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.

Tutti i lavori di pesce è stato approvato dal Comitato Istituzionale cura degli animali e l&#39;uso della University of California-Berkeley (numero di protocollo R330). L&#39;eutanasia è stata effettuata utilizzando immersione in 0,025% tricaine-S tamponata con 0,1% di bicarbonato di sodio 39. Tutte le fasi sono eseguite a temperatura ambiente. 1. Preparazione Nota: Eseguire i passi 1.1-1.5 in tubi conici o fiale di scintillazione in grado di sigillare ermeticamente ed essere fissate in orizzontale. I pesci non devono essere costantemente scosso, ma cercare di miscelare la soluzione più spesso possibile capovolgendo delicatamente o agitando il rack di tubi o fiale per esporre tutti i lati del pesce alla soluzione colorante e consentire macchia di penetrare il tessuto in modo uniforme. Non posizionare una grande partita di pesce su un agitatore piattaforma, come il peso del liquido si romperà lo shaker. <ol><li> Fissare o pesce appena eutanasia o pesce conservato in etanolo al 10% neutra tamponata formalina (NBF) durante la notte. In alternativa, utilizzare il 4% Paraformaldeide in soluzione PBS 1x invece del 10% NBF. Nota: se l&#39;estrazione del DNA, clip di una piccola porzione di caudale o pinne pettorali prima della fissazione e conservare in etanolo. </li><li> Smaltire correttamente fix in una cappa chimica e sostituire con acqua di rubinetto (cioè ~ pH 7,0) per 2 ore. Evitare di utilizzare acqua deionizzata come spesso può essere acido e può decalcificazione ossea. </li><li> Rimuovere l&#39;acqua e pesce macchia con 0,008% Alizarina Rossa S in 1% KOH in acqua per 24 ore. Per il pesce meno di 20 mm di lunghezza standard, utilizzare 0.004% Alizarina Rossa S. (Fai un 100x (0,8%) soluzione madre di Alizarina Rossa S che possono poi essere diluita). </li><li> Rimuovere macchia (mettendo in idoneo contenitore per rifiuti in cappuccio) e il luogo di pesce in acqua di rubinetto per pochi ore. Cambiare l&#39;acqua, se necessario fino a quando l&#39;acqua di risciacquo è per lo più chiara. </li><li> Rimuovere l&#39;acqua e il luogo di pesce in glicerolo al 50%, 0,25% KOH per la compensazione mite e la successiva dissezione. Nota: Questo protocollo di colorazione viene modificato dai metodi descritti in precedenza 40,41. </li></ol><pclass = &quot;jove_title&quot;&gt; 2. Dissezione Nota: Vedere la Figura 1 per una revisione delle pertinenti testa morfologia scheletrica. <img alt="Figura 1" src="/files/ftp_upload/54056/54056fig1.jpg" /> Figura 1:. Stickleback testa morfologia scheletrica Alizarina Rossa macchiato testa Spinarello ripreso con fluorescenza sotto un set di filtri rodamina B. morfologia Utile è etichettato: Op = opercolo, Subop = subopercle, BSRS = branchiostegi raggi, preop = preopercolo, Infraorb 1-3 = infraorbitale 1-3 (chiamati anche circumorbitals o suborbitals), Dent = dentario, Premax = premaxilla, Max = mascellare , Nas = nasale, Lat. ETHM = etmoide laterale, Psph = parasphenoid, Fron = osso frontale. Per una descrizione più dettagliata della testa di scheletro spinarello, vedere Anker (1974) 31. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54056/54056fig1large.jpg" target="_blank">Si prega di cliccare qui per vedere una versione più grande</a>questa figura. <ol><li> Appoggiare il pesce piatto (Figura 2A) e inserire pinze taglienti # 5 orologiai nel lato dell&#39;occhio ad un angolo di ~ 45 ° per forare la membrana che ricopre l&#39;occhio. </li><li> Peel la membrana distanza dall&#39;occhio, simile alla pelatura un coperchio yogurt (Figura 2B). </li><li> Inserire una pinza aperta dietro l&#39;occhio, afferrare del nervo ottico dietro l&#39;occhio, e rimuovere l&#39;occhio (Figura 2C). Non forare l&#39;occhio in quanto perderà melanina. Se