Dual oxidase (duox)-derived reactive oxygen species (ROS) have been correlated with neuronal polarity, cerebellar development, and neuroplasticity. However, there have not been many comprehensive studies of the effect of individual duox isoforms on central-axon regeneration in vivo. Here, we explored this question in zebrafish, an excellent model organism for central-axon regeneration studies. In our research, mutation of the duox gene with CRISPR/Cas9 significantly retarded the single-axon regeneration of the zebrafish Mauthner cell in vivo. Using deep transcriptome sequencing, we found that the expression levels of related functional enzymes in mitochondria were down-regulated in duox mutant fish. In vivo imaging showed that duox mutants had significantly disrupted mitochondrial transport and redox state in single Mauthner-cell axon. Our research data provide insights into how duox is involved in central-axon regeneration by changing mitochondrial transport.
Reactive oxygen species (ROS) are chemically reactive molecules or free radicals containing oxygen, such as superoxide anion radicals (O2−) and hydrogen peroxide (H2O2) . ROS were known to be inevitable and damage-inducing by-products of cellular respiration long before the discovery of nicotinamide adenine dinucleotide phosphate (NADPH) oxidase family members [2, 3]. The NADPH oxidase (NOX) family is involved in the production of ROS in response to various stages of cellular differentiation, growth, and maintenance [4,5,6]. Dual oxidase (Duox), a member of the NOX family, was originally identified as thyroid NADPH oxidase . However recent studies have reported that Duox enzymes are also expressed in the salivary glands, rectum, trachea, and bronchium [8, 9]. In zebrafish, there is only a single duox gene, in contrast to the two duox genes in both humans and rodents, Duox1 and Duox2 . Recently, it has been shown that Duox-generated H2O2 is critical for the recruitment of leukocytes to initiate inflammation in zebrafish larvae [11, 12]. Studies in zebrafish have also shown that Duox deficiency presents as congenital hypothyroidism [7, 13,14,, 14].
In addition, Duox has been reported to be vital in heart regeneration  and sensory-axon regeneration . However, little is known about its role in the regeneration of axons in the central nervous system (CNS). In mammals, axon regeneration in the CNS is extremely limited [17, 18], unlike in the peripheral nervous system (PNS) . Conversely, several types of neurons have robust regenerative abilities in the CNS of zebrafish [20,21,22,23,24]. Recent studies in zebrafish have used laser axotomy to precisely damage single axons in the CNS, making it possible to explore the potential factors impacting axon regeneration [25, 26]. For example, miRNAs , mitochondria , and Ca2+ activity  have been demonstrated to be involved in Mauthner-cell axon regeneration after two-photon laser ablation. Hence, zebrafish have become an attractive vertebrate model for studying the impact of Duox on axon regeneration in the CNS.
In our study, we found that loss of Duox retarded Mauthner cell axon regeneration in zebrafish in vivo, and that this was due to disrupted mitochondrial transport and redox state.
Zebrafish Lines and Maintenance
Wild-type (WT), duox+/- mutant, duox-/- mutant, Tg (Tol-056) , Tg (Tol-056); duox+/- mutant, and Tg (Tol-056); duox-/- mutant zebrafish (Danio rerio) were used in this study. Zebrafish embryos were bred with a laboratory stock and maintained at 28.5°C with a 14/10 h light/dark cycle. Embryos were collected from natural spawning and staged by dpf (days post-fertilization) according to established criteria . To prevent dark pigment formation, larvae were raised in embryo medium containing 0.2 mm N-phenylthiourea (Sigma). All animal manipulations in this study were conducted in strict accordance with the guidelines and regulations set forth by the University of Science and Technology of China (USTC) Animal Resources Center and the University Animal Care and Use Committee. The protocol was approved by the Committee on the Ethics of Animal Experiments of the USTC (Permit Number: USTCACUC1103013).
Design of Mutant Sites and Synthesis of Cas9 mRNA and sgRNA
The sgRNAs were designed to target the fifth exon of the duox gene by “SeqBuilder” software (DNAStar, USA). The targeting sequences started with GG, ended with NGG (PAM), and contained the restrictive enzyme BamH-I site near the PAM for genotyping. The Cas9 mRNA and sgRNAs were synthesized with little modification as previously described . In brief, the Cas9 mRNA was synthesized with a T7 mMESSAGE mMACHINE Kit (Promega, USA). The DNA fragments of the sgRNAs were amplified with pairs of primers (Table S1), and then purified with phenol and chloroform. The sgRNAs were transcribed in vitro with T7 Riboprobe Systems (Promega, USA).
Construction and Identification of duox Mutants
Cas9 mRNA (250 ng/μL) and sgRNAs (45 ng/μL) were co-microinjected into one-cell zebrafish embryos. Genomic DNAs, which were extracted from the injected embryos at 20 h post-fertilization, were used as templates for the following identifications. DNA fragments containing the duox target sequences were amplified by PCR and digested with BamH-I restriction endonucleases (Takara, Japan) at 37°C for 0.5 h. The uncleaved bands were cleaned after gel electrophoresis and cloned into pMD-19T (Takara, Japan). Monoclonal colonies were picked up for PCR and restriction enzyme digestion and were then sequenced by Sanger sequencing (Genewiz, Inc.). Primers used in the experiment are listed in Supplementary Table S1.
