<p class="jove_content">Le projet actuel a été financé par la preuve de l’Université du Kansas de fonds Concept. Nous tenons à remercier le personnel soignant à KUMC (Kansas City, KS) pour nous aider à obtenir des humains des cordons ombilicaux et David Jorgensen pour aider avec les cultures de limaçon.</p>

<p class="jove_content">Toutes les procédures, y compris l’euthanasie animale, ont été effectuées selon le protocole institutionnel animalier et utilisation Comité (IACUC) approuvé (APUC #2014-2234) à l’Université du Kansas Medical Center (KUMC).</p>
<p class="jove_content">Remarque : HWJCs ont été isolées de cordons ombilicaux humains qui ont été donnés par les patients qui a donné son consentement éclairé et spécimens ont été utilisés conformément aux protocoles approuvés par le Comité de sujets humains Université du Kansas (KU-CISR #15402).</p>
<p class="jove_title">1. l’OS Temporal récolte et l’isolement de la cochlée</p>
<ol>
	<li>Décapiter une souris convenablement euthanasiée, 15 semaines de coupe entre le crâne et la première vertèbre cervicale avec des ciseaux chirurgicaux aiguisés, puis vaporiser vers le bas de la tête avec l’éthanol à 70 % pour désinfecter (<strong class="xfig">Figure 1 a</strong>).</li>
	<li>Coupent le crâne dans un plan longitudinal à l’aide de la même paire forte de ciseaux chirurgicaux (<strong class="xfig">Figure 1 b</strong><strong>-C</strong>, lignes pointillées).</li>
	<li>Retirez le tissu cérébral du crâne à l’aide d’une paire de pinces (<strong class="xfig">Figure 1</strong>, flèche).</li>
	<li>Identifier l’OS temporal à travers la présence de l’auditif et les racines de nerf vestibulaire (<strong class="xfig">Figure 1E</strong><strong>-F</strong>, pointes de flèche) à l’intérieur du crâne et le conduit auditif de l’extérieur (<strong class="xfig">Figure 1F</strong>, flèche). Découpez soigneusement à travers le crâne pour isoler l’OS temporal du reste du crâne (<strong class="xfig">Figure 1</strong>).<br>Remarque : Dans certains cas, il peut être possible de dégager délicatement l’OS environnants du crâne de l’OS temporal sans nécessiter de coupe.</li>
	<li>Insérer la pince fine dans l’ouverture du conduit auditif environ 5 mm (<strong class="xfig">Figure 1 H</strong>), donc les conseils ne risquent pas de percer la cochlée. Casser délicatement l’os de la bulle tympanique (<strong class="xfig">Figure 1 H</strong>, haut rouge zone ombrée) donc il fractures (ligne<strong class="xfig">Figure 1I</strong>, rouge pointillée).<br>Remarque : Si nécessaire, un microscope à dissection peut aider dans ce processus afin d’assurer un positionnement précis et la manipulation des instruments.</li>
	<li>À l’aide de pinces fines, dégager loin le reste de la bulle tympanique de l’OS temporal, exposant le labyrinthe osseux de la cochlée (<strong class="xfig">Figure 1J</strong>, supports de rouges).</li>
	<li>Placez une boîte de Pétri assez grande pour contenir de l’OS temporal (<em>p. ex.</em>, 30 mm ou plus) sous la dissection étendue de champ et remplissez-le avec assez Phosphate Buffered Saline (PBS) pour couvrir l’OS temporal (<em>par exemple</em>, 0,5 à 1 mL).</li>
	<li>Plonger l’OS temporal avec la bulle tympanique supprimée dans le PBS avec les fenêtres ovales et rondes de la cochlée vers le haut.<br>Remarque : Suppression du système vestibulaire de la cochlée n’est pas nécessaire, car le système vestibulaire sert bien une poignée de manipulation et d’orienter les caractéristiques de la cochlée (<strong class="xfig">Figure 1I</strong> et <strong class="xfig">Figure 2 b</strong>).</li>
	<li>Avec un ultra fine paire de pinces, retirer l’artère stapédiens, qui traverse l’étrier.</li>
	<li>Insérez une extrémité de la pince ultra fine à travers l’arche de l’étrier où l’artère est transmise et soulever délicatement l’étrier. L’étrier désarticulés devrait soulever hors de la fenêtre ovale.</li>