forato, la melanina può essere lavato via durante le fasi successive. </li><li> Ripetere sull&#39;altro lato. </li><li> A partire dal posteriore, posizionare una piccola lama da dissezione forbice sotto il lembo opercolo, lama trascinare forbice dorsale sopra opercolo, quindi tagliare i tessuti molli fino alla cavità oculare (Figura 2D). Tagliare dorsale all&#39;osso opercolo. </li><li> Tagliare l&#39;osso frontale (dorsale alla cavità oculare) (Figura 2E). </li><li> Tagliare l&#39;osso parasphenoid linea mediana intorno tegli centro delle orbite (Figura 2F). </li><li> taglio opercolo Ripetere sul lato opposto. </li><li> Inserire una pinza sotto l&#39;opercolo e sbucciare lentamente la faccia lontano dal corpo, tagliare qualsiasi tessuto molle ancora attaccato (Figura 2G - H). Fare attenzione a non interrompere la prima fila di branchiospine. <ol><li> Con una pinza, staccare i ceratohyals su entrambi i lati del basihyal linea mediana, mentre staccando e rimuovendo lo scheletro cranio anteriore (l&#39;intera mascella compreso il dentario, premaxilla, e della mascella, l&#39;intero scheletro ioide compreso il opercolo dermico esterno, preopercolo, subopercle, e raggi branchiostegi e dorsale alla base e gli elementi endochondral ventrali, e la parte anteriore del cranio, compresi il naso, etmoide laterali e ossa infraorbitale, vedi figure 1 e 2I). </li><li> spine pelvico possono essere ripiegati fuori dal corpo, e può servire come una maniglia per pinze per afferrare hold of quando presente. Spine bloccano in posizione. Per sbloccare, tirare delicatamente colonna vertebrale con una pinza direttamente via dal corpo di pesce, poi flettere leggermente posteriormente a premere schiena piatta contro il pesce. </li></ol></li><li> Inserire chiuso pinza posteriore e ventrale di scheletro branchiale (appena al di sotto del tubo intestinale) e pinze trascinare anteriormente, prendere in giro a parte i restanti muscoli e legamenti attaccati allo scheletro branchiale (Figura 2J - K). </li><li> Utilizzando la punta delle pinze chiuse, raschiare via i muscoli connessi dorsale scheletro branchiale alla scatola cranica ventrale in un posteriore in direzione anteriore (Figura 2L). </li><li> Ripetere 2.9 e 2.10 sul lato opposto. </li><li> Afferrare la base del tubo intestinale e tirare anteriormente per rimuovere lo scheletro e intestino branchiale tubo (Figura 2 M - N). </li><li> Separare il tubo intestinale effettuando un taglio perpendicolare posteriore alla fine del quinto ceratobranchial (Figura 2O </strong&gt;). </li><li> Dopo aver rimosso eventuali frammenti ossei rimanenti dalla scatola cranica sul lato dorsale dello scheletro branchiale, inserire le forbici nel cestello branchiale per fare un taglio dorsale (taglio anteriore a posteriore) tra i set bilaterali di piatti dorsale dei denti (Figura 3A - D). Assicurarsi taglio è centrato per evitare di danneggiare le piastre dorsali dei denti. </li><li> Fare due tagli laterali poco profonde nel lume intestinale gommose alla fine posteriore dello scheletro branchiale (estremità anteriore del tubo intestinale) per assistere con l&#39;apertura dello scheletro branchiale (Figura 3E). </li><li> Mettere pesce e tutti i pezzi di tessuto in un vaso e posizionare lo scheletro branchiale in una provetta con il 50% glicerolo, 0,25% KOH continuare compensazione dolce, o 100% glicerolo, se non è necessaria un&#39;ulteriore compensazione. vasetti di etichette e tubi con un identificatore univoco in modo che possano essere monitorati. La quantità di compensazione previsto è in gran parte una funzione della dimensione del pesce, grandi pesci adulti (oltre 40 mm di lunghezza standard) in genere richiedono ulteriore compensazione. </li></ol><img alt="figura 2" src="/files/ftp_upload/54056/54056fig2.jpg" /> Figura 