The microinjected founder (F0) larvae were raised to adulthood and then crossed with WT zebrafish to generate F1 larvae. The F1 larvae that carried heterozygous knockout alleles were raised to adulthood, and their genotypes were confirmed by PCR amplification and sequencing analysis of DNA from fin clippings. Homozygous knockout zebrafish were generated and selected by genotyping and then crossing male and female zebrafish carrying heterozygous knockout alleles.
Retro-complementation and Single-Cell Electroporation In Vivo
For duox gene retro-complementation, duox gene and duox-/- mutant gene CDS sequences were cloned via primer (Table S1), then inserted into the plasmid UAS-mCherry to construct the retro-complementation plasmids UAS-duox-mCherry and UAS-duox-/--mCherry. Then the CMV-GAL4-VP16/UAS-mCherry plasmids (control group), CMV-GAL4-VP16/UAS-duox-mCherry plasmids (duox gene retro-complementation group), and CMV-GAL4-VP16/UAS-duox-/--mCherry plasmids (duox-/- gene retro-complementation group) were co-transfected through single-cell electroporation into unilateral Mauthner cells at 4 dpf in duox-/- mutant fish as described previously . Different concentrations of rhodamine-dextran (a fluorescent dye that labels Mauthner cells), CMV-GAL4-VP16 (a plasmid that drives the expression of GAL4-VP16), UAS-mCherry/UAS-duox-mCherry/ UAS-duox-/--mCherry (plasmids that drive the expression of the red fluorescent protein, mCherry), UAS-mito-EGFP (a plasmid that labels mitochondria), and UAS-mito-roGFP2-Orp-1 (plasmids that the drive expression of redox state probes; Addgene, no. 64997) were combined (using 100–200 ng/µL of each plasmid) and co-delivered into unilateral Mauthner cells. Zebrafish larvae were returned to embryo media containing N-phenylthiourea and allowed to recover. Two days after electroporation, morphologically normal and healthy larvae were selected for subsequent experiments.
Before axotomy, each 6-dpf zebrafish larva expressing mCherry/GFP fluorescence in unilaterally in Mauthner cells was anesthetized in MS222 (Sigma) and immobilized in 1% low-melting agarose. A Zeiss two-photon microscope (LSM710) was used to ablate Mauthner cell axons over the cloacal pores [26, 34]. We used a two-photon laser at 800 nm and an intensity of 16%–22% to ablate the Mauthner cell axons.
In Vivo Imaging of Axon Regeneration and Mitochondrial Movement
To observe Mauthner cell axon regeneration, anesthetized larvae were imaged at 2 days-post axotomy (dpa) with a confocal system (Olympus FV1000) and a water-immersion lens (40×, 0.85 numerical-aperture objective) at 2-μm intervals. All of the images were spliced using Adobe Photoshop CS4. The point just above the cloacal pores was defined as the starting point of regrowth, and the axonal terminal of regeneration was defined as the end-point of regrown axons. All of the fluorescent live images and time-lapse movies show a lateral view of the spinal cord, with the anterior to the left and dorsal above. The regenerative length was calculated using Fiji-ImageJ.
To obtain static images of mitochondrial morphology in the axon terminal, larvae were screened for co-labeled mito-EGFP and rhodamine-dextran in Mauthner cells. We collected images of larval axonal terminals at 6, 7, and 8 dpf using an Olympus microscope equipped with a water-immersion lens (60×, 0.9 numerical-aperture objective) at 1-µm intervals. Dynamic imaging and analysis of mitochondrial movement in axons in vivo were as described previously . Mitochondrial motility was defined as the percentage of moving mitochondria. The speed of a mitochondrion was defined as the total distance moved divided by the time spent moving.
To image the redox state of mitochondria in Mauthner cells, Mauthner cell soma was scanned with a confocal system (Zeiss, LSM710). Biosensor fluorescence was excited using 405-nm and 488-nm lasers sequentially and via line-by-line scanning. Emission was detected at 500–570 nm. The fluorescent intensity of an Mauthner cell soma in a single horizontal plane was measured. The fluorescent ratio (405 nm/488 nm) of each Mauthner cell soma was calculated using Fiji-ImageJ.
RNA Extraction and qRT-PCR
Total RNAs were extracted from 50 larvae of the WT and duox-/- lines using TRIzol (Takara, Japan) reagent. Quantitative real-time PCR (qRT-PCR) was performed with the SYBR green kit (Invitrogen, USA). qRT-PCR was performed in triplicate with three individual biological samples (nine replicates). The results were normalized to the expression level of the housekeeping gene β-actin and are shown as a relative expression level calculated using the 2-ΔΔCt method .
