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.

Todo el trabajo de los peces fue aprobado por el Comité de Cuidado y Uso de Animales Institucional de la Universidad de California en Berkeley (número de protocolo R330). La eutanasia se realizó mediante inmersión en 0,025% tricaína-S tamponada con bicarbonato de sodio 0,1% 39. Todas las etapas se realizaron a temperatura ambiente. 1. Preparación Nota: Realice los pasos 1.1-1.5 en tubos cónicos o viales de centelleo que puede sellar herméticamente y mantenerse en posición horizontal. Los peces no deben ser agitadas constantemente, pero tratan de mezclar la solución tan a menudo como sea posible invirtiendo suavemente o se agita el bastidor de tubos o viales para exponer todos los lados de los peces a la solución de tinción y permitir que la mancha penetre en el tejido de manera uniforme. No coloque un gran lote de pescado en un agitador de plataforma, como el peso pesado del líquido romperá el agitador. <ol><li> Fijar bien el pescado recién sacrificados o pescado almacenado en etanol con 10% de formalina tamponada neutra (NBF) durante la noche. Como alternativa, utilice el 4% paraformaldehído en solución 1x PBS en lugar de 10% NBF. Nota: Si la extracción de ADN, clip de una pequeña porción de las aletas caudales o pectorales antes de la fijación y almacenar en etanol. </li><li> Disponer de corrección adecuadamente en una campana química y reemplazar con agua del grifo (es decir ~ pH 7,0) durante 2 horas. Evitar el uso de agua desionizada, ya que a menudo puede ser ácida y puede descalcificar el hueso. </li><li> Eliminar el agua y los peces mancha con 0,008% rojo de alizarina S en el 1% de KOH en agua durante 24 horas. Para los peces de menos de 20 mm de longitud estándar, utilice 0,004% rojo de alizarina S. (Haga una solución de reserva de 100x (0,8%) de rojo de alizarina S que a continuación se puede diluir). </li><li> Retirar la mancha (poner en un recipiente apropiado para desechos en la campana) y el lugar de peces en el agua del grifo para unos pocos horas. Cambiar el agua según sea necesario hasta que el agua de enjuague es mayormente despejado. </li><li> Eliminar el agua y el pescado en lugar de 50% de glicerol, 0,25% de KOH para la limpieza suave y posterior disección. Nota: Este protocolo de tinción es una modificación de los métodos descritos anteriormente 40,41. </li></ol><pclass = &quot;jove_title&quot;&gt; 2. Disección Nota: Véase la Figura 1 para una revisión de la morfología del esqueleto cabeza relevante. <img alt="Figura 1" src="/files/ftp_upload/54056/54056fig1.jpg" /> Figura 1:. Stickleback cabeza de la morfología del esqueleto rojo de alizarina manchada cabeza espinoso de tres espinas fotografiado con fluorescencia bajo un conjunto de filtro de rodamina B. la morfología de utilidad tiene la etiqueta: OP = opérculo, SubOp = subopérculo, BSR = branquiostegales rayos, Preop = preopérculo, Infraorb 1-3 = 1-3 infraorbitarios (también llamados circumorbitals o suborbitales), Dent = dentario, Premax = premaxilar, maxilar = Max , NAS = nasal, Lat. ETHM = etmoidal lateral, PSPH = parasphenoid, Fron = hueso frontal. Para una descripción más detallada del esqueleto de la cabeza del espinoso, véase Anker (1974) 31. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54056/54056fig1large.jpg" target="_blank">Haga clic aquí para ver una versión más grande de</a>esta figura. <ol><li> Coloque el pescado plano (Figura 2A) e insertar una pinza afilada # 5 de relojero en el lado del ojo en un ángulo de 45 ° ~ para perforar la membrana que cubre el ojo. </li><li> Pelar la membrana lejos del ojo, similar a la descamación una tapa yogur (Figura 2B). </li><li> Inserte fórceps abierto detrás del ojo, agarrar el nervio óptico detrás del ojo, y quitar el ojo (Figura 2C). No perforar el ojo, ya que se fuga de melanina. En caso de pinchazo, la melanina se puede lavar durante las etapas posteriores. </li><li> Repita del otro lado. </li><li> A partir de la parte posterior, coloque una pequeña cuchilla de disección tijera debajo de la solapa opérculo, la cuchilla de arrastre de tijera dorsal por encima de opérculo, luego se corta el tejido blando a través de la cuenca del ojo (Figura 2D). Cortar dorsal de la médula opérculo. </li><li> Cortar el hueso frontal (dorsal a la cuenca del ojo) (Figura 2E). </li><li> Cortar el hueso alrededor de la línea media parasphenoid tque el centro de las cuencas de los ojos (Figura 2F). </li><li> corte opérculo Repita en el lado opuesto. </li><li> Inserte fórceps bajo el opérculo y lentamente la cáscara de la cara lejos del cuerpo, el recorte de cualquier tejido blando todavía unido (Figura 2 G - H). Tenga cuidado de no perturbar la primera fila de hendiduras branquiales. <ol><li> Con unas pinzas, separar