2:. Stickleback branchiale scheletro dissezione Alizarina Rossa macchiato pesci Spinarello pronto per la dissezione. L&#39;occhio è depigmented da un&#39;ampia radura. Le frecce blu indicano la direzione del moto. (A) Vista laterale della testa spinarello, anteriore è a destra. (B) La rimozione della membrana che ricopre l&#39;occhio. (C) La rimozione dell&#39;occhio. (D) dorsale taglio sopra il opercolo. Taglio dell&#39;osso (E) frontale. (F) Parasphenoid tagliato. (G - I) Rimozione dello scheletro facciale. (J) Rimozione delle connessioni dei tessuti molli scheletro branchiale ventrali. (K - L) Rimozione di dorsali connessioni scheletro branchiali. (&lt;strong&gt; M - N) Rimozione dello scheletro branchiale. (O) Separare il tubo intestinale dallo scheletro branchiale. Vedere i punti da 2.1 a 2.16 per maggiori dettagli. Barra di scala = 5 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54056/54056fig2large.jpg" target="_blank">Clicca qui per vedere una versione più grande di questa figura.</a> 3. branchiale scheletro Re-colorazione (se necessario) <ol><li> Per colorare la più scura scheletro branchiale o tessuto chiaro più, rimuovere il 50% glicerolo, soluzione di KOH 0,25% e lavare con 1% KOH due volte (un lavaggio cinque minute seguito da un secondo lavaggio 24 ore agitando orizzontalmente su un agitatore piattaforma). </li><li> Rimuovere 1% KOH e ri-macchia con 0,008% Alizarina Rossa S in 1% KOH per 24 ore. </li><li> Rimuovere macchia e sostituirlo con 1% KOH per 24 ore. </li><li> Rimuovere la soluzione KOH e sostituirla con glicerolo al 50%, 0,25% KOH. </li></ol> 4. Montaggio branchiale scheletro <ol><li> Rimuovere scheletro branchialedal 50% glicerolo, 0,25% KOH o 100% glicerolo e posto vicino al fondo di un 22 mm x 60 millimetri copertura in vetro antiscivolo con lato dorsale rivolto verso l&#39;alto (Figura 3F). Aggiungere qualche goccia di glicerolo al 50%, 0,25% KOH o 100% glicerolo in cima scheletro branchiale. Se la transizione dal 50% glicerolo, 0,25% KOH al 100% glicerolo, soluzione cambiamento in una provetta e agitare per&gt; 5 min prima del montaggio per equilibrare il tessuto. </li><li> Stendete due piccole sfere di plastilina e posto su una delle estremità del vetrino di agire come distanziatori. </li><li> Liberamente posizionare un secondo vetrino in cima con una pressione sufficiente per appiattire lo scheletro branchiale anteriore (Figura 3G). </li><li> Peel aprire lo sportello dorsale sinistra tra cui lastre dei denti dorsali, appiattire, e scorrevole tra le lamelle (Figura 3H). </li><li> Ripetere la tecnica con diritto lembo dorsale e spingere intero scheletro branchiale lontano dal bordo del vetrino (Figura 3I). <ol><li> alterarenativamente, tenere entrambi i lembi dorsali aperti con una pinza e con attenzione inserire il vetrino su, appiattendo lo scheletro branchiale in un unico movimento fluido. </li><li> In alternativa, montare lo scheletro branchiale a testa in giù su un vetrino, dispiegamento ogni lato dorsale lateralmente in modo gravità non permette lo scheletro branchiale per chiudere il backup. Poi coprire con il secondo 22 mm x 60 millimetri copertura in vetro antiscivolo e capovolgere prep. Nota: diverse tecniche di montaggio tendono a lavorare meglio o peggio per ogni individuo. Prova ogni e vedere che cosa si sente più comodo. </li></ol></li><li> Premere leggermente sulla parte superiore vetrino per appiattire le palle d&#39;argilla abbastanza per mantenere lo scheletro branchiale montato piatta, ma fare attenzione a non schiacciare il campione. <ol><li> Durante il processo di montaggio, ceratobranchials possono ruotare e oscurare una fila di rakers. Porre rimedio a questo da una pinza tra i coprioggetti e ri-orientare il ceratobranchials o l&#39;intero scheletro