To examine the protein expression in duox-/- mutant zebrafish, 5-dpf WT and duox-/- mutant larvae were collected and lysed with RIPA buffer. Samples were boiled for 5 min and run on a 12% SDS-PAGE gel. Mouse monoclonal anti-Duox1 (Santa Cruz, USA) was used to assess the protein levels in each group.
Deep-Sequencing-Based Transcriptomic Analysis
Total RNAs were extracted from 4-dpf duox-/- and WT zebrafish, and 3 g of RNA per sample was prepared for constructing a transcriptional library. Sequencing libraries were generated with an UltraTM RNA Library Prep Kit (NEB, USA) according to the manufacturer’s instructions. Clustering of the index-coded samples was conducted with a cBot Cluster Generation System using a TruSeq PE Cluster Kit (Illumina, USA), per the instructions. After clustering, the library preparations were sequenced on an Illumina Hiseq 2000 platform. Perl scripts were used to remove the adapter for clean reads, calculating the Q20/Q30 duplicate data, and generating the raw reads. Transcriptomic assembly was performed according to a protocol described previously . For the Gene Ontology (GO) enrichment assay, the differentially-expressed genes (DEGs) were identified using Wallenius’ non-central hypergeometric distribution, implemented in the GOseq R packages . Kobas software was used to determine the statistical enrichment of DEGs in KEGG pathways to predict and classify the functions of the assembled sequences .
Transmission Electron Microscopy
Each larval zebrafish fixed in 2.0% formaldehyde and 2.5% glutaraldehyde solution (Electron Microscopy Sciences) overnight at 4 °C. Following washes, they were washed with 0.1 mol/L phosphate buffer (pH 7.4). Specimens were then incubated in post-fixation solution containing 1% osmium tetroxide for 2 h, washed with water, washed three times with 0.1 mol/L phosphate buffer (pH 7.4) for 15 min each time. Next, specimens were washed with water, dehydrated with serial dilutions of ethanol in water (50%, 70%, 80%, 90%, 100%, 100%) and 100% acetone twice for 15 min each. The samples were then embedded in Epon/Araldite resin with surrounding support tissue and hardened for 2–3 days at 60°C. Ultrathin (80 nm) transverse sections of the brain from larvae were stained with uranyl acetate and lead citrate. Sections were viewed and photographed with an FEI Tecnai Spirit (120 kV) transmission electron microscope (TEM).
All of the data are reported as the mean ± SEM, or as relative proportions of 100%, as indicated in the figure legends. We used either Student’s two-tailed t tests or one-way analyses of variance (ANOVAs) for all of analyses, as indicated in the figure legends. All graphs were constructed and statistical tests were performed in GraphPad Prism 7. We considered P < 0.05 statistically significant (*P <0.05; **P <0.01; ***P <0.001). We defined the length from the cloacal pore to the tail end as the whole length of the Mauthner cell, as the head end was too thick to sufficiently image in vivo.
Generation and Identification of duox Mutant Zebrafish
To investigate the functions of duox in vivo, we attempted to obtain a duox mutant zebrafish line. We designed a CRISPR-Cas9-targeted site in the fifth exon of the zebrafish duox gene that contained a BamH-I restriction site for further identifying the resultant mutants (Fig. 1A). Cas9 mRNA and duox sgRNA were co-microinjected into one-cell embryos. To identify specific mutagenesis, a 440-bp DNA fragment was amplified by PCR from the genomic DNA and digested with a BamH-I restriction enzyme. The results of gel electrophoresis showed that F2 zebrafish larvae were heritably homozygous mutants (Fig. 1B). Moreover, the representative sequencing results indicated that F2 zebrafish had a two-bp deletion (Fig. 1C–E). Furthermore, bioinformatics analysis revealed that the functional sequence domains (HLH-PAS-PAS) of the F2 zebrafish Duox protein were frame-shifted (Fig. 1F). Finally, we used Western blotting to determine whether our duox mutation was a null allele mutation. The results showed that the expression of Duox protein was completely abolished in duox mutant zebrafish (Fig. 1G and H). Taken together, these results verified that we successfully generated a duox null-mutant zebrafish line.
