This work was funded in part by NIH R01 #DE021475 to CTM and an NSF Graduate Research Fellowship to NAE. Thanks to Miles Johnson for assistance with imaging and Priscilla Erickson for critical reading of the manuscript.

Tous les travaux de poisson a été approuvé par le soin et l&#39;utilisation des animaux Commission institutionnelle de l&#39;Université de Californie à Berkeley (numéro de protocole R330). Euthanasie a été réalisée en utilisant l&#39; immersion dans 0,025% Tricaine-S tamponnée avec 0,1% de bicarbonate de sodium 39. Toutes les étapes sont effectuées à température ambiante. 1. Préparation Remarque: Effectuer les étapes 1.1-1.5 dans des tubes coniques ou des flacons de scintillation qui peut sceller hermétiquement et être posés horizontalement. Les poissons ne doivent être constamment secoué, mais essayez de mélanger la solution aussi souvent que possible en retournant doucement ou en secouant la grille de tubes ou flacons pour exposer tous les côtés du poisson à la solution de coloration et de permettre aux taches de pénétrer le tissu uniformément. Ne placez pas un grand lot de poisson sur une plate-forme shaker, comme le poids lourd du liquide va briser le shaker. <ol><li> Fixer soit du poisson ou du poisson fraîchement euthanasiés stocké dans de l&#39;éthanol avec 10% de formol neutre tamponné (FBN) pendant la nuit. Vous pouvez également utiliser 4% paraformaldéhyde en solution 1 x PBS au lieu de NBF à 10%. Remarque: Si l&#39;extraction de l&#39;ADN, attachez une petite partie des caudale ou nageoires pectorales avant la fixation et à stocker dans l&#39;éthanol. </li><li> Dispose de fixer correctement dans une hotte chimique et remplacer l&#39;eau du robinet (qui est ~ pH 7,0) pendant 2 heures. Évitez d&#39;utiliser l&#39;eau de-ionisée car il peut souvent être acide et peut détartrer os. </li><li> Enlever l&#39;eau et le poisson tache avec 0,008% Alizarine Rouge S dans 1% de KOH dans de l&#39;eau pendant 24 heures. Pour les poissons de moins de 20 mm de longueur standard, utilisez 0,004% Alizarine Red S. (Faire un 100x (0,8%) solution mère de Alizarine Red S qui peut ensuite être diluée). </li><li> Retirer la tache (mise en conteneur à déchets approprié dans la hotte) et placer le poisson dans l&#39;eau du robinet pendant quelques heures. Changer l&#39;eau au besoin jusqu&#39;à ce que l&#39;eau de rinçage est la plupart du temps clair. </li><li> Retirer le poisson de l&#39;eau et le lieu dans 50% de glycérol, 0,25% de KOH pour la compensation légère et la dissection ultérieure. Remarque: Ce protocole de coloration est modifiée à partir de méthodes 40,41 décrites précédemment. </li></ol><pclass = &quot;jove_title&quot;&gt; 2. Dissection Note: Voir la Figure 1 pour un examen de la tête pertinente morphologie squelettique. <img alt="Figure 1" src="/files/ftp_upload/54056/54056fig1.jpg" /> Figure 1:. Tête Stickleback morphologie du squelette Alizarine rouge tachée tête épinoche à trois épines imagé avec fluorescence sous un ensemble de rhodamine filtre B. morphologie utile est étiqueté: Op = opercule, Sous-op = subopercle, BSR = branchiostèges rayons, Preop = préopercule, Infraorb 1-3 = infraorbital 1-3 (également appelés circumorbitals ou suborbitaux), Dent = du dentaire, Premax = prémaxillaire, Max = maxillaires , Nas = nasale, Lat. ETHM = ethmoïde latéral, PSPH = parasphénoïde, Fron = os frontal. Pour obtenir une description plus détaillée de l&#39;épinoche tête squelette, voir Anker (1974) 31. <a href="https://www.jove.com/files/ftp_upload/54056/54056fig1large.jpg" target="_blank">S&#39;il vous plaît cliquer ici pour voir une version plus grande</a>cette figure. <ol><li> Poser le poisson plat (figure 2A) et insérer une pince de forte # 5 horloger dans le côté de l&#39;œil à un angle de 45 ° ~ pour perforer la membrane qui recouvre l&#39;œil. </li><li> Peler la membrane loin de l&#39;œil, semblable à peler un couvercle de yogourt (figure 2B). </li><li> Insérez une pince ouverte derrière l&#39;œil, attraper le nerf optique derrière l&#39;œil, et enlever l&#39;œil (figure 2C). Ne pas percer l&#39;oeil car il fuira la mélanine. Si perforé, la mélanine peut être lavé au cours des étapes ultérieures. </li><li> Répétez de l&#39;autre côté. </li><li> A partir de la partie postérieure, placez une petite lame de ciseaux de dissection sous le rabat de l&#39; opercule, la lame de ciseaux glisser dorsalement ci - dessus opercule, puis couper les tissus mous grâce à la prise de l&#39; oeil (figure 2D). Couper dorsale à l&#39;os de l&#39;opercule. </li><li> Couper l&#39;os frontal (dorsale à l&#39;orbite de l&#39; oeil) (Figure 2F). </li><li> Couper l&#39;os parasphénoïde midline autour til centre des orbites (Figure 2F). </li><li> Répétez l&#39;opercule coupe sur le côté opposé. </li><li> Insérez une pince sous l&#39;opercule et pelez lentement le visage loin du corps, le parage tout tissu mou encore attaché (Figure 2G - H). Prenez soin de ne pas perturber la première rangée de branchiospines. <ol><li> Avec des pinces, détacher les cératohyaux des deux côtés de la basihyal médiane tout en détachant et en enlevant le squelette craniofacial antérieure (l&#39;ensemble de la mâchoire y compris le dentary, prémaxillaire et maxillaire, l&#39;ensemble du squelette de hyoïde y compris l&#39;opercule dermique extérieur, préopercule, subopercle, et rayons branchiostèges et la dorsale sous - jacente et des éléments endochondrale ventrales, et la partie antérieure du crâne , y compris le nez, ethmoïde latéral, et les os sous - orbitaire, voir les figures 1 et 2I). </li><li> épines pelviennes peuvent être pliés hors du corps, et peut servir de poignée pour une pince pour agripper of lorsqu&#39;il est présent. Épines verrouillent en place. Pour déverrouiller, tirez doucement la colonne vertébrale avec une pince directement loin du corps de poisson, puis pliez doucement en arrière pour presser la colonne vertébrale à plat contre les poissons. </li></ol></li><li> Insérer fermé pince postérieure et ventrale au squelette branchial (juste en dessous de tube de l&#39; intestin) et des pinces de glisser en avant, à démêler les muscles et les ligaments qui restent attachés au squelette branchial (Figure 2J - K). </li><li> En utilisant les conseils de pinces fermées, gratter les muscles attachés dorsal squelette branchial la braincase ventrale dans un postérieur à la direction antérieure (Figure 2L). </li><li> Répétez 2.9 et 2.10 sur le côté opposé. </li><li> Saisissez la base du tube de l&#39; intestin et tirez en avant pour enlever le squelette et le tube intestin branchial (Figure 2M - N). </li><li> Séparer le tube de l&#39; intestin en