branchiale scorrevole. </li></ol></li><li> Conservare prepara piatto in vassoi di diapositive a rotemperatura om. Montato in 100% glicerolo, preparazioni possono essere memorizzati tra lamelle a ponte per almeno un decennio. pinze pulite e forbici con isopropanolo o etanolo e suggerimenti copertura. </li></ol><img alt="Figura 3" src="/files/ftp_upload/54056/54056fig3.jpg" /> Figura 3:. Piatto di montaggio branchiale scheletro di manipolazione e di montaggio dello scheletro branchiale è mostrato. Le frecce blu indicano la direzione del moto. (A) branchiale scheletro dorsale verso l&#39;alto. (B - D) Rotazione e incisione tra le piastre dorsali dente. (E) taglio laterale in tessuto molle per aprire ulteriormente la base del tubo intestinale. (F) branchiale scheletro posta sul fondo di un coprioggetto pronto per il montaggio. (G) Seconda coprioggetto posto sulla metà anteriore dello scheletro branchiale (sopra piastre dente dorsali). (H - I)montaggio piatto dello scheletro branchiale aprendo sportelli piastra dorsale denti e scorrevole tra due vetrini. Vedere i passaggi 4.1 tramite 4.6 per maggiori dettagli. Barra di scala = 5 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54056/54056fig3large.jpg" target="_blank">Clicca qui per vedere una versione più grande di questa figura.</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 branchial skeletal elements in a simple two-dimensional plane for ease of imaging and quantification of multiple traits. 
One limitation of this protocol is that although teeth, gill rakers, and most endochondral bones within this flat-mounted two-dimensional prep can be easily imaged and quantified, the second through fourth epibranchials, that have much more three-dimensional shapes, are not easily visualized and would require additional dissection to accurately measure. A second limitation of this protocol is the method is minimally destructive to the fish head. However, only two bones are damaged with a single cut each, the parasphenoid and the frontal bone. All other craniofacial skeletal elements can be returned to and measured at later points in time.
This protocol, although developed for sticklebacks, can be used in many other fish species, although sometimes modifications are necessary. For example, some fish species like zebrafish have a non-ossified dorsal chewing pad44 requiring the dorsal cut to be lateral, around this plate. The posterior end of the fifth ceratobranchial is curved is some species (e.g., zebrafish and other Cypriniformes) and must be trimmed or much larger clay spacers used to allow flat mounting. Particularly large specimens can be mounted between two pieces of glass or acrylic in the same manner. Species with a particularly thick epithelial layer covering tooth plates can be restained and cleared after dissection or more aggressively cleared with 2% KOH. Fish in this situation may also be presoaked in 1 or 2% KOH prior to Alizarin staining to increase dye penetrance. 
An alternative more technologically advanced method for imaging the branchial skeleton is Micro Computed Tomography (&#181;CT)23,45. However, &#181;CT requires an expensive scanner, is much more time consuming, and requires considerable training. Instead, this dissection protocol is inexpensive, quick, and amenable to high throughput studies required for quantitative studies such as quantitative trait loci (QTL) mapping genomic regions controlling evolved changes in craniofacial patterning. Additionally, these dissections can be performed at any larval or adult time point22,35,36,38 to study skeletal development. Although fish head anatomy is complex, this simple protocol allows easy visualization of the internal branchial skeleton, a critically important component of trophic morphology.