Duox Deficiency Retards Mauthner-Cell Axon Regeneration at an Early Stage In Vivo
Duox is known to be required for promoting injury-induced peripheral sensory axon regeneration in zebrafish skin . However, the in vivo role of Duox in CNS axon regeneration has remained unclear. Mauthner cells are a pair of myelinated hindbrain neurons projecting to the spina cord in zebrafish and have been demonstrated to exhibit a strong regenerative capacity in our previous studies [26,27,28]. To investigate the roles of Duox in Mauthner cell axon regeneration, we crossed the transgenic line Tg (T056: EGFP) with the duox-/- mutant for two consecutive generations to obtain the Tg (T056: EGFP)/duox-/- zebrafish line (Fig. 2A). We used two-photon laser axotomy to transect one of the Mauthner cell unilateral axons over the cloacal pores at 6 dpf (Fig. 2B). Furthermore, in vivo live imaging showed that homozygous and heterozygous duox mutant larvae displayed a reduced length of Mauthner cell axon regeneration compared with that of WT larvae (Fig. 2C and D). We analyzed the whole length of Mauthner cell axons and found no significant difference among WT, homozygous and heterozygous duox mutant larvae, indicating that the original length of Mauthner-cell axon was unaffected by duox mutant (Fig. 2E and F). For the retro-complementation experiment, we co-transfected CMV-GAL4-VP16/UAS-mCherry plasmids (duox-/- mutant + UAS-mCherry group), CMV-GAL4-VP16/UAS-duox-mCherry plasmids (duox-/- mutant + UAS-duox-mCherry group), and CMV-GAL4-VP16/UAS-duox-/--mCherry plasmids (duox-/- mutant + UAS-duox-/--mCherry group) through single-cell electroporation into unilateral Mauthner cells at 4 dpf in duox-/- mutant fish (Fig. 2G and H). Subsequently, we ablated red-fluorescent Mauthner cell axons at 6 dpf with a two-photon laser-scanning microscope and continued to image the regenerated length of Mauthner cells at 8 dpf (2 dpa) (Fig. 2I). Live imaging results showed that overexpression of the duox gene remarkably rescued the weak regenerative ability in duox-/- mutant fish and duox-/- gene retro-complementation had no significance impact on Mauthner cell axon regeneration (Fig. 2J and K). Taken together, these results suggest that Duox is required for Mauthner cell axon regeneration in zebrafish.
Duox Modulates Genes Associated with Mitochondrial Function
Robust Mauthner cell axon regeneration requires high mobility of mitochondrial transport along axons . ROS is known to play important roles in both mitochondrial H2O2 generation and the electron transport chain . To investigate the Duox-affected genes involved in mitochondria, we performed high-throughput RNA sequencing in duox-/- mutant and WT larvae at 4 dpf. Transcriptomic analysis revealed that there were 360 down-regulated genes and 375 up-regulated genes in duox-/- mutant fish (Fig. 3A and Table S2) (The data have been deposited in the NCBI Gene Expression Omnibus, GEO accession number GSE144689). Furthermore, GO analyses revealed that these DEGs participated in rhythmic processes, responded to stimuli, and were involved in growth, antioxidant activity, catalytic activity, and transcription factor activity, and were significantly altered in duox-/- mutants (Fig. S1). Importantly, genes related to mitochondria, such as cyp2p9, alas2, cyp11a1, mfn2, opa1, atp5f1b, cyp2x8, gpx4b, and gsr, were also down-regulated (Fig. 3B), which was further corroborated via qRT-PCR (Fig. 3C). These results suggest that Duox plays an important role in mitochondria.
Duox Affects Mitochondrial Morphology and Dynamics
Considering our previous study showing that mitochondria play a crucial role in the process of Mauthner cell axon regeneration , we next investigated whether duox-/- mutations also affect mitochondrial transport in zebrafish larvae. We hypothesized that Duox mutations impair Mauthner cell axon regeneration length via disrupting mitochondrial transport.
To test the above hypothesis, we used TEM to observe mitochondrial morphology in the zebrafish brain (i.e., the mitochondria in the Mauthner cell axon were difficult to localize via TEM). TEM imaging results showed that mitochondrial sizes in duox-/- mutants had dramatically larger surface areas, perimeters, and Feret’s diameters than those in the WT group (Fig. S2). Duox mutations disrupting mitochondrial fusion and fission processes due to harmful substance invasion or other unknown mechanisms may have participated in shaping the differences in mitochondrial sizes and morphology in duox-/- mutants compared to those in WTs.
To further investigate the role of Duox in mitochondrial dynamics, we imaged mitochondrial movement in Mauthner cell axons (Fig 4A). Mitochondrial movement directly affects mitochondrial transport which has deleterious effects on the delivery of energy to the regeneration site [40, 41]. Mitochondrial motility was defined as the percentage of moving mitochondria. The speed of a moving mitochondrion was defined as the total distance that a mitochondrion moved divided by the time spent moving. Mitochondrial speed was significantly slower in the Mauthner cell axons of duox mutants (in 12 fish; total: 0.3481 ± 0.01513 μm/s, n = 82 mitochondria; anterograde: 0.3307 ± 0.01468 μm/s, n = 73 mitochondria; retrograde: 0.4896 ± 0.05067 μm/s, n = 9 mitochondria) than that of the WT group (in 12 fish; total: 0.5743 ± 0.02923 μm/s, n = 54 mitochondria; anterograde: 0.5475 ± 0.02838 μm/s, n = 49 mitochondria; retrograde: 0.8364 ± 0.0927 μm/s, n = 5 mitochondria; Fig. 4B and D). However, mitochondrial motility in Mauthner cell axons of duox mutant larvae (in 12 fish; total: 30.72% ± 3.744%; anterograde: 24.01% ± 3.999%; retrograde: 6.879% ± 3.273%) was not significantly different from that in WT larvae (in 12 fish; total: 33.11% ± 4.593%; anterograde: 29.32% ± 4.229%; retrograde: 3.781% ± 1.704%; Fig. 4B and C). Taken together, although the ratio of mobile mitochondria was unaffected in duox-/- mutants, duox gene deficiency affected the speed of mitochondria along axons, which may have disrupted the efficacy of mitochondrial transport for providing energy.