faisant un postérieur perpendiculaire de coupe à la fin de la cinquième ceratobranchial (Figure 2O </strong&gt;). </li><li> Après avoir enlevé tous les fragments d&#39;os restants de la boîte crânienne sur la face dorsale du squelette branchial, insérer des ciseaux dans le panier branchiale pour faire une coupe dorsale (coupe antérieure à postérieure) entre les ensembles bilatéraux de plaques dorsales de dents (Figure 3A - D). Assurez-coupe est centrée pour éviter d&#39;endommager les plaques dorsales de dents. </li><li> Faire deux coupes latérales peu profondes dans la lumière intestinale caoutchouteux à la fin postérieure du squelette branchial (extrémité antérieure du tube de l&#39; intestin) pour aider à ouvrir le squelette branchial (Figure 3E). </li><li> Placer le poisson et tous les morceaux de tissu dans un bocal et placez-le squelette branchial dans un tube à centrifuger avec 50% de glycérol, 0,25% de KOH continuer compensation douce, ou 100% de glycérol si aucune autre compensation est nécessaire. bocaux d&#39;étiquetage et les tubes avec un identifiant unique afin qu&#39;ils puissent être suivis. Le montant de la compensation requise est en grande partie fonction de la taille des poissons, gros poissons adultes (plus de 40 mm de longueur standard) nécessitent généralement de compensation supplémentaire. </li></ol><img alt="Figure 2" src="/files/ftp_upload/54056/54056fig2.jpg" /> Figure 2:. Stickleback branchiale squelette dissection Alizarine rouge tachée poisson épinoche à trois épines prête pour la dissection. L&#39;œil est dépigmentée d&#39;une vaste clairière. Les flèches bleues indiquent la direction du mouvement. (A) Vue latérale de la tête de l&#39; épinoche, antérieure se trouve à droite. (B) Enlèvement de la membrane recouvrant l&#39;œil. (C) L&#39; élimination de l&#39;oeil. (D) Dorsale coupé au- dessus de la opercule. (E) Frontal coupe osseuse. (F) parasphenoid coupé. (G - I) Retrait du squelette facial. (J) Suppression des connexions ventrales des tissus mous du squelette branchial. (K - L) Enlèvement des dorsales branchiaux connexions squelette. (&lt;strong&gt; M - N) Retrait du squelette branchial. (O) Séparer le tube intestinal du squelette branchial. Voir les étapes 2.1 à 2.16 pour plus de détails. Barre d&#39; échelle = 5 mm. <a href="https://www.jove.com/files/ftp_upload/54056/54056fig2large.jpg" target="_blank">S&#39;il vous plaît cliquer ici pour voir une version plus grande de cette figure.</a> 3. branchiale Squelette Re-coloration (si nécessaire) <ol><li> Pour colorer le plus sombre squelette branchial ou d&#39;un tissu clair plus, supprimer 50% de glycérol, solution de KOH 0,25% et laver avec 1% KOH deux fois (un lavage de cinq minutes suivi d&#39;un second lavage de 24 heures tout en secouant horizontalement sur une plate-forme shaker). </li><li> Retirer 1% de KOH et re-tache avec 0,008% Alizarine Rouge S dans 1% de KOH pendant 24 heures. </li><li> Retirer la tache et le remplacer par 1% de KOH pendant 24 heures. </li><li> Retirer la solution KOH et le remplacer par 50% de glycérol, 0,25% de KOH. </li></ol> 4. Montage branchiale Squelette <ol><li> Retirer le squelette branchialà partir de 50% de glycérol, 0,25% de KOH ou 100% de glycérol et lieu près du fond d&#39;un 22 mm x 60 mm lamelle en verre avec la face dorsale vers le haut (figure 3F). Ajouter quelques gouttes de glycérol à 50%, 0,25% de KOH ou 100% de glycérol sur le dessus du squelette branchial. Si la transition de 50% de glycérol, 0,25% de KOH à 100% de glycérol, le changement solution dans un tube à centrifuger et agiter pendant&gt; 5 min avant de monter pour équilibrer les tissus. </li><li> Rouler deux petites boules de pâte à modeler et placer sur chaque extrémité de la lamelle à agir comme entretoises. </li><li> Librement placer une deuxième lamelle sur le dessus avec une pression suffisante pour aplatir le squelette branchial antérieur (figure 3G). </li><li> Peel ouvrir le rabat dorsal gauche y compris les plaques de dents dorsales, aplatir, et glisser entre les lamelles (figure 3H). </li><li> Répéter technique avec rabat droit dorsal et pousser tout le squelette branchial loin du bord de la lamelle (figure 3I). <ol><li> Modifiernativement, tenir les deux volets dorsales ouvertes avec une pince et placer soigneusement la lamelle sur, aplatir le squelette branchial dans un mouvement fluide. </li><li> Vous pouvez également monter le squelette branchial à l&#39;envers sur une lamelle, évasement chaque côté dorsal latéralement de façon gravité ne permet pas le squelette branchial pour fermer back up. Puis couvrir avec la deuxième 22 mm x 60 mm couvercle en verre de glissement et inverser prep. Remarque: les techniques de montage différentes ont tendance à travailler mieux ou pire pour chaque individu. Essayez chacune et de voir ce qui se sent plus à l&#39;aise. </li></ol></li><li> Appuyez légèrement sur le dessus lamelle pour aplatir des balles assez pour garder le squelette branchial d&#39;argile monté à plat, mais prendre soin de ne pas écraser l&#39;échantillon. <ol><li> Au cours du processus de montage, cératobranchiaux peuvent tourner et obscurcir une rangée de rakers. Remédier au forceps entre les lamelles et réorientant les cératobranchiaux ou squelette entier branchiale coulissant. </li></ol></li><li> Magasin preps plat dans des plateaux coulissants à rotempérature om. Monté dans 100% de glycerol, préparations peuvent être conservés entre les lamelles couvre-pontés pendant au moins dix ans. pinces propres et des ciseaux avec de l&#39;isopropanol ou de l&#39;éthanol et des conseils de couverture. </li></ol><img alt="Figure 3" src="/files/ftp_upload/54056/54056fig3.jpg" /> Figure 3:. Montage plat squelette branchial Manipulation et montage du squelette branchial est représenté. Les flèches bleues indiquent la direction du mouvement. (A) branchiale dorsale squelette vers le haut. (B - D) Rotation et incision entre les plaques dorsale de la dent. (E) coupé dans un tissu mou latéral pour ouvrir davantage la base du tube intestinal. (F) branchiale squelette placé au fond d&#39;une lamelle couvre -objet prêt pour le montage. (G) Deuxième lamelle placée sur la moitié antérieure du squelette branchial ( au- dessus des plaques de dents dorsales). (H - I)montage plat du squelette branchial par l&#39;ouverture dorsale dents volets de plaque et de glissement entre deux lamelles. Voir les étapes 4.1 à 4.6 pour plus de détails. Barre d&#39; échelle = 5 mm. <a href="https://www.jove.com/files/ftp_upload/54056/54056fig3large.jpg" target="_blank">S&#39;il vous plaît cliquer ici pour voir une version plus grande de cette figure.</a>