los ceratohyals en ambos lados de la basihyal línea media, mientras que pelar y retirar el esqueleto craneofacial anterior (toda la mandíbula incluyendo el dentario, premaxilar y maxilar; todo el esqueleto hioides incluyendo el opérculo dérmica exterior, preopérculo, subopérculo, y branquiostegales rayos y los dorsales y elementos que subyace endochondral ventral, y la parte anterior del cráneo, incluidos el nasal, etmoidal lateral, y los huesos infraorbitarios, véanse las figuras 1 y 2I). </li><li> espinas pélvicas se pueden plegar hacia fuera del cuerpo, y pueden servir como un asa para unas pinzas para agarrar of cuando está presente. Espinas encajen en su sitio. Para desbloquear, tirar suavemente de la columna vertebral con una pinza en dirección opuesta a cuerpo de los peces, y doblar suavemente en sentido posterior para presionar contra la columna vertebral plana peces. </li></ol></li><li> Insertar cerrado fórceps posterior y ventral al esqueleto branquial (justo debajo de tubo intestinal) y las pinzas de arrastre en sentido anterior, las burlas aparte los músculos y ligamentos que quedan unidos al esqueleto branquial (Figura 2J - K). </li><li> Usando la punta de unas pinzas cerradas, raspar los músculos que sujetan el dorsal esqueleto branquial a la caja craneana ventral en una posterior a la dirección anterior (Figura 2 l). </li><li> Repetir 2,9 y 2,10 en el lado opuesto. </li><li> Agarre la base del tubo intestinal y tire hacia delante para eliminar el esqueleto y el intestino branquial tubo (Figura 2 M - N). </li><li> Separar el tubo intestinal haciendo una posterior corte perpendicular al final de la quinta ceratobranquial (Figura 2O </strong&gt;). </li><li> Después de eliminar cualquier fragmento de hueso restante de la caja craneal en el lado dorsal del esqueleto branquial, insertar tijeras en la canasta branquial para hacer un corte dorsal (corte de anterior a posterior) entre los conjuntos bilaterales de placas de dientes dorsal (Figura 3A - D). corte se centra garantizar para evitar dañar las placas de los dientes dorsal. </li><li> Haga dos cortes laterales poco profundas en la luz del intestino de caucho en el extremo posterior del esqueleto branquial (extremo anterior del tubo intestinal) para ayudar en la apertura de la esqueleto branquial (Figura 3E). </li><li> Coloque el pescado y todas las piezas de tejido en un frasco y colocar el esqueleto branquial en un tubo de microcentrífuga con 50% de glicerol, 0,25% de KOH para continuar limpieza suave, o el 100% de glicerol, si se requiere ninguna otra compensación. frascos y tubos de etiquetas con un identificador único para que puedan ser rastreados. La cantidad de aclaramiento requerido es en gran parte una función del tamaño de los peces, peces grandes adulto (mayor de 40 mm de longitud estándar) requieren normalmente de compensación adicional. </li></ol><img alt="Figura 2" src="/files/ftp_upload/54056/54056fig2.jpg" /> Figura 2:. Stickleback disección esqueleto branquial manchado rojo de alizarina pez espinoso de tres espinas listo para la disección. El ojo está despigmentada de amplio legrado. Las flechas azules indican la dirección del movimiento. (A) Vista lateral de la cabeza del espinoso, es anterior a la derecha. (B) La eliminación de la membrana que cubre el ojo. (C) La eliminación del ojo. (D) Dorsal cortado por encima del opérculo. Corte del hueso (E) frontal. (F) Parasphenoid cortar. (G - I) La eliminación del esqueleto facial. (J) La eliminación de las conexiones de los tejidos blandos esqueleto branquial ventrales. (K - L) La eliminación de las conexiones esqueleto branquial dorsales. (&lt;strong&gt; M - N) La eliminación del esqueleto branquial. (O) Separar el tubo intestinal del esqueleto branquial. Consulte los pasos 2.1 a 2.16 para obtener más detalles. Barra de escala = 5 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54056/54056fig2large.jpg" target="_blank">Haga clic aquí para ver una versión más grande de esta figura.</a> 3. branquial Esqueleto Re-tinción (si es necesario) <ol><li> Para teñir el más oscuro esqueleto branquial o tejido claro más, retire 50% de glicerol, solución de KOH 0,25% y se lava con 1% KOH dos veces (un lavado de 5 minutos seguido de un segundo lavado 24 hr mientras se agita horizontalmente en un agitador de plataforma). </li><li> Retire 1% KOH y re-mancha con 0.008% rojo de alizarina S en 1% de KOH durante 24 h. </li><li> Retire las manchas y reemplazar con 1% de KOH durante 24 horas. </li><li> Retire la solución de KOH y reemplazar con 50% de glicerol, 0,25% de KOH. </li></ol> 4. Montaje branquial Esqueleto <ol><li> Retire esqueleto branquialde 50% de glicerol, 0,25% de KOH o 100% de glicerol y el lugar cerca de la parte inferior de un 22 mm x 60 mm cubreobjetos