To further investigate the roles of Duox in mitochondrial function, we assessed the mitochondrial redox state in single unilateral Mauthner cell axons via redox-sensitive GFPs (roGFPs), which allow subcellular redox-coupled-specific in vivo imaging in model organisms [42,43,44]. Therefore, we recognized that roGFP2 could be converted into a specific probes for H2O2 via coupling it to the microbial H2O2 sensor, oxidant receptor peroxidase 1 (Orp1) .
Since H2O2 is the major oxidant species involved in protein thiol oxidation and redox regulation, we generated a UAS-mito-roGFP2-Orp1 plasmid for measuring H2O2 in the mitochondrial matrix. H2O2 levels in Mauthner cell mitochondria were lower in the duox mutant group (0.9358 ± 0.04672, n = 8) than in the WT group (2.125 ± 0.2215, n = 4; Fig. 4E and F). In addition, H2O2 levels and ROS levels in whole-mount larvae were also down-regulated due to duox gene deficiency (Fig. S3). Collectively, these results suggest that duox gene mutation induces a low level of H2O2 formation, thus implying that mitochondrial redox in Mauthner cell axons is unbalanced.
In summary, we observed that duox mutants have disrupted mitochondrial morphology and function, especially in terms of a slower speed of transport in Mauthner cells, which may contribute to the retarded Mauthner cell axon regeneration in duox mutants.
Duox Affects Mitochondrial Mobility and Speed During an Early Stage of Mauthner-Cell Axon Regeneration
To directly examine the role of Duox in mitochondrial transport in Mauthner cell axon regeneration, we measured mitochondrial mobility and speed at 24 h post-ablation (hpa) and 48 hpa. The results showed that in 24 hpa larvae, duox gene deficiency resulted in a slower mitochondrial speed (in 26 fish; total: 0.5313 ± 0.01123 μm/s, n = 208 mitochondria; anterograde: 0.524 ± 0.01117 μm/s, n = 186 mitochondria; retrograde: 0.5904 ± 0.04542 μm/s, n = 23 mitochondria) and lower mobility (in 26 fish; total: 25.22% ± 2.306%; anterograde: 22.09% ± 2.493%; retrograde: 3.121% ± 1.041%) than in the WT group (mitochondrial speed in 10 fish; total: 0.5814 ± 0.01365 μm/s, n = 56 mitochondria; anterograde: 0.576 ± 0.014 μm/s, n = 53 mitochondria; retrograde: 0.6764 ± 0.02918 μm/s, n = 3 mitochondria; Fig. 5A and B; mitochondrial mobility in 10 fish; total: 40.69% ± 3.422%; anterograde: 40.27% ± 3.307%; retrograde: 0.417% ± 0.417%; Fig. 5A and C). However, at 48 hpa, there was no significant difference in mitochondrial speed or mobility between the duox-/- mutant and WT groups (Fig. 5B and C). Taken together, these results suggest that Duox mutants affect mitochondrial speed and motility in the early stages of regeneration, resulting in a lower energy supply for Mauthner cell axon regeneration.
Duox, as a member of the NADPH oxidase family, was originally identified as thyroid NADPH oxidase, and has been demonstrated to be involved in the production of ROS in response to different extracellular signals. Studies of innate immunity have established Duox-generated H2O2 candidates for wound-to-leukocyte signaling in zebrafish . Duox has been reported to play important roles in heart regeneration and sensory axon regeneration [15, 16], both of which are parts of the PNS. However, our previous work has shown that Mauthner cells have robust axon regeneration after two-photon axotomy, indicating that Mauthner cells are a promising model for studying CNS axon regeneration at single-cell resolution in vivo; in addition, Mauthner cells are large cells that can be viewed clearly and easily in vivo [26, 27]. Here, we found that Mauthner cell axon regeneration was inhibited in duox-/- mutant zebrafish. This result was not likely due to a mere developmental delay because the whole length of the Mauthner cell axon in mutant larvae was not significantly different before ablation compared to that in WTs. Our results are consistent with the above findings of PNS regeneration in that we also found that Duox plays a vital role in CNS regeneration due to H2O2 production.