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T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evolution+and+development+of+complex+biomechanical+systems:+300+million+years+of+fish+jaws.">Evolution and development of complex biomechanical systems: 300 million years of fish jaws.</a> Zebrafish. 2 (4), 243-257 (2005).</li><li>Lauder, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+design+and+evolution+of+the+pharyngeal+jaw+apparatus+in+euteleostean+fishes.">Functional design and evolution of the pharyngeal jaw apparatus in euteleostean fishes.</a> Zool. J. Linn. Soc. 77, 1-38 (1983).</li><li>Fraser, G. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+ancient+gene+network+is+co-opted+for+teeth+on+old+and+new+jaws.">An ancient gene network is co-opted for teeth on old and new jaws.</a> PLoS Biol. 7 (2), e1000031 (2009).</li><li>Stock, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+dentition+in+comparative+context.">Zebrafish dentition in comparative context.</a> J. Exp. Zool. B. Mol. Dev. Evol. 308, 523-549 (2007).</li><li>Liem, K., Greenwood, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+functional+approach+to+the+phylogeny+of+the+pharyngognath+teleosts.">A functional approach to the phylogeny of the pharyngognath teleosts.</a> Am. Zool. 21 (1), 83-101 (1981).</li><li>Anker, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphology+and+kinetics+of+the+head+of+the+stickleback,+Gasterosteus+aculeatus.">Morphology and kinetics of the head of the stickleback, Gasterosteus aculeatus.</a> Trans. Zool. Soc. London. 32 (5), 311-416 (1974).</li><li>Meyer, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphometrics+and+allometry+in+the+trophically+polymorphic+cichlid+fish,+Cichlusomu+citrinelfum:+Alternative+adaptations+and+ontogenetic+changes+in+shape.">Morphometrics and allometry in the trophically polymorphic cichlid fish, Cichlusomu citrinelfum: Alternative adaptations and ontogenetic changes in shape.</a> J. Zool., Lond. 221, 237-260 (1990).</li><li>Huysseune, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phenotypic+plasticity+in+the+lower+pharyngeal+jaw+dentition+of+Astatoreochromis+alluaudi+(Teleostei:+Cichlidae).">Phenotypic plasticity in the lower pharyngeal jaw dentition of Astatoreochromis alluaudi (Teleostei: Cichlidae).</a> Arch. Oral Biol. 40 (11), 1005-1014 (1995).</li><li>Muschick, M., Indermaur, A., Salzburger, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Convergent+Evolution+within+an+adaptive+radiation+of+cichlid+fishes.">Convergent Evolution within an adaptive radiation of cichlid fishes.</a> Curr. Biol. 22 (24), 2362-2368 (2012).</li><li>Cleves, P. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evolved+tooth+gain+in+sticklebacks+is+associated+with+a+cis-regulatory+allele+of+Bmp6.">Evolved tooth gain in sticklebacks is associated with a cis-regulatory allele of Bmp6.</a> Proc. Natl. Acad. Sci. 111 (38), 13912-13917 (2014).</li><li>Ellis, N. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+developmental+and+genetic+mechanisms+underlie+convergently+evolved+tooth+gain+in+sticklebacks.">Distinct developmental and genetic mechanisms underlie convergently evolved tooth gain in sticklebacks.</a> Development. (142), 2442-2451 (2015).</li><li>Tucker, A. S., Fraser, G. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evolution+and+developmental+diversity+of+tooth+regeneration.">Evolution and developmental diversity of tooth regeneration.</a> Semin. Cell Dev. Biol. 25-26, 71-80 (2014).</li><li>Erickson, P. A., Glazer, A. M., Cleves, P. A., Smith, A. S., Miller, C. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two+developmentally+temporal+quantitative+trait+loci+underlie+convergent+evolution+of+increased+branchial+bone+length+in+sticklebacks.">Two developmentally temporal quantitative trait loci underlie convergent evolution of increased branchial bone length in sticklebacks.</a> Proc. R. Soc. B. 281, (2014).</li><li>Leary, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> AVMA Guidelines for the Euthanasia of Animals. , (2013).</li><li>Bell, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evolutionary+phenetics+and+genetics.">Evolutionary phenetics and genetics.</a> Evol. Genet. Fishes. , 431-528 (1984).</li><li>Taylor, W. R., Van Dyke, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Revised+procedures+for+staining+and+clearing+small+fishes+and+other+vertebrates+for+bone+and+cartilage+study.">Revised procedures for staining and clearing small fishes and other vertebrates for bone and cartilage study.</a> Cybium. 9 (2), 107-119 (1985).</li><li>Erickson, P. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+190+base+pair,+TGF-&#946;+responsive+tooth+and+fin+enhancer+is+required+for+stickleback+Bmp6+expression.">A 190 base pair, TGF-&#946; responsive tooth and fin enhancer is required for stickleback Bmp6 expression.</a> Dev. Biol. 401 (2), 310-323 (2015).</li><li>Miller, C. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=cis-Regulatory+changes+in+Kit+ligand+expression+and+parallel+evolution+of+pigmentation+in+sticklebacks+and+humans.">cis-Regulatory changes in Kit ligand expression and parallel evolution of pigmentation in sticklebacks and humans.</a> Cell. 131 (6), 1179-1189 (2007).</li><li>Aigler, S. R., Jandzik, D., Hatta, K., Uesugi, K., Stock, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Selection+and+constraint+underlie+irreversibility+of+tooth+loss+in+cypriniform+fishes.">Selection and constraint underlie irreversibility of tooth loss in cypriniform fishes.</a> Proc. Natl. Acad. Sci. U. S. A. 111 (21), 7707-7712 (2014).</li><li>Pasco-Viel, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evolutionary+trends+of+the+pharyngeal+dentition+in+Cypriniformes+(Actinopterygii+Ostariophysi).">Evolutionary trends of the pharyngeal dentition in Cypriniformes (Actinopterygii Ostariophysi).</a> PLoS One. 5 (6), e11293 (2010).</li></ol>