	<li>À l’aide d’une pointe de la pince ultra-fine, perforer l’ovale et ronde de windows.</li>
	<li>À l’aide d’une solution de PBS 1 x rempli de 28,5 calibre seringue avec tuyau (diamètre intérieur 0,28 mm, diamètre extérieur 0,61 mm) attaché, positionner le tube sur la fenêtre ovale (<strong class="xfig">Figure 2 a</strong> <strong> -B</strong>).<br>Remarque : L’ouverture du tube peut être manipulé à l’aide de la pince ultra-fine contribuant à assurer un positionnement précis.<ol>
		<li>Utiliser une seringue fraîche et les tubes pour chaque cochlée.</li>
	</ol>
	</li>
	<li>Perfuse 2 mL d’antibiotique-antimycosiques (anti-anti) 10 % et 10 % la pénicilline-streptomycine (stylo-strep) dans du PBS par le biais de la cochlée pendant 5 min pour enlever la périlymphe. Perfusant trop vite avec trop de pression peut endommager les structures fines dans la cochlée.<br>Remarque : Conduire manuellement la seringue à la main pour perfuse le liquide dans la cochlée est efficace, bien qu’une pompe à perfusion pourrait être utilisée comme une alternative.<ol>
		<li>Si perfusant manuellement, utilisez une paire de fines, fermeture automatique des pinces pour stabiliser la cochlée (<strong class="xfig">Figure 2 a</strong>) au cours de la perfusion en saisissant la partie vestibulaire de l’OS temporal.<br>Remarque : Bien que non obligatoire, ce qui laisse les deux mains libres pour manipuler les instruments ; une main peut conduire à la seringue (<strong class="xfig">Figure 2 a</strong>) tandis que l’autre permet de positionner le tube de façon appropriée (<strong class="xfig">Figure 2 b</strong>).</li>
		<li>De cette étape vers l’avant, prenez soin de ne pas exposer la cochlée inutilement à l’air. Introduire des bulles d’air dans les espaces fluides se bloque la circulation du fluide, et les sécher n’endommage les structures fines dans la cochlée.</li>
	</ol>
	</li>
	<li>Retirez et jetez n’importe quel muscle tissu et l’OS fragments restants à l’aide de pinces fines.</li>
	<li>Si nécessaire, stocker limaçon isolé à 4 ° C en solution de strep-stylo de 10 % dans du PBS et anti-anti 10 % jusqu'à sept jours, avec des changements réguliers de solution antibiotique toutes les 48 h.</li>
</ol>
<p class="jove_title">2. cochléaire traitement</p>
<ol>
	<li><strong>DÉCELLULARISATION</strong>
	<ol>
		<li>Pour chaque cochlée, remplir un flacon à scintillation de verre de 20 mL avec un 1 % sodium dodécyl sulfate solution (SDS) dans de l’eau déminéralisée (DI), qui sert à decellularize la cochlée.</li>
		<li>Couper l’extrémité hors d’une pipette de transfert 7 mL en plastique à l’aide d’une lame de rasoir afin que l’ouverture soit assez grande pour une cochlée de passer à travers.</li>
		<li>À l’aide de la pipette de transfert, doucement dresser la cochlée dans la pipette alors il passe juste au bord de la coupure de quelques millimètres.</li>
		<li>Expulser la cochlée dans le flacon 1 % de scintillation rempli de solution SDS.</li>
		<li>Faire circuler 2 mL de SDS de 1 % dans de l’eau déminéralisée (DI) par le biais de la cochlée dans le flacon à scintillation à l’aide d’une pipette de transfert.</li>
		<li>Placer le flacon à scintillation dans une coiffe et laisser le flacon à scintillation de tourner à 10 tr/min pendant 72 h à température ambiante.<ol>
			<li>Changez 1 % SDS solution toutes les 24 h sur une période de 72 h. Faites attention lors du changement de SDS pour ne pas exposer la cochlée dans l’atmosphère.</li>
		</ol>
		</li>
		<li>Laver la cochlée trois fois de 30 min chacun, avec l’eau distillée.</li></ol></li></ol>Effectuer les lavages dans le même flacon à scintillation, utilisé pour des charges de la SDD ; à ce stade, la cochlée est DECELLULARISE.