In developing neurons, mitochondria are involved in many critical functions, including respiration/ATP production, Ca2+ buffering, apoptotic regulation, heme and Fe/S cluster biosynthesis, and ROS signaling/quenching [46, 47]. During the process of oxidative phosphorylation, mitochondria utilize oxygen to generate ATP from organic fuel molecules but in the process also produce ROS. H2O2 is a major type of ROS in organisms and is a central hub in redox signaling; H2O2 is mainly generated from NADPH oxidases or Complex III of the mitochondrial respiratory chain . As mitochondria are the main source of cellular energy, we speculated that Duox affecting Mauthner cell axon regeneration was correlated with mitochondrial dynamics and H2O2 levels. To test this hypothesis, we investigated the effects of Duox mutations on mitochondrial motility and speed in Mauthner cell axons. We found that mitochondrial speed was slower in duox-/- mutant larvae but that the mobility in Mauthner cell axons was unaffected. In addition, we used enzyme-coupled roGFP2-based probes to visualize chemically defined redox species in vivo. Orp-1-based probes specifically and reversibly report roGFP2 oxidation by H2O2 . Our results showed that H2O2 levels were lower in duox-/- mutant larvae than in WTs. These results indicate that Duox induced a decrease in the amount of energy transported to axons and resulted in poor axon regeneration. Our results are consistent with findings from a previous study in giant pandas that used a mutation in the duox gene to allow them to have a lower metabolic rate through a lower level of thyroid hormone synthesis . This study revealed that Duox deficiency inhibited Mauthner cell axon regeneration due to a lower metabolic rate through a lower level of mitochondrial energy production. However, these findings represented the Mauthner cell axonal state in duox mutants without laser ablation.
To further investigate the effect of Duox in Mauthner cell axon regeneration after laser ablation, we measured mitochondrial speed and mobility at 24 hpa and 48 hpa. We found that mitochondrial speed and mobility were both lower in duox-/- mutant larvae than in WT larvae at 24 hpa, but there was no significant difference at 48 hpa; hence, we further focused on the changes at 24 hpa. Compared to non-ablation and 48 hpa results, we found that mitochondrial mobility played more vital roles at 24 hpa; we propose two possible accounts for this finding. First, mitochondria are mainly trafficked along microtubules by ATPase-dependent kinesin and dynein motor proteins and dendrite- and axon-specific adaptor proteins. In distal dendrites and the entire axon, the plus-end of microtubules is oriented toward the growth cone, and kinesins mediate anterograde transport while dyneins mediate retrograde transport . In proximal dendrites, microtubule polarity is mixed, and motor proteins are not selective for either anterograde or retrograde transport . After ablation, the growth cone was closer to our imaging position at the distal axonal area (Fig. 1A in our previous study)  compared to that during non-ablation conditions, so we speculated that Duox affects kinesin. Second, H2O2 is rapidly produced from axonal laser ablation; to avoid damage from high levels of H2O2, Mauthner cell axons increase mitochondrial mobility to consume more H2O2  in WT larvae. However, this mechanism is unnecessary in duox-/- mutant larvae since H2O2 generation is inhibited. These findings collectively suggest that during axon regeneration, axons require highly integrated metabolic machinery and more anterograde trafficking of mitochondria for local ATP synthesis to meet the large energy demands of regeneration, but these processes appear to be impeded in duox-/- mutant larvae.
In summary, we found that zebrafish Duox inhibited Mauthner cell axon regeneration via affecting mitochondrial mobility and speed. Our present study indicates that metabolic insufficiency induced by duox mutant may impair Mauthner cell regeneration due to impeded mitochondrial dynamics, which provides insight into the interactions among energy metabolism, mitochondrial trafficking, and axon regeneration in the CNS.
The datasets generated during and/or analyzed during the current study are available from the corresponding author on reasonable request.
Bolduc JA, Collins JA, Loeser RF. Reactive oxygen species, aging and articular cartilage homeostasis. Free Radic Biol Med 2019, 132: 73–82.
Buvelot H, Jaquet V, Krause KH. Mammalian NADPH Oxidases. Methods Mol Biol 2019, 1982: 17–36.
Xu FF, Zhang ZB, Wang YY, Wang TH. Brain-derived glia maturation factor beta participates in lung injury induced by acute cerebral ischemia by increasing ROS in endothelial cells. Neurosci Bull 2018, 34: 1077–1090.
Veith C, Boots AW, Idris M, van Schooten FJ, van der Vliet A. Redox imbalance in idiopathic pulmonary fibrosis: A role for oxidant cross-talk between NADPH oxidase enzymes and mitochondria. Antioxid Redox Signal 2019, 31: 1092–1115.
Marrali G, Casale F, Salamone P, Fuda G, Ilardi A, Manera U, et al. NADPH oxidases 2 activation in patients with Parkinson’s disease. Parkinsonism Relat Disord 2018, 49: 110–111.
Ma MW, Wang J, Dhandapani KM, Wang R, Brann DW. NADPH oxidases in traumatic brain injury—Promising therapeutic targets? Redox Biol 2018, 16: 285–293.
Park JS, Choi TI, Kim OH, Hong TI, Kim WK, Lee WJ, et al. Targeted knockout of duox causes defects in zebrafish growth, thyroid development, and social interaction. J Genet Genomics 2019, 46: 101–104.
Geiszt M, Witta J, Baffi J, Lekstrom K, Leto TL. Dual oxidases represent novel hydrogen peroxide sources supporting mucosal surface host defense. FASEB J 2003, 17: 1502–1504.
