Fgf-dependent glial cell bridges facilitate spinal cord regeneration in Zebrafish

Adult Zebrafish show a remarkable capacity to regenerate their spinal column after injury, an ability that stands in stark contrast to the limited repair that occurs within the mammalian CNS post-injury. The reasons for this interspecies difference in regenerative capacity remain unclear. Here we demonstrate a novel role for Fgf signaling during glial cell morphogenesis in promoting axonal regeneration after spinal cordinjury. Zebrafish glia are induced by Fgf signaling, to form anelongated bipolarmorphology that formsabridge between the two sides of the resected spinal cord, over which regenerating axons actively migrate. Loss of Fgf function inhibits formation of this "glial bridge" and prevents axon regeneration. Despite the poor potential for mammalian axonal regeneration, primate astrocytes activated by Fgf signaling adopt a similar morphology to that induced in Zebrafish glia. This suggests that differential Fgf regulation, rather than intrinsic cell differences, underlie the distinct responses of mammalian and Zebrafish glia to injury

A Cytochrome P450 Conserved in Insects Is Involved in Cuticle Formation

The sequencing of numerous insect genomes has revealed dynamic changes in the number and identity of cytochrome P450 genes in different insects. In the evolutionary sense, the rapid birth and death of many P450 genes is observed, with only a small number of P450 genes showing orthology between insects with sequenced genomes. It is likely that these conserved P450s function in conserved pathways. In this study, we demonstrate the P450 gene, Cyp301a1, present in all insect genomes sequenced to date, affects the formation of the adult cuticle in Drosophila melanogaster. A Cyp301a1 piggyBac insertion mutant and RNAi of Cyp301a1 both show a similar cuticle malformation phenotype, which can be reduced by 20-hydroxyecdysone, suggesting that Cyp301a1 is an important gene involved in the formation of the adult cuticle and may be involved in ecdysone regulation in this tissue

Characterization of the laminin gene family and evolution in zebrafish

Laminins are essential components of all basement membranes and are fundamental to tissue development and homeostasis. Humans possess at least 16 different heterotrimeric laminin complexes formed through different combinations of alpha, beta, and gamma chains. Individual chains appear to exhibit unique expression patterns, leading to the notion that overlap between expression domains governs the constitution of complexes found within particular tissues. However, the spatial and temporal expression of laminin genes has not been comprehensively analyzed in any vertebrate model to date. Here, we describe the tissue-specific expression patterns of all laminin genes in the zebrafish, throughout embryonic development and into the "post-juvenile" period, which is representative of the adult body form. In addition, we present phylogenetic and microsynteny analyses, which demonstrate that the majority of our zebrafish sequences are orthologous to human laminin genes. Together, these data represent a fundamental resource for the study of vertebrate laminins

Testing of therapies in a novel nebulin nemaline myopathy model demonstrate a lack of efficacy

Abstract Nemaline myopathies are heterogeneous congenital muscle disorders causing skeletal muscle weakness and, in some cases, death soon after birth. Mutations in nebulin, encoding a large sarcomeric protein required for thin filament function, are responsible for approximately 50% of nemaline myopathy cases. Despite the severity of the disease there is no effective treatment for nemaline myopathy with limited research to develop potential therapies. Several supplements, including L-tyrosine, have been suggested to be beneficial and consequently self-administered by nemaline myopathy patients without any knowledge of their efficacy. We have characterized a zebrafish model for nemaline myopathy caused by a mutation in nebulin. These fish form electron-dense nemaline bodies and display reduced muscle function akin to the phenotypes observed in nemaline myopathy patients. We have utilized our zebrafish model to test and evaluate four treatments currently self-administered by nemaline myopathy patients to determine their ability to increase skeletal muscle function. Analysis of muscle pathology and locomotion following treatment with L-tyrosine, L-carnitine, taurine, or creatine revealed no significant improvement in skeletal muscle function emphasizing the urgency to develop effective therapies for nemaline myopathy

Characterization of muscle phenotypes in <i>actc1b</i>^<i>-/-</i> mutants and Actc1b morphants.