The authors have nothing to disclose.

The branchial skeleton is a complex set of bones in the throat of a fish that manipulates, filters, and masticates food items on their way to the esophagus. Many interesting trophic traits including the patterning of gill rakers, pharyngeal teeth, and branchial bones vary across and within species. The majority of these traits are difficult to near impossible to accurately measure with the branchial skeleton in situ (e.g., gill raker length, branchial bone length). This flat-mounting protocol places all...

Une quantité incroyable de diversité existe dans le squelette de la tête chez les vertébrés, en particulier chez les poissons. Dans de nombreux cas , cette diversité facilite les stratégies d&#39;alimentation différentes : 1 - 4, et peut impliquer des changements majeurs à la fois un motif craniofaciale externe et interne. Le squelette branchial est situé à l&#39;intérieur de la gorge d&#39;un poisson et qui entoure la majeure partie de la cavité buccale. Le squelette branchial est composé de 5 segments en série homologues, les quatre antérieure qui soutiennent les branchies. Ensemble , ces cinq segments fonctionnent comme une interface entre les poissons et leur nourriture 5. Variation dans une multitude de traits, y compris branchiospines, dents pharyngiennes, et les os branchiaux contribuent à la recherche de nourriture efficace sur les différents types d&#39;aliments. Épinoches ont subi une radiation adaptative après des formes océaniques ancestrales colonisées lacs d&#39;eau douce et les criques tout l&#39;hémisphère nord. Le changement dans le régime alimentairedu petit zooplancton dans l&#39;océan à des proies plus grosses en eau douce a donné lieu à une variation trophique dramatique dans plusieurs traits craniofaciales 6. Alors que de nombreuses études ont porté sur les différences craniofaciales externes dans épinoches 7 - 13, des changements importants craniofaciales évoluent de façon répétée dans le squelette branchial interne. La possibilité de créer des hybrides fertiles entre les populations d&#39;épinoches morphologiquement distinctes offre une excellente occasion de cartographier la base génétique des changements ont évolué vers le squelette branchial. Un trait trophique d&#39;importance écologique est la structuration des branchicténies, des os dermiques périodiques qui tapissent les faces antérieure et postérieure des os branchies et sont utilisés pour filtrer les proies. Les poissons qui se nourrissent généralement sur ​​de petits objets de proies ont tendance à avoir plus et plus densément maillant espacées branchicténies par rapport aux poissons qui se nourrissent de proies plus grosses 14,15. Variation branchicténies a été rapporté à la fois wSEIN et entre les espèces 14-19, et les aspects de la branchicténies patterning contribuent à des niches trophiques et remise en forme 16. Des décennies de recherche ont largement documenté nombre maillant racloir et la variation de longueur dans épinoches à trois épines 17 - 21; Toutefois, ces études se concentrent généralement sur la première rangée de branchiospines. Des travaux récents ont montré la modularité dans le contrôle génétique du nombre maillant racloir à travers le squelette branchial 22,23 et à travers une seule ligne dans branchicténies espacement 23 et la longueur 24 mettant en évidence l&#39;importance d&#39;étudier plus de ramer un ou une seule raker maillant pour comprendre le base génétique du développement de la réduction des branchies racloir. Un second trait trophique de signification à la fois écologique et biomédicale est le motif de dents pharyngées. Dents chez les poissons peuvent être situés à la fois dans la mâchoire, et par voie orale dans le squelette branchial, connu sous le nom de dents pharyngées. dents orales sont utilisées principalement pour prey capturer tout les dents pharyngées sont utilisés pour la mastication et la proie manipulation 25 - 27. Les deux ensembles forment par l&#39; intermédiaire des mécanismes de développement partagés et sont considérés comme de développement homologue 28. Modularité intéressante se produit où certaines espèces, comme le poisson zèbre, le manque de dents pharyngée orale et dorsale 29 tandis que d&#39; autres espèces ont plusieurs cératobranchiaux dentées, pharyngobranchials, et parfois dentées basihyal et hypobranchiaux 30. Dans épinoches dents pharyngiennes se trouvent sur ​​le côté ventral et cinquième ceratobranchial dorsalement sur ​​la partie antérieure et postérieure pharyngobranchials 31. Cinématique sur l&#39; épinoche alimentation montrent la mâchoire orale est principalement utilisé pour la capture des proies et de faciliter l&#39; aspiration d&#39; alimentation 9 en laissant la mastication à la mâchoire du pharynx. Dans cichlidés, inférieure morphologie pharyngée de la mâchoire varie considérablement 32,33 et a été montré pour être adaptatif et en corrélation avec niche trophique 34. Multiles populations d&#39; eau douce d&#39;épinoches ples ont évolué une augmentation spectaculaire de pharyngée ventral nombre de dents 23,35,36. Des travaux récents ont montré que la base génétique du développement de ce gain de dent évolué est en grande partie distincte dans deux populations dérivées indépendamment des épinoches d&#39;eau douce 36. Contrairement à dents de mammifères, les poissons se régénèrent leurs dents en permanence tout au long de la vie adulte 37. Ces deux populations précédemment décrites en eau douce dentées élevées ont évolué un taux de remplacement de la dent accélérée, fournissant un système de vertébrés rares pour étudier la base génétique de la régénération 36. Un troisième trait trophique qui a évolué à plusieurs reprises dans les épinoches d&#39;eau douce est plus long os épibranchiales et ceratobranchial, les branchies homologues arc segmentaire de la mâchoire supérieure et inférieure, respectivement 38. Plus longs os branchiaux confèrent une plus grande cavité buccale et probablement sont adaptatifs pour permettre à de plus grandes proies soient consumed. En outre, dans d&#39; autres poissons, les os épibranchiales sont importants pour la dépression des dorsale pharynx dents plaques 25. Comme branchicténies et les dents pharyngées, les os branchiaux sont internes et donc, difficile de visualiser facilement ou quantifier. Nous présentons ici un protocole détaillé pour disséquer et plat monter le squelette branchial, permettant une visualisation facile et la quantification d&#39;une variété de traits craniofaciales importants. Bien que ce protocole décrit une dissection de l&#39;épinoche, cette même méthode fonctionne sur une variété d&#39;autres poissons.