Sirokmany G, Donko A, Geiszt M. Nox/Duox family of NADPH oxidases: Lessons from knockout mouse models. Trends Pharmacol Sci 2016, 37: 318–327.
Flores MV, Crawford KC, Pullin LM, Hall CJ, Crosier KE, Crosier PS. Dual oxidase in the intestinal epithelium of zebrafish larvae has anti-bacterial properties. Biochem Biophys Res Commun 2010, 400: 164–168.
Niethammer P, Grabher C, Look AT, Mitchison TJ. A tissue-scale gradient of hydrogen peroxide mediates rapid wound detection in zebrafish. Nature 2009, 459: 996–999.
Yoo SK, Starnes TW, Deng Q, Huttenlocher A. Lyn is a redox sensor that mediates leukocyte wound attraction in vivo. Nature 2011, 480: 109–112.
Moreno JC, Bikker H, Kempers MJE, van Trotsenburg P, Baas F, de Vijlder JJM, et al. Inactivating mutations in the gene for thyroid oxidase 2 (THOX2) and congenital hypothyroidism. N EngI J Med 2002, 347: 95–102.
Park KJ, Park HK, Kim YJ, Lee KR, Park JH, Park JH, et al. DUOX2 mutations are frequently associated with congenital hypothyroidism in the korean population. Ann Lab Med 2016, 36: 145–153.
Han P, Zhou XH, Chang N, Xiao CL, Yan S, Ren H, et al. Hydrogen peroxide primes heart regeneration with a derepression mechanism. Cell Res 2014, 24: 1091–1107.
Rieger S, Sagasti A. Hydrogen peroxide promotes injury-induced peripheral sensory axon regeneration in the zebrafish skin. PLoS Biol 2011, 9: e1000621.
Chen MF, Zheng BH. Axon plasticity in the mammalian central nervous system after injury. Trends Neurosci 2014, 37: 583–593.
Becker T, Becker CG. Axonal regeneration in zebrafish. Curr Opin Neurobiol 2014, 27: 186–191.
Li WY, Zhang WT, Cheng YX, Liu YC, Zhai FG, Sun P, et al. Inhibition of KLF7-targeting microRNA 146b promotes sciatic nerve regeneration. Neurosci Bull 2018, 34: 419–437.
Taylor JSH, Jack JL, Easter SS. Is the Capacity for optic-nerve regeneration related to continued retinal ganglion-cell production in the frog—a test of the hypothesis that neurogenesis and axon regeneration are obligatorily linked. Eur J Neurosci 1989, 1: 626–638.
Nagashima M, Fujikawa C, Mawatari K, Mori Y, Kato S. HSP70, the earliest-induced gene in the zebrafish retina during optic nerve regeneration: Its role in cell survival. Neurochem Int 2011, 58: 888–895.
Saijilafu, Zhang BY, Zhou FQ. Signaling pathways that regulate axon regeneration. Neurosci Bull 2013, 29: 411–420.
Vajn K, Plunkett JA, Tapanes-Castillo A, Oudega M. Axonal regeneration after spinal cord injury in zebrafish and mammals: differences, similarities, translation. Neurosci Bull 2013, 29: 402–410.
Mu Z, Zhang S, He C, Hou H, Liu D, Hu N, et al. Expression of SoxC transcription factors during zebrafish retinal and optic nerve regeneration. Neurosci Bull 2017, 33: 53–61.
Yanik MF, Cinar H, Cinar HN, Chisholm AD, Jin YS, Ben-Yakar A. Neurosurgery—Functional regeneration after laser axotomy. Nature 2004, 432: 822.
Hu BB, Chen M, Huang RC, Huang YB, Xu Y, Yin W, et al. In vivo imaging of Mauthner axon regeneration, remyelination and synapses re-establishment after laser axotomy in zebrafish larvae. Exp Neurol 2018, 300: 67–73.
Huang R, Chen M, Yang L, Wagle M, Guo S, Hu B. MicroRNA-133b Negatively regulates zebrafish single mauthner-cell axon regeneration through targeting tppp3 in vivo. Front Mol Neurosci 2017, 10: 375.
Xu Y, Chen M, Hu B, Huang R, Hu B. In vivo imaging of mitochondrial transport in single-axon regeneration of zebrafish mauthner cells. Front Cell Neurosci 2017, 11: 4.
Chen M, Huang RC, Yang LQ, Ren DL, Hu B. In vivo imaging of evoked calcium responses indicates the intrinsic axonal regenerative capacity of zebrafish. FASEB J 2019, 33: 7721–7733.
Satou C, Kimura Y, Kohashi T, Horikawa K, Takeda H, Oda Y, et al. Functional role of a specialized class of spinal commissural inhibitory neurons during fast escapes in zebrafish. J Neurosci 2009, 29: 6780–6793.
Kimmel CB, Ballard WW, Kimmel SR, Ullmann B, Schilling TF. Stages of embryonic-development of the zebrafish. Dev Dyn 1995, 203: 253–310.