Una quantità incredibile di diversità esiste nello scheletro testa tra i vertebrati, soprattutto tra i pesci. In molti casi questa diversità facilita differenti strategie di alimentazione 1 - 4, e può comportare importanti modifiche sia modello craniofacciale esterna e interna. Lo scheletro branchiale è situato internamente nella gola di un pesce e circonda la maggior parte della cavità buccale. Lo scheletro branchiale comprende 5 segmenti omologhi serialmente, anteriore quattro delle quali supportano branchie. Insieme, queste cinque segmenti funzionano da interfaccia tra i pesci e il loro cibo 5. Variazione in una moltitudine di tratti, tra cui branchiospine, denti faringei, e le ossa branchiali contribuire a foraggiare efficace su diversi tipi di cibo. Spinarelli sono stati sottoposti a una radiazione adattativa dopo forme ancestrali oceaniche colonizzati laghi d&#39;acqua dolce e torrenti in tutto l&#39;emisfero settentrionale. Il cambiamento nella dietadal piccolo zooplancton nell&#39;oceano di grande preda in acqua dolce ha provocato drammatica variazione trofico in diversi tratti cranio-facciali 6. Mentre molti studi si sono concentrati sulle differenze cranio esterni in spinarelli 7 - 13, importanti cambiamenti cranio-facciali evolvono più volte nello scheletro branchiale interna. La possibilità di creare ibridi fecondi tra le popolazioni spinarello morfologicamente distinte offre un&#39;eccellente opportunità per mappare la base genetica delle modifiche si sono evoluti per lo scheletro branchiale. Una caratteristica trofico di importanza ecologica è il patterning di branchiospine, ossa dermiche periodici che costeggiano le facce anteriore e posteriore delle ossa branchiali e sono utilizzati per filtrare prede. I pesci che in genere si nutrono di piccoli oggetti preda tendono ad avere più a lungo e più densamente spaziati Gill rakers rispetto ai pesci che si nutrono sulla preda più grande 14,15. Variazione branchiospine stato segnalato sia wntro e tra le specie 14-19, e gli aspetti di Gill raker patterning contribuiscono a nicchie trofiche e fitness 16. Decenni di ricerche hanno ampiamente documentato numero Gill raker e variazione di lunghezza in spinarelli threespine 17 - 21; Tuttavia, questi studi in genere si concentrano sulla prima fila di branchiospine. Studi recenti hanno dimostrato modularità nel controllo genetico del numero branchiali raker attraverso lo scheletro branchiale 22,23 e attraverso una singola riga in branchiali raker spaziatura 23 e lunghezza 24 mettendo in evidenza l&#39;importanza di studiare più di remare uno o un singolo rastrello branchia per capire il sviluppo basi genetiche della riduzione Gill raker. Un secondo tratto trofico sia di importanza ecologica e biomedica è il patterning di denti faringei. Denti in pesci possono essere posizionati sia nella mascella orale e nello scheletro branchiale, noto come i denti faringei. denti orali sono utilizzati principalmente per prey catturare mentre denti faringei sono utilizzati per la masticazione e la preda manipolazione 25 - 27. Entrambi i gruppi si formano tramite meccanismi di sviluppo condivisi e sono considerati evolutivamente omologa 28. Modularità Interessante si verifica per cui alcune specie, come zebrafish, mancanza orale e della faringe dorsale denti 29, mentre le altre specie hanno più ceratobranchials dentate, pharyngobranchials, e talvolta dentate basihyal e hypobranchials 30. In spinarelli, denti faringei si trovano ventrale al quinto ceratobranchial e dorsalmente sul anteriore e posteriore pharyngobranchials 31. Cinematica di alimentazione stickleback mostrano la mascella orale viene utilizzato principalmente per la cattura la preda e facilitare l&#39;alimentazione di aspirazione 9 lasciando la masticazione alla mascella faringea. In ciclidi, inferiore faringeo mascella morfologia varia notevolmente 32,33 e ha dimostrato di essere adattivo e correlato con nicchia trofica 34. MultiPLE popolazioni stickleback d&#39;acqua dolce si sono evoluti gli aumenti drammatici nella ventrale faringea numero di denti 23,35,36. Studi recenti hanno dimostrato che la base genetica evolutiva di questo guadagno dente evoluto è in gran parte distinta in due popolazioni derivate indipendentemente spinarelli d&#39;acqua dolce 36. A differenza di denti di mammiferi, pesci rigenerare i denti continuamente per tutta la vita adulta 37. Entrambe queste popolazioni di acqua dolce dentate alti precedentemente descritti si sono evoluti un tasso di sostituzione dei denti accelerato, fornendo un sistema di vertebrati rara per studiare le basi genetiche della rigenerazione 36. Una terza caratteristica trofica che si è evoluta più volte in spinarelli d&#39;acqua dolce è più ossa epibranchial e ceratobranchial, le branchiali omologhi arco ribassato della mascella superiore e inferiore, rispettivamente 38. Più lunghe ossa branchiali conferiscono una grande cavità orale e probabilmente sono adattativi per consentire prede più grandi per essere Consumed. Inoltre, in altri pesci, ossa epibranchial sono importanti per la depressione delle lastre dei denti dorsale faringei 25. Come rastrelli branchiali e denti faringei, le ossa branchiali sono interni e, quindi, difficile da visualizzare o quantificare facilmente. Qui vi presentiamo un protocollo dettagliato per sezionare e TV a montare lo scheletro branchiale, consentendo una facile visualizzazione e la quantificazione di una varietà di importanti caratteristiche cranio-facciali. Mentre questo protocollo descrive una dissezione stickleback, questo stesso metodo funziona su una varietà di altri pesci.