	
	
	<li><strong>Décalcification</strong>
	<ol>
		<li>Retirez l’eau DI le flacon à scintillation, laissant juste assez pour garder la cochlée submergés.</li>
		<li>Pour commencer une décalcification, ajouter 5 mL de 10 % d’acide tétraacétique (EDTA) (0,02 M) dans l’eau distillée dans le flacon à scintillation contenant de la cochlée.</li>
		<li>Faire circuler 2 mL d’EDTA 10 % dans l’eau distillée par le biais de la cochlée dans le flacon à scintillation à l’aide d’une pipette de transfert.</li>
		<li>Remettre le flacon à scintillation dans la coiffe et laisser le flacon à scintillation de tourner à 10 tr/min pendant 72 h à température ambiante. Changer la solution d’EDTA 10 % toutes les 24 h sur une période de 72 h. Faites attention lors du changement des solutions afin que la cochlée n’est pas exposée à l’air.</li>
		<li>Rincer la cochlée 3 fois pendant 2 h chacune avec du PBS 1 x ; à ce stade, la cochlée est détartrée.</li>
	</ol>
	</li>
	<li><strong>Stockage</strong>
	<ol>
		<li>Stocker le limaçon pendant 72 h dans une anti-anti, 10 % plume strep-solution à 10 % dans du PBS à 4 ° C.<br>Remarque : Les plus longues périodes de stockage peuvent être possibles avec des changements réguliers de solution antibiotique ; Toutefois, un stockage prolongé de limaçon transformés n’a pas été validé dans le présent protocole.</li>
	</ol>
	</li>

<p class="jove_title">3. achats et l’Expansion des hWJCs</p>
<ol>
	<li>Isoler les hWJCs selon les protocoles publiés antérieurement<sup class="xref">22</sup>. En bref, les cordons ombilicaux segment en sections de 3 cm.</li>
	<li>Faites une incision peu profonde avec un scalpel dans la longueur sur chaque segment du cordon ombilical de dérouler et d’exposer la gelée de Wharton du cordon ombilical. Forceps permet de supprimer les vaisseaux sanguins.</li>
	<li>Laver les segments de cordon ombilical deux fois avec assez de PBS (<em>p. ex.</em>, 40 mL dans une boîte de Pétri de 60 mm) pour submerger le tissu. Finement hacher des segments de tissus avec un scalpel. Digérer le tissu dans une boîte de Pétri avec 50 mL de milieu de digestion de 100 mm (collagénase de type 2 de 0,2 %, 1 % la pénicilline-streptomycine, en basse-glucose moyen (DMEM d’aigle modifié de Dulbecco)).</li>
	<li>Placer le tissu en digérant moyen dans un agitateur orbital à 50 tr/min pendant la nuit dans un 5 % CO<sub>2</sub>, incubateur de 37 ° C. Le lendemain, diluer milieu de digestion à un ratio de 01:16 dans 2 % antibiotique-antimycosiques dans du PBS et cellules de granule par centrifugation à 500 g à la température ambiante (~ 27 ° C) pour 10 min. jetez surnageant.</li>
	<li>Remettre en suspension les boulettes dans le cellules souches mésenchymateuses croissance moyen (MSCGM) et recombiner des cellules. Cellules de la plaque à une densité de 7 x 10<sup>3</sup> cellules/cm<sup>2 </sup>en culture de tissus traitement flacons T-75. HWJCs en MSCGM de la culture et étendre au passage 5 pour des expériences.<br>Remarque : HWJCs peut être Sub cultivées dans plus grands T-fioles (<em>p. ex.</em>, T-150, T-300) lors de passage pour augmenter le rendement des hWJCs pour des expériences.</li>
</ol>
<p class="jove_title">4. l’injection de hWJCs dans Decellularized limaçon</p>
<ol>
	<li><strong>Préparer le limaçon DECELLULARISE</strong>
	<ol>
		<li>À l’aide d’une technique aseptique, laver limaçon stockées dans du PBS 1 x trois fois pour chaque 30 min.</li>
		<li>Transférer le limaçon dans puits séparés d’une plaque 24 puits et ajouter 1 mL de 37 ° C préchauffé MSCGM.</li>
		<li>Incuber le limaçon dans un 5 % CO<sub>2</sub>, 37 ° C incubateur de culture cellulaire pendant au moins 1 h avant l’injection de cellules.</li>
	</ol>
	</li>
	<li><strong>Dissocier hWJCs et remettre en suspension</strong>
	<ol>
		<li>HWJCs de lavage avec 37 ° C préchauffé solution 1 PBS x deux fois.</li>
		<li>Ajouter suffisamment 37 ° C préchauffé trypsine avec EDTA 0,5 % pour couvrir la surface du navire cellulaire.</li>
		<li>Incuber les hWJCs jusqu'à 5 min dans un 5 % CO<sub>2</sub>, 37 ° C incubateur de culture cellulaire.<ol>
			<li>Vérifiez que 90 % des cellules ont détaché de la surface de culture cellulaire en affichant des cellules sous un microscope inversé.<br>Remarque : Les cellules qui se déplacent lorsque le récipient de culture est tapoté doucement, ont détaché. Les cellules qui restent fixes, n’ont pas détaché.</li>
			<li>Tapoter légèrement les côtés du ballon pour déloger davantage les cellules attachées.</li>
		</ol>
		</li>