<table>
	<tbody>
		<tr>
			<td>Sodium Hydroxide (KOH)</td>
			<td>EMD</td>
			<td>PX1480-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Glycerol</td>
			<td>Sigma-Alderich</td>
			<td>G7893-4L</td>
			<td></td>
		</tr>
		<tr>
			<td>10% Neutral Buffered Formalin (NBF)</td>
			<td>Azer Scientific</td>
			<td>NBF-4-G</td>
			<td></td>
		</tr>
		<tr>
			<td>Alizarin Red S</td>
			<td>EMD</td>
			<td>116-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Cover Glasses 22x60mm</td>
			<td>VWR</td>
			<td>16004-350</td>
			<td></td>
		</tr>
		<tr>
			<td>100x10mm Glass Petri Dish</td>
			<td>Kimble Chase</td>
			<td>23064-10010</td>
			<td>To dissect samples on</td>
		</tr>
		<tr>
			<td>Sylgard 184 Silicone Elastomer Kit</td>
			<td>Ellsworth Adhesives</td>
			<td>184 SIL ELAST KIT 0.5KG</td>
			<td>Can be poured into glass or plastic petri dishes to make dissecting plates</td>
		</tr>
		<tr>
			<td>Modeling Clay</td>
			<td>Sargent Art</td>
			<td>22-4000</td>
			<td>1lb cream</td>
		</tr>
		<tr>
			<td>Scintillation Vials (case of 500)</td>
			<td>Wheaton</td>
			<td>66021-668</td>
			<td>Borosilicate Glass with Screw Cap</td>
		</tr>
		<tr>
			<td>Forceps-Dumont #5 Inox (Biologie tip)</td>
			<td>FST</td>
			<td>11252-20</td>
			<td>Dumostars are an alternative</td>
		</tr>
		<tr>
			<td>Dissecting Scissors&#160;</td>
			<td>FST</td>
			<td>15003-08</td>
			<td>Alternate sizes are available depending on size of sample</td>
		</tr>
		<tr>
			<td>Dissecting Microscope</td>
			<td>Leica</td>
			<td>S6E with KL300 LED</td>
			<td>Many other models work nicely, having a flat base helps</td>
		</tr>
		<tr>
			<td>Microcentrifuge Tubes 1.7mL</td>
			<td>Denville</td>
			<td>C-2170</td>
			<td></td>
		</tr>
		<tr>
			<td>Cardboard slide tray</td>
			<td>Fisher</td>
			<td>12-587-10</td>
			<td></td>
		</tr>
	</tbody>
</table>

dissection and flat-mounting of the threespine stickleback branchial skeleton

The posterior pharyngeal segments of the vertebrate head give rise to the branchial skeleton, the primary site of food processing in fish. The morphology of the fish branchial skeleton is matched to a species' diet. Threespine stickleback fish (Gasterosteus aculeatus) have emerged as a model system to study the genetic and developmental basis of evolved differences in a variety of traits. Marine populations of sticklebacks have repeatedly colonized countless new freshwater lakes and creeks. Adaptation to the new diet in these freshwater environments likely underlies a series of craniofacial changes that have evolved repeatedly in independently derived freshwater populations. These include three major patterning changes to the branchial skeleton: reductions in the number and length of gill raker bones, increases in pharyngeal tooth number, and increased branchial bone lengths. Here we describe a detailed protocol to dissect and flat-mount the internal branchial skeleton in threespine stickleback fish. Dissection of the entire three-dimensional branchial skeleton and mounting it flat into a largely two-dimensional prep allows for the easy visualization and quantification of branchial skeleton morphology. This dissection method is inexpensive, fast, relatively easy, and applicable to a wide variety of fish species. In sticklebacks, this efficient method allows the quantification of skeletal morphology in genetic crosses to map genomic regions controlling craniofacial patterning.