Jao LE, Wente SR, Chen WB. Efficient multiplex biallelic zebrafish genome editing using a CRISPR nuclease system. Proc Natl Acad Sci U S A 2013, 110: 13904–13909.
Chen M, Xu Y, Huang RC, Huang YB, Ge SC, Hu B. N-cadherin is involved in neuronal activity-dependent regulation of myelinating capacity of zebrafish individual oligodendrocytes in vivo. Mol Neurobiol 2017, 54: 6917–6930.
O’Donnell KC, Vargas ME, Sagasti A. WldS and PGC-1 alpha regulate mitochondrial transport and oxidation state after axonal injury. J Neurosci 2013, 33: 14778–14790.
VanGuilder HD, Vrana KE, Freeman WM. Twenty-five years of quantitative PCR for gene expression analysis. Biotechniques 2008, 44: 619–626.
Grabherr MG, Haas BJ, Yassour M, Levin JZ, Thompson DA, Amit I, et al. Full-length transcriptome assembly from RNA-Seq data without a reference genome. Nat Biotechnol 2011, 29: 644–652.
Young MD, Wakefield MJ, Smyth GK, Oshlack A. Gene ontology analysis for RNA-seq: accounting for selection bias. Genome Biol 2010, 11: R14.
Kanehisa M, Araki M, Goto S, Hattori M, Hirakawa M, Itoh M, et al. KEGG for linking genomes to life and the environment. Nucleic Acids Res 2008, 36: D480–D484.
Dan Dunn J, Alvarez LA, Zhang X, Soldati T. Reactive oxygen species and mitochondria: A nexus of cellular homeostasis. Redox Biol 2015, 6: 472–485.
Cartoni R, Norsworthy MW, Bei F, Wang C, Li S, Zhang Y, et al. The mammalian-specific protein armcx1 regulates mitochondrial transport during axon regeneration. Neuron 2016, 92: 1294–1307.
Zhou B, Yu P, Lin MY, Sun T, Chen Y, Sheng ZH. Facilitation of axon regeneration by enhancing mitochondrial transport and rescuing energy deficits. J Cell Biol 2016, 214: 103–119.
Hanson GT, Aggeler R, Oglesbee D, Cannon M, Capaldi RA, Tsien RY, et al. Investigating mitochondrial redox potential with redox-sensitive green fluorescent protein indicators. J Biol Chem 2004, 279: 13044–13053.
Albrecht SC, Barata AG, Grosshans J, Teleman AA, Dick TP. In vivo mapping of hydrogen peroxide and oxidized glutathione reveals chemical and regional specificity of redox homeostasis. Cell Metab 2011, 14: 819–829.
Dooley CT, Dore TM, Hanson GT, Jackson WC, Remington SJ, Tsien RY. Imaging dynamic redox changes in mammalian cells with green fluorescent protein indicators. J Biol Chem 2004, 279: 22284–22293.
Gutsche M, Sobotta MC, Wabnitz GH, Ballikaya S, Meyer AJ, Samstag Y, et al. Proximity-based protein thiol oxidation by H2O2-scavenging peroxidases. J Biol Chem 2009, 284: 31532–31540.
Mattson MP, Gleichmann M, Cheng A. Mitochondria in neuroplasticity and neurological disorders. Neuron 2008, 60: 748–766.
McWilliams TG, Prescott AR, Allen GFG, Tamjar J, Munson MJ, Thomson C, et al. mito-QC illuminates mitophagy and mitochondrial architecture in vivo. J Cell Biol 2016, 214: 333–345.
Sies H. Role of metabolic H2O2 generation. J Biol Chem 2014, 289: 8735–8741.
Nie YG, Speakman JR, Wu Q, Zhang CL, Hu YB, Xia MH, et al. Exceptionally low daily energy expenditure in the bamboo-eating giant panda. Science 2015, 349: 171–174.
Baas PW, Deitch JS, Black MM, Banker GA. Polarity Orientation of microtubules in hippocampal-neurons—uniformity in the axon and nonuniformity in the dendrite. Proc Natl Acad Sci U S A 1988, 85: 8335–8339.
Treberg JR, Braun K, Selseleh P. Mitochondria can act as energy-sensing regulators of hydrogen peroxide availability. Redox Biol 2019, 20: 483–488.
This work was supported by the National Natural Science Foundation of China (31771183 and 31701027) and the National Key Research and Development Program of China (2019YFA0405603 and 2019YFA0405600). We thank LetPub (www.letpub.com) for its linguistic assistance during the preparation of this manuscript.
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The authors declare no competing financial interest.
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Yang, LQ., Chen, M., Ren, DL. et al. Dual Oxidase Mutant Retards Mauthner-Cell Axon Regeneration at an Early Stage via Modulating Mitochondrial Dynamics in Zebrafish. Neurosci. Bull. 36, 1500–1512 (2020). https://doi.org/10.1007/s12264-020-00600-9
- Mauthner cell
- Axon regeneration
- Mitochondrial dynamics