		<li>À l’aide d’une technique aseptique, transfert hWJCs du récipient de culture dans un tube conique de 50 mL contenant un volume équivalent de MSCGM pour le volume de la trypsine permettant de dissocier les cellules.</li>
		<li>Le hWJCs de granule par centrifugation à 500 g à la température ambiante (~ 27 ° C) pendant 5 min.</li>
		<li>Aspirer le surnageant et remettre en suspension les hWJCs en MSCGM à une concentration de 500 000 cellules/mL.</li>
	</ol>
	</li>
	<li><strong>Perfuse limaçon</strong>
	<ol>
		<li>Limaçon de transfert à une nouvelle plaque 24 puits à l’aide d’une pipette de transfert.</li>
		<li>Ajouter une goutte de MSCGM à chaque cochlée afin d’empêcher toute cochlée ne se dessèchent pas.</li>
		<li>Délicatement, orienter la cochlée avec une pincette ultra fines pour que la fenêtre ovale et ronde pointent vers le haut.<br>Remarque : Faire extrêmement attention lorsque vous manipulez directement chaque cochlée comme la cochlée est facilement endommagée.</li>
		<li>À l’aide d’une seringue stérile calibre 28,5 insuline avec tuyau connecté, établir 0,2 mL resuspendue hWJCs (100 000 cellules).</li>
		<li>À l’aide de pinces fines stérilisé, positionner le tube au-dessus de la fenêtre ovale.</li>
		<li>Lentement et délicatement perfuse la cochlée avec 0,2 mL de hWJCs remises en suspension à l’aide d’une seringue à insuline 28,5-Manomètre reliée au tuyau de polyéthylène (diamètre extérieur : 0,61 mm, diamètre intérieur : 0,28 mm) pendant environ 5 min.</li>
		<li>Après la perfusion, ajouter 0,8 mL de 37 ° C préchauffé MSCGM dans la cupule contenant la cochlée pour porter le volume total dans le puits à 1 mL.</li>
		<li>Répétez les étapes 4.3.3-4.3.7 pour chaque cochlée, à l’aide d’une seringue fraîche et tube à chaque fois.</li>
		<li>Place perfusés limaçon dans un 5 % de CO<sub>2</sub>et 37 ° C incubateur de culture cellulaire changement médias trois fois par semaine.</li>
	</ol>
	</li>
</ol>
<p class="jove_title">5. préservation et la récolte de la cochlée</p>
<p class="jove_content">NOTE : Limaçon peut-être être cultivés et récoltés à n’importe quel point du temps de perfusion après de jusqu'à 30 jours.</p>
<ol>
	<li>Afin de préserver le limaçon, difficulté à la paraformaldéhyde à 4 % dans du PBS 1 x nuit à 4 ° C sur une plate-forme bascule.</li>
	<li>Après fixation, laver le limaçon avec du PBS 1 x trois fois de 5 min chacun.</li>
	<li>Peu à peu déshydrater la cochlée avec de l’éthanol et clair avec un solvant hydrocarbure aliphatique avant l’intégration à la paraffine.</li>
	<li>Section des échantillons jusqu'à une épaisseur de 10 µm à l’aide d’un microtome et monter sur lames de microscope de verre.<br>Remarque : Les échantillons sont maintenant prêts à histologique ou immunohistochimique de traitement à l’aide de protocoles standard.</li>
</ol>

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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=Improving+Viability+and+Transfection+Efficiency+with+Human+Umbilical+Cord+Wharton's+Jelly+Cells+Through+Use+of+a+ROCK+Inhibitor.">Improving Viability and Transfection Efficiency with Human Umbilical Cord Wharton's Jelly Cells Through Use of a ROCK Inhibitor.</a> <em style='font-style: italic'>Cell Reprogram</em>. , (2014).</li><li>Mellott, A. J., Shinogle, H. E., Moore, D. S., Detamore, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+Photo-conversion:+A+second+chance+to+label+unique+cells.">Fluorescent Photo-conversion: A second chance to label unique cells.</a> <em style='font-style: italic'>Cell Mol Bioeng</em>. <b>8</b> (1), 187-196 (2015).</li><li>Mitchell, K. 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=Matrix+cells+from+Wharton's+jelly+form+neurons+and+glia.">Matrix cells from Wharton's jelly form neurons and glia.</a> <em style='font-style: italic'>Stem Cells</em>. <b>21</b> (1), 50-60 (2003).</li><li>Mellott, A. 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=Nonviral+Reprogramming+of+Human+Wharton's+Jelly+Cells+Reveals+Differences+Between+ATOH1+Homologues.">Nonviral Reprogramming of Human Wharton's Jelly Cells Reveals Differences Between ATOH1 Homologues.</a> <em style='font-style: italic'>Tissue Eng Part A</em>. <b>21</b> (11-12), 1795-1809 (2015).</li><li>Buytaert, J. A., Johnson, S. B., Dierick, M., Salih, W. H., Santi, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MicroCT+versus+sTSLIM+3D+imaging+of+the+mouse+cochlea.">MicroCT versus sTSLIM 3D imaging of the mouse cochlea.</a> <em style='font-style: italic'>J Histochem Cytochem</em>. <b>61</b> (5), 382-395 (2013).