<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> Questo protocollo si traduce in uno scheletro branchiale montato sezionato e piatta (Figura 4), ​​dove una serie di importanti caratteristiche trofiche può essere quantificato. Dal punto di vista dorsale, tutte le righe di branchiospine, tutte le lastre della faringe denti, e quasi tutte le ossa branchiali possono essere facilmente visualizzati e quantificati 22 - 24,35,36,38,42. Alizarina Rossa S reagisce anche su rodamina o filtro rosso simile permettendo doppia marcatura con altri marcatori (ad esempio, GFP transgenici 42) e un metodo alternativo di visualizzazione. Fluorescenza svanisce rapidamente in luce, in modo da conservare preparazioni al buio, se è prevista l&#39;imaging a fluorescenza o fenotipizzazione. Dal punto di vista ventrale, le branchie possono essere visualizzati e la loro pigmentazione quantificati 43. Preps possono essere memorizzati in 100% glicerolo per anni. 4.jpg &quot;/&gt; Figura 4:. Rappresentante spinarello scheletro branchiale Due esempi di scheletri branchiali macchiati e liquidati sono mostrati. (A) Brightfield immagine che mostra l&#39;osso etichetta rossa. (B) immagine fluorescente sotto un set di filtri rodamina B. Esempi di rastrelli, denti e le ossa sono etichettati con le parentesi acute, punte di freccia, e asterischi, rispettivamente. Barre di scala = 2 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54056/54056fig4large.jpg" target="_blank">Clicca qui per vedere una versione più grande di questa figura.</a>

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.

Dissezione e Flat-montaggio del Threespine Stickleback branchiale scheletro

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

Nanyang Technological University

Research

JoVE Journal

Biology

9.7K Views.  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. 

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

Dissection of Drosophila Ovaries

Dissezione di ovaie Drosophila

Ricerca

Biologia

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.

Drosophila larvali NMJ 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).

Dissezione lordo dello Stomaco del Gambero, 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.

Dissezione di giro dentato ippocampale di topo adulto

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.

Dissezione di organi dal Zebrafish adulti

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.

Dissezione e colorazione di Drosophila Larvali ovaie

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.

Dissezione del rene Zebrafish adulti

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.

Dissezione e Montaggio di<em&gt; Drosophila</em&gt; pupa Dischi Eye

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.

La dissezione del Auditory Bulla nei topi postnatale: Isolamento delle ossa dell&#39;orecchio medio e l&#39;analisi istologica

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.

Invertire la dissezione e dati DiceCT Reveal altrimenti nascosti nell'evoluzione del Primate viso

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