<p>A) Antibody labelling against Actinin2 and Phalloidin of <i>actc1b</i>^<i>-/-</i> mutants and wildtype siblings with Actinin2 (green) and F-actin (red) at 2 dpf and 6 dpf showing normal muscle morphology. Scale bar represents 50μm. B) Locomotion assays show a significant reduction in distance travelled by <i>actc1b</i>^<i>-/-</i> mutants compared to siblings (<i>actc1b</i>^<i>+/-</i> and <i>actc1b</i>^<i>+/+</i>) zebrafish. Error bars represent SEM for three independent experiment (n = 6,11,16 for <i>actc1b</i>^<i>+/+</i>; n = 24,23,18 for <i>actc1b</i>^<i>+/-</i>; and n = 13,9,10 for <i>actc1b</i>^<i>-/-</i> per experiment), *p<0.05 using a one-way ANOVA. C) Locomotion assays showing a significant reduction in distance travelled by Actc1b ex2 and UTR morphants compared to both Standard Control MO injected and uninjected zebrafish. No significant difference in locomotion is observed for Standard Control MO injected and uninjected zebrafish. Error bars represent median values and interquartile range (pooled samples from 3 independent experiments n = 45,48,46 for Actc1b ex2 MO; n = 45,48,33 for Actc1b UTR MO; n = 45,48,47 for Standard Control MO; and n = 48,47,45 for uninjected zebrafish), #p<0.0001 using a Kruskal-Wallis Test.</p

Quantification of RNA and protein levels following Actc1b morpholino knockdown.

<p>A) Representative data from western blot analysis for α-actin protein expression in <i>actc1b</i>^<i>-/-</i> and their wildtype siblings (<i>actc1b</i>^<i>+/-</i> and <i>actc1b</i>^<i>+/+</i>) at 2 dpf injected with either an Actc1b UTR, Actc1b ex2, or Standard Control MO. β-tubulin was used as a loading control. B) Quantification of western blot analysis from three independent replicate experiments of A) (<a href="http://www.plosgenetics.org/article/info:doi/10.1371/journal.pgen.1007212#pgen.1007212.s005" target="_blank">S5 Fig</a>) consisting of a pooled sample of 20 tails whereby α-actin protein levels were normalized against the -tubulin loading control. Error bars represent SEM for three independent experiments, *p<0.05, **p<0.01, and #p<0.0001 using a two-way ANOVA. C) Quantitative RT-PCR analysis for zebrafish <i>acta1a</i>, <i>acta1b</i>, <i>actc1a</i>, and <i>actc1b</i> genes in tail samples from <i>actc1b</i>^<i>+/+</i>, <i>actc1b</i>^<i>+/-</i>, and <i>actc1b</i>^<i>-/-</i> at 2 dpf. Error bars represent SEM for three independent experiments each consisting of a pooled sample of 30 tail samples, *p<0.05 indicates difference from <i>actc1b</i>^<i>+/+</i> using a one-way ANOVA. D) Quantitative RT-PCR analysis for zebrafish <i>acta1a</i>, <i>acta1b</i>, <i>actc1a</i>, and <i>actc1b</i> genes in whole embryos from Actc1b ex2 and Actc1b UTR morphants compared to Standard control MO injected zebrafish at 2 dpf. Error bars represent SEM for three independent experiments each consisting of a pooled sample of 20–30 embryos, **p<0.01, and #p<0.001 indicate difference from control MO using a one-way ANOVA.</p

Quantitative RT-PCR analysis for zebrafish α-actin genes.

<p>mRNA expression of <i>acta1a</i>, <i>acta1b</i>, <i>actc1a</i>, and <i>actc1b</i> genes in the A) head (comprising the heart tissue) and B) tail (comprising predominantly skeletal muscle tissue) during zebrafish development. Error bars for A) and B) represent SEM for three independent biological replicates each consisting of a pooled sample of 20–30 embryos.</p

Characterization of phenotypic severity following Actc1a morpholino knockdown.