</li><li>Nonoyama, H., 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=Investigation+of+the+ototoxicity+of+gadoteridol+(ProHance)+and+gadodiamide+(Omniscan)+in+mice.">Investigation of the ototoxicity of gadoteridol (ProHance) and gadodiamide (Omniscan) in mice.</a> <em style='font-style: italic'>Acta Otolaryngol</em>. <b>136</b> (11), 1091-1096 (2016).</li><li>Dai, C., 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=Rhesus+Cochlear+and+Vestibular+Functions+Are+Preserved+After+Inner+Ear+Injection+of+Saline+Volume+Sufficient+for+Gene+Therapy+Delivery.">Rhesus Cochlear and Vestibular Functions Are Preserved After Inner Ear Injection of Saline Volume Sufficient for Gene Therapy Delivery.</a> <em style='font-style: italic'>J Assoc Res Otolaryngol</em>. , (2017).</li><li>Sutherland, A. J., Converse, G. L., Hopkins, R. A., Detamore, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+bioactivity+of+cartilage+extracellular+matrix+in+articular+cartilage+regeneration.">The bioactivity of cartilage extracellular matrix in articular cartilage regeneration.</a> <em style='font-style: italic'>Adv Healthc Mater</em>. <b>4</b> (1), 29-39 (2015).</li></ol>

<p class="jove_content">Les auteurs n’ont rien à divulguer.</p>

<p class="jove_content">Nous avons démontré avec succès que des cellules cochléaires natifs peuvent être retirés de la cochlée via un processus DÉCELLULARISATION, qui permet l’utilisation de la cochlée comme un échafaudage de tissu complexe, en trois dimensions. Santi <em>et al</em>. <sup class="xref">15</sup> mis au point la méthode initiale pour decellularizing limaçon et ont réussi à estimer les volumes de nombreuses structures cochléaires grâce à l’aide d’une nappe de lumière microscopie<sup class="xref">23</sup>. Ce premiers travaux a servi une base solide pour l’ingénierie tissulaire et des techniques de culture cellulaire présentés ici. Cochlée DECELLULARISE peut être avec succès infusée avec des cellules et puis cultivés pendant une période prolongée de temps. Limaçon d’une vaste gamme d’animaux pourrait théoriquement être utilisés en combinaison avec une même gamme de types de cellules. Ces combinaisons permettent les techniques présentées ici pour être utilisé dans nombreuses conceptions expérimentales possibles, telles que les applications qui examinent l’épithélium sensoriel développement<sup class="xref">14</sup>, drug testing de nouveaux agents pharmaceutiques<sup class="xref"> en génie tissulaire 24</sup>, et élaboration de réparation cochléaire modèles<sup class="xref">25</sup>. Cependant, malgré la large applicabilité de ces techniques ils ne sont pas sans limites.</p>
<p class="jove_content">Une des limites de la technique actuelle est que la perfusion est variable et basé sur l’individu. Développer un titulaire, de la cochlée et à l’aide d’un pousse-seringue pour mener les perfusions peuvent augmenter la précision et la réussite du processus. En outre, le prouver définitivement DÉCELLULARISATION réussie pourrait potentiellement être accompli via plusieurs méthodes. Nous avons utilisé la coloration DAPI qui a été utilisé pour quantifier les noyaux des cellules restantes, et nous avons n’observé aucun post-DÉCELLULARISATION noyaux résiduels. En outre, des essais de quantification ADN standards utilisent le réactif, colorant vert, détecter des quantités picogramme de l’ADN, pour identifier les restes d’acide nucléique, qui est peut-être visuellement plus sensibles que le DAPI pour identifier la présence de petits fragments d’acides nucléiques acides dans les tissus DECELLULARISE<sup class="xref">26</sup>. Peu importe quelles techniques sont utilisées pour valider DÉCELLULARISATION réussie, ce n'est pas possible montrer DÉCELLULARISATION complète d’une cochlée individuelle avant d’utiliser comme un échafaudage. Néanmoins, nous avons trouvé le processus DÉCELLULARISATION soit efficace et robuste.</p>
<p class="jove_content">Les deux étapes les plus critiques dans le protocole actuel sont l’isolement et le traitement du limaçon et la perfusion du limaçon. Grand soin doit être franchie avec la cochlée est initialement isolée, comme la cochlée est délicate et fragile. Si trop de force est appliquée lors de la manipulation de la cochlée avec une pince, la cochlée pourrait fragmenter. Ainsi, il est impératif de traiter chaque cochlée doucement avant DÉCELLULARISATION et décalcification. Le système vestibulaire, si laissé intact, sert de poignée grande pour attraper l’oreille interne complexe avec une pince et orienter la cochlée. De même, lorsque perfusant la cochlée avec des cellules, la pression derrière la perfusion ne doit pas être trop grande, sinon les cellules pourraient ne pas survivre à la perfusion. Comme indiqué dans le protocole actuel, raccorder le tube à la fin d’une seringue de 1 mL insuline permet une manipulation plus facile de la cochlée avec d’une part et une plus grande précision en perfusant les cellules dans la cochlée de l’autre main.</p>