de vidrio con el lado dorsal hacia arriba (Figura 3F). Añadir unas gotas de glicerina al 50%, 0,25% KOH o 100% de glicerol en la parte superior del esqueleto branquial. Si la transición de 50% de glicerol, 0,25% de KOH al 100% de glicerol, solución de cambio en un tubo de microcentrífuga y agitar durante&gt; 5 min antes de montar para equilibrar los tejidos. </li><li> Estirar dos pequeñas bolas de arcilla de modelar y colocar en cada extremo de la hoja de la cubierta para actuar como espaciadores. </li><li> Sin apretar colocar un segundo cubreobjetos en la parte superior con la presión suficiente para aplanar el esqueleto branquial anterior (Figura 3G). </li><li> Abra retirando la tapa dorsal izquierda que incluye placas de dientes dorsales, aplanar, y se deslizan entre los cubreobjetos (Figura 3H). </li><li> Repita técnica con la aleta dorsal derecha y empujar toda esqueleto branquial de distancia desde el borde del cubreobjetos (Figura 3I). <ol><li> Alterarde forma nativa, mantenga ambas aletas dorsales abiertas con unas pinzas y colocar con cuidado el cubreobjetos sobre, aplanando el esqueleto branquial con un movimiento suave. </li><li> Como alternativa, montar el esqueleto branquial al revés sobre una hoja de la cubierta, extendiendo cada lado dorsal lateralmente para que la gravedad no permite que el esqueleto branquial para cerrar una copia de seguridad. A continuación, cubrir con la segunda x 60 mm cubreobjetos de vidrio de 22 mm e invierta preparación. Nota: Las diferentes técnicas de montaje tienden a funcionar mejor o peor para cada individuo. Pruebe cada uno y ver lo que se siente más cómodo. </li></ol></li><li> Presione ligeramente en la parte superior cubreobjetos para aplanar las bolas de arcilla suficientes para mantener el esqueleto branquial montado plana, pero tenga cuidado de no aplastar el espécimen. <ol><li> Durante el proceso de montaje, ceratobranchials pueden rotar y oscurecer una fila de rastrillos. Remediar esto mediante una pinza entre el cubreobjetos y reorientando los ceratobranchials o de todo el esqueleto branquial deslizante. </li></ol></li><li> Tienda prepara plana en bandejas deslizantes en rotemperatura om. Montado en el 100% de glicerol, preparaciones se pueden almacenar entre cubreobjetos puenteados durante al menos una década. pinzas limpias y tijeras con isopropanol o etanol y consejos de cubierta. </li></ol><img alt="figura 3" src="/files/ftp_upload/54056/54056fig3.jpg" /> Se muestra de montaje plana del esqueleto branquial manipulación y el montaje del esqueleto branquial: Figura 3.. Las flechas azules indican la dirección del movimiento. (A) branquial dorsal esqueleto hacia arriba. (B - D) La rotación y la incisión entre las placas dentales dorsal. (E) de corte lateral en el tejido blando para abrir aún más la base del tubo intestinal. (F) branquial esqueleto colocado en la parte inferior de un cubreobjetos listo para el montaje. (G) En segundo cubreobjetos colocado en la mitad anterior del esqueleto branquial (por encima de las placas dorsales de los dientes). (H - I)montaje plano del esqueleto branquial mediante la apertura de las aletas de la placa de los dientes dorsales y deslizamiento entre dos hojas de la cubierta. Consulte los pasos 4.1 a 4.6 para más detalles. Barra de escala = 5 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54056/54056fig3large.jpg" target="_blank">Haga clic aquí para ver una versión más grande de esta 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 cantidad increíble de la diversidad existente en el esqueleto de la cabeza entre los vertebrados, en especial entre los peces. En muchos casos esta diversidad facilita diferentes estrategias de alimentación 1-4, y puede implicar cambios importantes en tanto patrón craneofacial externa e interna. El esqueleto branquial está situado internamente en la garganta de un pez y rodea la mayor parte de la cavidad bucal. El esqueleto branquial se compone de 5 segmentos serie homóloga, la parte anterior de cuatro de los cuales apoyan las branquias. En conjunto, estos cinco segmentos funcionan como una interfaz entre el pescado y sus alimentos 5. La variación en una multitud de rasgos incluyendo branquiespinas, dientes faríngeos, y los huesos branquiales contribuyen a forrajeo eficiente en diferentes tipos de alimentos. Espinosos han sido sometidos a una radiación adaptativa después de formas ancestrales oceánicas colonizados lagos de agua dulce y arroyos en todo el hemisferio norte. El cambio en la dietadel zooplancton pequeño en el océano para una presa más grande en el agua dulce se ha traducido en la variación trófica dramático en varios rasgos craneofaciales 6. Mientras que muchos estudios se han centrado en las diferencias craneofaciales externos en los