<html> Ce protocole se traduit par un squelette branchial monté disséqué et plat (figure 4) où une variété de traits trophiques importants peuvent être quantifiés. Du point de vue dorsale, toutes les lignes de branchicténies, toutes les plaques pharyngée à dents, et presque tous les os branchies peuvent être facilement visualisées et quantifiées 22 - 24,35,36,38,42. Alizarine Red S fluoresces également sur ​​un rhodamine...

Watch this Scientific Journal Video about Dissection and Flat-mounting of the Threespine Stickleback Branchial Skeleton at JoVE.com

Dissection and Flat-mounting of the Threespine Stickleback Branchial Skeleton

The branchial skeleton, including gill rakers, pharyngeal teeth, and branchial bones, serves as the primary site of food processing in most fish. Here we describe a protocol to dissect and flat-mount this internal skeleton in threespine sticklebacks. This method is also applicable to a variety of other fish species.

Dissection et Flat-montage de l&#39;épinoche à trois épines branchiale Squelette

The posterior pharyngeal segments of the vertebrate head give rise to the branchial skeleton, the primary site of food processing in fish. The morphology ...

dissection-flat-mounting-threespine-stickleback-branchial

Research

JoVE Journal

Biology

9.7K vues.  University of California, Berkeley. The branchial skeleton, including gill rakers, pharyngeal teeth, and branchial bones, serves as the primary site of food processing in most fish. Here we describe a protocol to dissect and flat-mount this internal skeleton in threespine sticklebacks. This method is also applicable to a variety of other fish species. 

Vidéo: Dissection and Flat-mounting of the Threespine Stickleback Branchial Skeleton

Dissection of Drosophila Ovaries

Dissection des ovaires chez la drosophile

Recherche

Biologie

Drosophila Larval NMJ Dissection

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use of the Drosophila motor system is due to its high accessibility. It can be analyzed with single-cell resolution. There are 30 muscles per hemisegment whose arrangement within the peripheral body wall are known. A total of 35 motor neurons attach to these muscles in a pattern that has high fidelity. Using molecular biology and genetics, one can create transgenic animals or mutants. Then, one can study the developmental consequences on the morphology and function of the NMJ. In order to access the NMJ for study, it is necessary to carefully dissect each larva. In this article we demonstrate how to properly dissect Drosophila larvae for study of the NMJ by removing all internal organs while leaving the body wall intact. This technique is suitable to prepare larvae for imaging, immunohistochemistry, or electrophysiology.

This protocol demonstrates how to dissect Drosophila larvae in preparation for immunohistochemistry and/or imaging of the neuromuscular junction.

Drosophile larvaire JNM Dissection

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use ...

Gross Dissection of the Stomach of the Lobster, Homarus Americanus

The stomach of the American lobster (Homarus americanus) is located in the cephalothorax, between the rostrum and the cervical groove. The anterior end of the stomach is defined by the mouth opening and the posterior end by the bottom of the pylorus. Along the dorsal side of the stomach lies the stomatogastric nervous system (STNS). This nervous system, which contains rhythmic networks that underlie feeding behavior, is an established model system for studying rhythm generating networks and neuromodulation 1,2. While it is possible to study this system in vivo 3, the STNS continues to produce its rhythmic activity when isolated in vitro. In order to study this system in vitro the stomach must be removed from the animal. This video article describes how the stomach can be dissected from the American lobster. In an accompanying video article4 we demonstrate how the STNS can be isolated from the stomach.

Gross Dissection of the Stomach of the Lobster, Homarus Americanus

We describe the gross dissection of the stomach of the American lobster (Homarus americanus).

Dissection brut de l&#39;estomac du homard, Homarus americanus</em

The stomach of the American lobster (Homarus americanus) is located in the cephalothorax, between the rostrum and the cervical groove. The anterior end of ...

Dissection of Hippocampal Dentate Gyrus from Adult Mouse

The hippocampus is one of the most widely studied areas in the brain because of its important functional role in memory processing and learning, its remarkable neuronal cell plasticity, and its involvement in epilepsy, neurodegenerative diseases, and psychiatric disorders. The hippocampus is composed of distinct regions; the dentate gyrus, which comprises mainly granule neurons, and Ammon's horn, which comprises mainly pyramidal neurons, and the two regions are connected by both anatomic and functional circuits. Many different mRNAs and proteins are selectively expressed in the dentate gyrus, and the dentate gyrus is a site of adult neurogenesis; that is, new neurons are continually generated in the adult dentate gyrus. To investigate mRNA and protein expression specific to the dentate gyrus, laser capture microdissection is often used. This method has some limitations, however, such as the need for special apparatuses and complicated handling procedures. In this video-recorded protocol, we demonstrate a dissection technique for removing the dentate gyrus from adult mouse under a stereomicroscope. Dentate gyrus samples prepared using this technique are suitable for any assay, including transcriptomic, proteomic, and cell biology analyses. We confirmed that the dissected tissue is dentate gyrus by conducting real-time PCR of dentate gyrus-specific genes, tryptophan 2,3-dioxygenase (TDO2) and desmoplakin (Dsp), and Ammon's horn enriched genes, Meis-related gene 1b (Mrg1b) and TYRO3 protein tyrosine kinase 3 (Tyro3). The mRNA expressions of TDO2 and Dsp in the dentate gyrus samples were detected at obviously higher levels, whereas Mrg1b and Tyro3 were lower levels, than those in the Ammon's horn samples. To demonstrate the advantage of this method, we performed DNA microarray analysis using samples of whole hippocampus and dentate gyrus. The mRNA expression of TDO2 and Dsp, which are expressed selectively in the dentate gyrus, in the whole hippocampus of alpha-CaMKII+/- mice, exhibited 0.037 and 0.10-fold changes compared to that of wild-type mice, respectively. In the isolated dentate gyrus, however, these expressions exhibited 0.011 and 0.021-fold changes compared to that of wild-type mice, demonstrating that gene expression changes in dentate gyrus can be detected with greater sensitivity. Taken together, this convenient and accurate dissection technique can be reliably used for studies focused on the dentate gyrus.