<p class="jove_content">L’étape suivante, il serait naturel pour évaluer les marqueurs du développement et fonctionnelles pour déterminer si l’ECM cochléaire est induisant la différenciation des cellules souches, et si oui, quels composants spécifiques d’ECM dans la cochlée sont induisant des changements dans les cellules. Perfusant les différents types de cellules (<em>par exemple</em>, les cellules souches, neuroprogenitors, fibroblastes, <em>etc.</em>) par le biais de la cochlée au moniteur la différenciation cellulaire et le comportement serait une autre enquête de suivi fascinante. En outre, les cellules qui ont été génétiquement modifiées ou sont exposés à un protocole de différenciation de facteur de croissance peuvent apporter un éclairage nouveau sur comment l’ECM cochléaire prend en charge le développement neurosensoriel et peut-être même fonction dans l’audience. Ainsi, le protocole actuel est une première étape importante dans l’utilisation de l’ECM cochléaire pour étudier le développement neurosensoriel oreille interne et la maintenance, qui pourrait un jour mener à l’élaboration de nouveaux traitements pour restaurer la perte de l’audition.</p>

<p class="jove_content">La cochlée est une structure complexe en spirale dans l’OS temporal. Il est composé d’un labyrinthe osseux externe et un labyrinthe membraneux concentriques, intérieure<sup class="xref">1</sup>. Le labyrinthe membraneux se compose de trois espaces fluides : Scala vestibulaire, Scala media et Scala timbales<sup class="xref">1</sup>. Les médias de scala abrite l’épithélium sensoriel, qui est composé d’une multitude de types de cellules, mais les cellules sensorielles de cheveux (HC), qui transmettre l’énergie mécanique en ondes sonores d’influx nerveux<sup class="xref">2</sup>, sont particulièrement intéressants. Exposition au traumatisme acoustique<sup class="xref">3</sup><sup>,</sup><sup class="xref">4</sup><sup>,</sup><sup class="xref">5</sup>, médicament<sup class="xref">6</sup>, maladie<sup class="xref">7</sup><sup>,</sup><sup class="xref">8</sup>et vieillissement,<sup class="xref">9</sup> peut tous entraîner dysfonctionnement auditif par l’intermédiaire de la mort HC. Perte des cellules ciliées chez les mammifères est permanente, contrairement à aviaires HCs, qui peuvent se régénérer après blessure<sup class="xref">10</sup>.</p>
<p class="jove_content">Une variété de recherches contemporaines ont cherché à restaurer perdus HCs, bien que les approches expérimentales spécifiques varient. Manipulation de l’expression génique dans l’épithélium sensoriel et l’implantation de cellules souches différenciées en dehors du corps sont des approches dominantes dans le domaine, bien que les méthodes d’induction qui cherchent à différencier des cellules souches dans cochlear organoïdes ont été tentative de<sup class="xref">11</sup><sup>,</sup><sup class="xref">12</sup><sup>,</sup><sup class="xref">13</sup>. Chacune de ces approches est soit directement sur les cellules souches, ou les indices de développement utilisés par les cellules souches ; Cependant, une seconde partagé, et potentiellement critiques, élément est l’ECM de la cochlée elle-même<sup class="xref">14</sup><sup>,</sup><sup class="xref">15</sup>.</p>
<p class="jove_content">L’ECM propose non seulement un support physique des cellules et des tissus, ce qui inclut une surface pour l’adhésion cellulaire, la prolifération, la survie et la migration, mais joue également un rôle crucial dans le développement du HCs et la spirale ganglion<sup class="xref">15</sup><sup>,</sup><sup class="xref">16 </sup> <sup>,</sup>,<sup class="xref">17</sup>. Naturellement présents ECM propose des signaux inductifs qui peuvent guider la détermination du phénotype cellulaire et/ou d’adhérence, la prolifération et la survie cellulaire<sup class="xref">18</sup>. Par conséquent, l’utilisation de la cochlée DECELLULARISE en combinaison avec des hWJCs cultivées offrent une occasion unique d’explorer le rôle de la régénération de l’ECM et HC. HWJCs sont un type de cellule facilement disponible, non sujettes à controverse isolé de cordons ombilicaux humains qui se comportent comme des cellules souches mésenchymateuses<sup class="xref">19</sup>. HWJCs ont démontré la capacité de différencier vers le bas de la cellule neurosensorielle lignées<sup class="xref">20</sup><sup>,</sup><sup class="xref">21</sup>. Ainsi, le protocole actuel en détail l’isolement, DÉCELLULARISATION et la perfusion du limaçon de carcasses de souris C57BL avec hWJCs pour l’ingénierie tissulaire oreille interne.</p>