<p>A) <i>actc1b</i>^<i>-/-</i> and wildtype siblings (<i>actc1b</i>^<i>+/-</i> and <i>actc1b</i>^<i>+/+</i>) injected with either an Actc1a splice or Standard Control MO were stained with Actinin2 and phenotypes were scored as either wildtype, mild (small outgrowth of aggregates at the myosepta (arrowheads)) or severe (large outgrowth of aggregates at the myosepta (arrowheads) and Actinin2 positive aggregates throughout the muscle fibers (arrows)). Scale bar represents 50μm. B) Quantification of the phenotypic severity for <i>actc1b</i>^<i>-/-</i> and wildtype siblings (<i>actc1b</i>^<i>+/+</i> and <i>actc1b</i>^<i>+/-</i>) injected with Actc1a splice compared to Standard Control MO injected zebrafish. Error bars represent SEM for three independent experiments (for Actc1a MO n = 8,13,7 <i>actc1b</i>^<i>+/+</i>, n = 27,16,16 <i>actc1b</i>^<i>+/-</i> and n = 7,15,7 <i>actc1b</i>^<i>+/+</i> and for Standard Control MO n = 7,3,7 <i>actc1b</i>^<i>+/+</i>, n = 16,21,20 <i>actc1b</i>^<i>+/-</i> and n = 4,8,6 <i>actc1b</i>^<i>-/-</i>), *p<0.05 indicates a significant difference in phenotype proportions using a Chi-square test.</p

Characterization of phenotypic severity following Actc1b morpholino knockdown.

<p>A) <i>actc1b</i>^<i>-/-</i> and wildtype siblings (<i>actc1b</i>^<i>+/-</i> and <i>actc1b</i>^<i>+/+</i>) injected with either an Actc1b ex2, Actc1b UTR or Standard Control morpholino were stained with Actinin2 (green) and F-actin (red) and phenotypes were scored as either wildtype, mild (small outgrowth of aggregates at the myosepta (arrowheads)) or severe (large outgrowth of aggregates at the myosepta (arrowheads) and Actinin2 positive aggregates throughout the muscle fibers (arrows)). Scale bar represents 50μm. B) Quantification of phenotypic severity for <i>actc1b</i>^<i>-/-</i> and wildtype siblings injected with Actc1b ex2 and Actc1b UTR MOs compared to Standard Control MO injected zebrafish. Error bars represent SEM for three independent experiments (for Actc1b ex2 MO: n = 26,14,16 <i>actc1b</i>^<i>+/+</i>, n = 31,35,29 <i>actc1b</i>^<i>+/-</i> and n = 12,11,23 <i>actc1b</i>^<i>-/-</i>, for Actc1b UTR MO: n = 23,21,17 <i>actc1b</i>^<i>+/+</i>, n = 28,30,23 <i>actc1b</i>^<i>+/-</i> and n = 8,11,8 <i>actc1b</i>^<i>-/-</i> and for Standard Control MO: n = 11,8,10 <i>actc1b</i>^<i>+/+</i>, n = 11,13,11 <i>actc1b</i>^<i>+/-</i> and n = 9,12,4 <i>actc1b</i>^<i>-/-</i>), <i>#</i>p<0.0001 using a Chi-square test. C) Locomotion assays show a significant reduction in distance travelled by <i>actc1b</i>^<i>+/+</i> injected with an Actc1b UTR MO and Actc1b ex2 MO compared to Standard Control MO, using a Kruskal-Wallis Test. Locomotion assays show a significant reduction in distance travelled by <i>actc1b</i>^<i>+/-</i> injected with an Actc1b ex2 MO compared to Control MO, using a Kruskal-Wallis Test. No difference in distance travelled is observed between <i>actc1b</i>^<i>-/-</i> mutants injected with either an Actc1b UTR MO, Actc1b ex2 MO or Standard Control MO. Error bars represent median values with interquartile range (pooled samples from 3 independent experiments for Actc1b ex2 MO: n = 25,21,20 <i>actc1b</i>^<i>+/+</i>, n = 41,53,49 <i>actc1b</i>^<i>+/-</i> and n = 20,14,15 <i>actc1b</i>^<i>-/-</i>, for Actc1b UTR MO: n = 30,24,19 <i>actc1b</i>^<i>+/+</i>, n = 42,47,57 <i>actc1b</i>^<i>+/-</i> and n = 21,23,17 <i>actc1b</i>^<i>-/-</i> and for Standard Control MO: n = 31,28,26 <i>actc1b</i>^<i>+/+</i>, n = 41,50,52 <i>actc1b</i>^<i>+/-</i> and n = 21,14,17 <i>actc1b</i>^<i>-/-</i>). *p<0.05 and #p<0.0001.</p