A dissection technique for removal of the dentate gyrus from adult mouse under a stereomicroscope was demonstrated in this video-recorded protocol.

Dissection de l&#39;hippocampe gyrus denté de souris adulte

The hippocampus is one of the most widely studied areas in the brain because of its important functional role in memory processing and learning, its ...

Dissection of Organs from the Adult Zebrafish

Over the last 20 years, the zebrafish has become a powerful model organism for understanding vertebrate development and disease.  Although experimental analysis of the embryo and larva is extensive and the morphology has been well documented, descriptions of adult zebrafish anatomy and studies of development of the adult structures and organs, together with techniques for working with adults are lacking. The organs of the larva undergo significant changes in their overall structure, morphology, and anatomical location during the larval to adult transition.  Externally, the transparent larva develops its characteristic adult striped pigment pattern and paired pelvic fins, while internally, the organs undergo massive growth and remodeling.  In addition, the bipotential gonad primordium develops into either testis or ovary.  This protocol identifies many of the organs of the adult and demonstrates methods for dissection of the brain, gonads, gastrointestinal system, heart, and kidney of the adult zebrafish. The dissected organs can be used for in situ hybridization, immunohistochemistry, histology, RNA extraction, protein analysis, and other molecular techniques.  This protocol will assist in the broadening of studies in the zebrafish to include the remodeling of larval organs, the morphogenesis of organs specific to the adult and other investigations of the adult organ systems.

This protocol describes a procedure for identifying and dissecting organs from the adult zebrafish.

Dissection d&#39;organes sur le poisson zèbre pour adultes

Over the last 20 years, the zebrafish has become a powerful model organism for understanding vertebrate development and disease.  Although experimental ...

Dissection and Staining of Drosophila Larval Ovaries

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells form, and how they interact with their environment is therefore crucial for understanding development, homeostasis and disease. The ovary of the fruit fly Drosophila melanogaster has served as an influential model for the interaction of germ line stem cells (GSCs) with their somatic support cells (niche) 1, 2. The known location of the niche and the GSCs, coupled to the ability to genetically manipulate them, has allowed researchers to elucidate a variety of interactions between stem cells and their niches 3-12.
Despite the wealth of information about mechanisms controlling GSC maintenance and differentiation, relatively little is known about how GSCs and their somatic niches form during development. About 18 somatic niches, whose cellular components include terminal filament and cap cells (Figure 1), form during the third larval instar 13-17. GSCs originate from primordial germ cells (PGCs). PGCs proliferate at early larval stages, but following the formation of the niche a subgroup of PGCs becomes GSCs 7, 16, 18, 19. Together, the somatic niche cells and the GSCs make a functional unit that produces eggs throughout the lifetime of the organism.
Many questions regarding the formation of the GSC unit remain unanswered. Processes such as coordination between precursor cells for niches and stem cell precursors, or the generation of asymmetry within PGCs as they become GSCs, can best be studied in the larva. However, a methodical study of larval ovary development is physically challenging. First, larval ovaries are small. Even at late larval stages they are only 100&mu;m across. In addition, the ovaries are transparent and are embedded in a white fat body. Here we describe a step-by-step protocol for isolating ovaries from late third instar (LL3) Drosophila larvae, followed by staining with fluorescent antibodies. We offer some technical solutions to problems such as locating the ovaries, staining and washing tissues that do not sink, and making sure that antibodies penetrate into the tissue. This protocol can be applied to earlier larval stages and to larval testes as well.

Dissection and Staining of Drosophila Larval Ovaries

How niches and stem cells form during development is an important question with practical implications. In the Drosophila ovary, germ line stem cells and their somatic niches form during larval development. This video demonstrates how to dissect, stain and mount female gonads from late third instar (LL3) Drosophila larvae.

Dissection et la coloration des Drosophile Ovaires larvaires

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells ...

Dissection of the Adult Zebrafish Kidney

Researchers working in the burgeoning field of adult stem cell biology seek to understand the signals that regulate the behavior and function of stem cells during normal homeostasis and disease states. The understanding of adult stem cells has broad reaching implications for the future of regenerative medicine1. For example, better knowledge about adult stem cell biology can facilitate the design of therapeutic strategies in which organs are triggered to heal themselves or even the creation of methods for growing organs in vitro that can be transplanted into humans1. The zebrafish has become a powerful animal model for the study of vertebrate cell biology2. There has been extensive documentation and analysis of embryonic development in the zebrafish3. Only recently have scientists sought to document adult anatomy and surgical dissection techniques4, as there has been a progressive movement within the zebrafish community to broaden the applications of this research organism to adult studies. For example, there are expanding interests in using zebrafish to investigate the biology of adult stem cell populations and make sophisticated adult models of diseases such as cancer5. Historically, isolation of the zebrafish adult kidney has been instrumental for studying hematopoiesis, as the kidney is the anatomical location of blood cell production in fish6,7. The kidney is composed of nephron functional units found in arborized arrangements, surrounded by hematopoietic tissue that is dispersed throughout the intervening spaces. The hematopoietic component consists of hematopoietic stem cells (HSCs) and their progeny that inhabit the kidney until they terminally differentiate8. In addition, it is now appreciated that a group of renal stem/progenitor cells (RPCs) also inhabit the zebrafish kidney organ and enable both kidney regeneration and growth, as observed in other fish species9-11. In light of this new discovery, the zebrafish kidney is one organ that houses the location of two exciting opportunities for adult stem cell biology studies. It is clear that many outstanding questions could be well served with this experimental system. To encourage expansion of this field, it is beneficial to document detailed methods of visualizing and then isolating the adult zebrafish kidney organ. This protocol details our procedure for dissection of the adult kidney from both unfixed and fixed animals.
Dissection of the kidney organ can be used to isolate and characterize hematopoietic and renal stem cells and their offspring using established techniques such as histology, fluorescence activated cell sorting (FACS)11,12, expression profiling13,14, and transplantation11,15. 
We hope that dissemination of this protocol will provide researchers with the knowledge to implement broader use of zebrafish studies that ultimately can be translated for human application.