<table><tbody><tr><td>Allegra X-14R Centrifuge</td><td>Beckman-Coulter</td><td>B08861</td><td></td></tr><tr><td>Intramedic Semi-Rigid Tubing</td><td>Becton Dickinson</td><td>427401</td><td></td></tr><tr><td>New Brunswick Innova 2000 Orbital Shaler</td><td>Eppendorf</td><td>M1190-0002</td><td></td></tr><tr><td>Surgical Scissors</td><td>Fine Science Tools</td><td>14060-10</td><td></td></tr><tr><td>Fine Forceps</td><td>Fine Science Tools</td><td>11370-40</td><td></td></tr><tr><td>Ultra-Fine Forceps</td><td>Fine Science Tools</td><td>18155-13</td><td></td></tr><tr><td>50-mL Conical Tubes</td><td>Fisher Scientific</td><td>12565271</td><td></td></tr><tr><td>Petri Dish</td><td>Fisher Scientific</td><td>FB087579B</td><td></td></tr><tr><td>U-100 Insulin Syringe</td><td>Fisher Scientific</td><td>14-829-1B</td><td></td></tr><tr><td>Scintillation Vial</td><td>Fisher Scientific</td><td>03-341-73</td><td></td></tr><tr><td>Rotator</td><td>Fisher Scientific</td><td>88-861-049</td><td></td></tr><tr><td>Transfer Pipette</td><td>Fisher Scientific</td><td>22-170-404</td><td></td></tr><tr><td>Razor Blade</td><td>Fisher Scientific</td><td>12-640</td><td></td></tr><tr><td>Antibiotic-Antimycotic</td><td>Fisher Scientific</td><td>15-240-062</td><td></td></tr><tr><td>Penicillin-Streptomycin</td><td>Fisher Scientific</td><td>15-140-122</td><td></td></tr><tr><td>24-Well Plate</td><td>Fisher Scientific</td><td>07-200-84</td><td></td></tr><tr><td>SuperFrost PLUS Glass Microscope Slides</td><td>Fisher Scientific</td><td>12-550-15</td><td></td></tr><tr><td>Transfer Pipette</td><td>Fisher Scientific</td><td>22-170-404</td><td></td></tr><tr><td>ProLong Gold Antifade Mountant with DAPI</td><td>Fisher Scientific</td><td>P36935</td><td></td></tr><tr><td>Clear-Rite 3</td><td>Fisher Scientific</td><td>22-046341</td><td></td></tr><tr><td>Thermo Scientific Forma Series II 3110 Water-Jacekted CO&lt;sub&gt;2&lt;/sub&gt; Incubator</td><td>Fisher Scientific</td><td>13-998-078</td><td></td></tr><tr><td>Mesenchymal Stem Cell Growth Medium</td><td>Lonza</td><td>PT-3001</td><td></td></tr><tr><td>Trypsin-EDTA</td><td>Lonza</td><td>CC-3232</td><td></td></tr><tr><td>TPP T-75 Culture Flask</td><td>MidSci</td><td>TP90076</td><td></td></tr><tr><td>TPP T-150 Culture Flask</td><td>MidSci</td><td>TP90151</td><td></td></tr><tr><td>TPP T-300 Culture Flask</td><td>MidSci</td><td>TP90301</td><td></td></tr><tr><td>Dissection Microscope</td><td>Nikon Instruments</td><td>SMZ800</td><td></td></tr><tr><td>Nikon Eclipse Ts2R-FL Inverted Microscope</td><td>Nikon Instruments</td><td>MFA51010</td><td></td></tr><tr><td>NuAire Class II, Type A2 Biosafety Cabinet</td><td>NuAire</td><td>NU-425-600</td><td></td></tr><tr><td>1X PBS</td><td>Sigma-Aldrich</td><td>P5368-10PAK</td><td></td></tr><tr><td>1% SDS Solution</td><td>Sigma-Aldrich</td><td>436143-100G</td><td></td></tr><tr><td>10% EDTA</td><td>Sigma-Aldrich</td><td>E9884-100G</td><td></td></tr></tbody></table>

a protocol for decellularizing mouse cochleae for inner ear tissue engineering

<p class="jove_content">Chez les mammifères, les cellules ciliées mécanosensoriels qui facilitent l’absence d’audience la capacité de régénération, qui a limité les traitements pour la perte auditive. Les stratégies actuelles de la médecine régénérative ont mis l’accent sur la transplantation de cellules souches ou manipulation génétique d’entourant les cellules de soutien dans l’oreille interne pour encourager le remplacement des cellules endommagées de tige pour corriger une perte auditive. Pourtant, la matrice extracellulaire (ECM) peut jouer un rôle vital dans l’induction et de maintien de la fonction des cellules ciliées et n’a pas été bien étudiée. À l’aide de la vis d’Archimède ECM comme un échafaudage à cultiver des cellules souches adultes peut-être fournir un aperçu unique Comment la composition et l’architecture de l’environnement extracellulaire aide les cellules à maintenir une fonction d’audition. Nous présentons ici une méthode pour isoler et decellularizing limaçon des souris à utiliser comme échafaudages acceptant les cellules souches adultes perfusés. Dans le protocole actuel, limaçon est isolés des souris euthanasiés, DECELLULARISE et détartré. Par la suite, les cellules de la gelée de Wharton humaine (hWJCs) qui ont été isolés dans le cordon ombilical ont été soigneusement perfusés dans chaque cochlée. Le limaçon servaient de bioréacteurs, et les cellules ont été cultivés pendant 30 jours avant de subir le traitement pour l’analyse. Limaçon DECELLULARISE conservé des structures extracellulaires identifiables, mais n’a pas révélé la présence de cellules ou des fragments d’ADN visibles. Les cellules perfusés dans la cochlée envahirent la plus grande partie de l’intérieur et l’extérieur de la cochlée et grandit sans incident pendant une durée de 30 jours. Ainsi, la méthode actuelle peut servir à étudier comment cochléaire ECM affecte le développement de cellules et de comportement.</p>