espinosos 7 - 13, los cambios craneofaciales importantes evolucionan en repetidas ocasiones en el esqueleto branquial interno. La capacidad de crear híbridos fértiles entre poblaciones de espinosos morfológicamente distintos proporciona una excelente oportunidad para mapear la base genética de los cambios evolucionado hasta el esqueleto branquial. Un rasgo trófica de importancia ecológica es el patrón de hendiduras branquiales, los huesos dérmicos periódicas que recubren las caras anterior y posterior de los huesos branquiales y se utilizan para filtrar los elementos de presa. Los peces que normalmente se alimentan de pequeñas presas tienden a tener más largo y más densamente espaciados Gill RäKERS en comparación con los peces que se alimentan de presas más grandes 14,15. La variación en branquiespinas ha informado tanto wentro de las especies y entre 14-19, y los aspectos de enmalle de profundidad de corte de patrones contribuyen a nichos tróficos y gimnasio 16. Décadas de investigación han documentado ampliamente el número de enmalle de profundidad de corte y la variación de la longitud de los peces espinosos de tres espinas 17 - 21; Sin embargo, estos estudios suelen centrarse en la primera fila de hendiduras branquiales. Trabajos recientes han demostrado la modularidad en el control genético del número de enmalle de profundidad de corte a través del esqueleto branquial 22,23 ya través de una sola fila en la profundidad de corte de enmalle espaciamiento 23 y 24 de longitud que destaca la importancia de estudiar más de la fila uno o un solo diente de ataque de enmalle para entender la base genética del desarrollo de la reducción de enmalle de profundidad de corte. Un segundo rasgo trófica de tanto significado ecológico y biomédica es el patrón de los dientes faríngeos. Los dientes en los peces pueden estar situados tanto en la mandíbula oral y en el esqueleto branquial, conocido como dientes faríngeos. dientes orales se utilizan principalmente para pRey capturar mientras que los dientes faríngeos se utilizan para la masticación y la presa de manipulación 25 - 27. Ambos conjuntos se forman a través de los mecanismos de desarrollo compartidos y se consideran homólogas de desarrollo 28. Modularidad interesante se produce mediante el cual algunas especies, como el pez cebra, la falta de dientes de la faringe oral y dorsal 29, mientras que otras especies tienen múltiples ceratobranchials dentadas, pharyngobranchials y, a veces dentadas basihyal y hypobranchials 30. En los espinosos, dientes faríngeos se encuentran ventralmente en la quinta ceratobranquial y dorsalmente en la parte anterior y posterior pharyngobranchials 31. Cinemática sobre la alimentación del pez espinoso muestran la mandíbula oral se utiliza principalmente para la captura de presas y facilitar la alimentación de aspiración 9 dejando la masticación de la mandíbula faríngea. En cíclidos, menor morfología de la mandíbula faríngea varía dramáticamente 32,33 y se ha demostrado que ser adaptable y se correlacionó con nicho trófico 34. Multipoblaciones de espinosos de agua dulce ples han evolucionado aumentos espectaculares en la faringe ventral número de dientes 23,35,36. Trabajos recientes han demostrado que la base genética del desarrollo de esta ganancia diente evolucionado en gran medida es distinta en dos poblaciones derivadas independientemente de espinosos de agua dulce 36. A diferencia de los dientes de mamíferos, peces regeneran sus dientes de forma continua durante la vida adulta 37. Ambas poblaciones de agua dulce dentadas altas anteriormente descritos han evolucionado una tasa de reemplazo de dientes acelerado, proporcionando un sistema de vertebrados rara para estudiar la base genética de regeneración 36. Un tercer rasgo trófico que se ha desarrollado en varias ocasiones en los espinosos de agua dulce es más largo y huesos epibranchial ceratobranquiales, los homólogos de arco rebajado branquiales de la mandíbula superior e inferior, respectivamente 38. Más largos huesos branquiales confieren una cavidad bucal grande y probablemente son adaptativos para permitir que las presas más grandes para ser Consumed. Por otra parte, en otro pescado, huesos epibranchial son importantes para la depresión de las placas de dientes faríngeos dorsal 25. Al igual que branquiespinas y dientes faríngeos, los huesos branquiales son internos y, por tanto, difíciles de visualizar con facilidad o cuantificar. Aquí se presenta un protocolo detallado para diseccionar y de montaje en el esqueleto branquial plana, lo que permite una fácil visualización y cuantificación de una variedad de importantes rasgos craneofaciales. Si bien este protocolo describe una disección stickleback, este mismo método funciona en una variedad de otros peces.