The zebrafish kidney is home to both renal and hematopoietic adult stem/progenitor cells, and represents an outstanding opportunity to study these cell types and their progeny in a vertebrate model organism. Here, we demonstrate a detailed dissection procedure that enables the researcher to identify and surgically remove the adult zebrafish kidney, which can be used for applications such as cell isolation, transplantation, and expression studies of kidney and/or blood cell populations.

Dissection du rein Zebrafish adultes

Researchers working in the burgeoning field of adult stem cell biology seek to understand the signals that regulate the behavior and function of stem ...

Dissection and Mounting of Drosophila Pupal Eye Discs

The Drosophila melanogaster eye disc is a powerful system that can be used to study many different biological processes. It contains approximately 800 separate eye units, termed ommatidia1. Each ommatidium contains eight neuronal photoreceptors that develop from undifferentiated cells following the passage of the morphogenetic furrow in the third larval instar2. Following the sequential differentiation of the photoreceptors, non-neuronal cells develop, including cone and pigment cells, along with mechanosensory bristle cells3. Final differentiation processes, including the structured arrangement of all the ommatidial cell types, programmed cell death of undifferentiated cell types and rhodopsin expression, occurs through the pupal phase4-7. This technique focuses on manipulating the pupal eye disc, providing insight and instruction on how to dissect the eye disc during the pupal phase, which is inherently more difficult to perform than the commonly dissected third instar eye disc. This technique also provides details on immunostaining to allow the visualization of various proteins and other cell components.

Dissection and Mounting of Drosophila Pupal Eye Discs

The goal of this technique is to enable researchers to perform dissection, immunostaining and mounting of pupal eye discs from Drosophila melanogaster of any age.

Dissection et de montage de<em&gt; Drosophila</em&gt; pupes disques yeux

The Drosophila melanogaster eye disc is a powerful system that can be used to study many different biological processes.  It contains approximately 800 ...

Dissection of the Auditory Bulla in Postnatal Mice: Isolation of the Middle Ear Bones and Histological Analysis

In most mammals, auditory ossicles in the middle ear, including the malleus, incus and stapes, are the smallest bones. In mice, a bony structure called the auditory bulla houses the ossicles, whereas the auditory capsule encloses the inner ear, namely the cochlea and semicircular canals. Murine ossicles are essential for hearing and thus of great interest to researchers in the field of otolaryngology, but their metabolism, development, and evolution are highly relevant to other fields. Altered bone metabolism can affect hearing function in adult mice, and various gene-deficient mice show changes in morphogenesis of auditory ossicles in utero. Although murine auditory ossicles are tiny, their manipulation is feasible if one understands their anatomical orientation and 3D structure. Here, we describe how to dissect the auditory bulla and capsule of postnatal mice and then isolate individual ossicles by removing part of the bulla. We also discuss how to embed the bulla and capsule in different orientations to generate paraffin or frozen sections suitable for preparation of longitudinal, horizontal, or frontal sections of the malleus. Finally, we enumerate anatomical differences between mouse and human auditory ossicles. These methods would be useful in analyzing pathological, developmental and evolutionary aspects of auditory ossicles and the middle ear in mice.

We present a protocol to isolate the auditory bulla, capsule, and ossicles from postnatal mice for whole mount and histological analysis.

Dissection de l&#39;Auditory Bulla chez les souris postnatale: Isolation des os de l&#39;oreille moyenne et l&#39;analyse histologique

In most mammals, auditory ossicles in the middle ear, including the malleus, incus and stapes, are the smallest bones. In mice, a bony structure called ...

Reverse Dissection and DiceCT Reveal Otherwise Hidden Data in the Evolution of the Primate Face

Facial expressions, or facial displays, of social or emotional intent are produced by many mammalian taxa as a means of visually communicating with conspecifics at a close range. These displays are achieved by contraction of the mimetic muscles, which are skeletal muscle attached to the dermis of the face. Reverse dissection, removing the full facial mask from the skull and approaching mimetic muscles in reverse, is an effective but destructive way of revealing the morphology of mimetic muscles but it is destructive. DiceCT is a novel mechanism for visualizing skeletal muscles, including mimetic muscles, and isolating individual muscle fascicles for quantitative measurement. Additionally, DiceCT provides a non-destructive mechanism for visualizing muscles. The combined techniques of reverse dissection and DiceCT can be used to assess the evolutionary morphology of mimetic musculature as well as potential contraction strength and velocity in these muscles. This study further demonstrates that DiceCT can be used to accurately and reliably visualize mimetic muscles as well as reverse dissection and provide a non-destructive method for sampling mimetic muscles.

Facial expressions are a mode of visual communication produced by mimetic muscles. Here, we present protocols for the novel techniques of reverse dissection and DiceCT to fully visualize and assess mimetic muscles. These combined techniques can examine both morphological and physiological aspects of mimetic musculature to determine functional aspects.

Inverser la Dissection et données DiceCT Reveal autrement cachés dans l’évolution du visage de primates

Facial expressions, or facial displays, of social or emotional intent are produced by many mammalian taxa as a means of visually communicating with ...

Dissection et Flat-montage de l'épinoche à trois épines branchiale Squelette

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