<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> Este protocolo da como resultado un esqueleto branquial montada diseccionado y plana (Figura 4), ​​donde una variedad de importantes rasgos tróficos se puede cuantificar. Desde una vista dorsal, todas las filas de hendiduras branquiales, todas las placas de dientes faríngeos, y casi todos los huesos branquiales pueden ser fácilmente visualizados y cuantificaron 22 - 24,35,36,38,42. Rojo de alizarina S también emite fluorescencia en una rodamina o filtro rojo similares permitiendo el doble etiquetado con otros marcadores (por ejemplo, GFP transgénica 42) y un método alternativo de visualización. La fluorescencia se desvanece rápidamente a la luz, por lo que las tiendas preparaciones en la oscuridad si se planea imágenes fluorescentes o el fenotipo. Desde un punto de vista ventral, las branquias se pueden visualizar y cuantificar su pigmentación 43. Preps se pueden almacenar en 100% de glicerol durante años. 4.jpg &quot;/&gt; Figura 4:. Stickleback Representante esqueleto branquial Dos ejemplos de esqueletos branquiales manchadas y despejados se muestran. (A) Campo claro imagen que muestra el hueso etiqueta roja. (B) Imagen fluorescente bajo un conjunto de filtro de rodamina B. Ejemplos de rastrillos, los dientes y los huesos están etiquetados con símbolos de intercalación, puntas de flecha, y asteriscos, respectivamente. Las barras de escala = 2 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54056/54056fig4large.jpg" target="_blank">Haga clic aquí para ver una versión más grande de esta 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.

La disección y-montaje plana del esqueleto branquial de tres espinas Stickleback

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 vistas.  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

La disección de los ovarios Drosophila

Investigación

Biología

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 larval NMJ Disección

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).

La disección bruto del estómago de la langosta, 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.

La disección de Gyrus dentado del hipocampo de ratones adultos

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.

La disección de los órganos del pez cebra adultos

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.

La disección y la tinción de Drosophila Larvas Ovarios

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.

La disección del riñón adultos de pez cebra

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.

Disección y montaje de<em&gt; Drosophila</em&gt; Pupal discos 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 disección de la Auditivo Bulla en ratones postnatales: Aislamiento de los huesos del oído medio y el análisis histológico

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.

Inversa, disección y DiceCT revelan lo contrario oculta datos en la evolución de la cara de Primate

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