<p class="jove_content" fo:keep-together.within-page="1">En utilisant les méthodes présentées ici, DÉCELLULARISATION réussie du limaçon a été évaluée en examinant la présence ou l’absence de l’ADN à travers 4', 6-diamidino-2-phénylindole (DAPI) coloration. Limaçon était considérés comme pleinement DECELLULARISE si l’ADN n’a pas été identifié dans la cochlée DECELLULARISE. Une cochlée native d’une expérience antérieure qui n’a pas subi de DÉCELLULARISATION ou décalcification a été utilisée comme témoin positif pour illustrer les structures et les cellules, on trouves traditionnellement dans une cochlée de souris C57BL. La cochlée DECELLULARISE-décalcifiés, qui n’avait pas tout hWJCs infusé, a été utilisée comme contrôle négatif. Pas quantifiables noyaux cellulaires DAPI coloré ont été observés dans toutes les régions de la section de tissus (<strong class="xfig">Figure 3</strong>). Deux limaçon était DECELLULARISE décalcifiés et puis imprégné de hWJCs pour le projet actuel. Le limaçon DECELLULARISE-détartrer qui est perfusés avec des hWJCs ont été observés d’avoir de nombreux DAPI teinté noyaux dans la cochlée et ça grimpe sur la coque osseuse en dehors de la cochlée (<strong>tableau 1</strong>). Total estimé de la densité des cellules pour la cellule infusé cochlée première étaient 113 759 cellules/cochlée après 30 jours de culture (<strong class="xfig">Figure 4</strong> et <strong class="xfig">Figure 5</strong>) et pour la deuxième cochlée étaient 118 732 cellules/cochlée après 30 jours de culture (<strong class="xfig">Figure 6</strong> et <strong class="xfig">La figure 7</strong>). Ces estimations de la densité cellulaire incluent les timbales Scala Scala media et Scala vestibule et volumes des structures trouvées dans les trois espaces fluides, mais ces estimations exclu tout du labyrinthe osseux restant. En outre, des échantillons de chaque cochlée sont colorés à l’hématoxyline et éosine (H &#38; E) montrer la présence de cellules et des caractéristiques anatomiques dans chaque cochlée (<strong class="xfig">Figure 8</strong>). L’anatomie générale des microstructures cochléaires est resté essentiellement inchangé tout au long de toutes les sections colorées ; Toutefois, aucune cellule n’était identifiables dans la section DECELLULARISE-détartrer (<strong class="xfig">Figure 8 b</strong>). L’identification des noyaux des cellules était radicalement différente entre la cochlée native (<strong class="xfig">Figure 8 a</strong>) et le limaçon DECELLULARISE-détartrer imprégné de cellules (<strong class="xfig">Figure 8</strong><strong>–</strong><strong>D</strong>).</p>
<p class="jove_content" fo:keep-together.within-page="1"><img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56523/56523fig1.jpg"><br>
<strong>Figure 1 : souris OS temporal isolement.</strong> Une fois que la souris a été euthanasiée convenablement, décapiter l’animal par une coupe entre la base du crâne et la première vertèbre cervicale et la tête de pulvérisation avec de l’éthanol pour désinfecter (<strong>A</strong>). Coupent le crâne dans un plan sagittal de la section (<strong>C</strong>de<strong>B</strong><strong>–</strong>) à l’aide des ciseaux chirurgicaux, coupant à travers les tissus du cerveau et les os aiguisés. La tête de souris bissection (<strong>D</strong>, Mandibule inférieure supprimé pour des travaux expérimentaux indépendant) doit alors avoir le tissu de cerveau enlevé pour faire apparaître l’OS temporal. Les deux foramens à travers lequel le col auditif et vestibulaire nerfs (pointes de flèches<strong>E</strong>, rouge) sont utiles indices permettant d’identifier correctement l’OS temporal. Retirez la chair du crâne (<strong>F</strong>). Le conduit auditif externe (<strong>F</strong>, flèche, panneau supérieur) fournit un signal externe utile aussi bien. Soigneusement couper l’OS en excès, laissant juste l’OS temporal isolés (<strong>G</strong>). La bulle tympanique (<strong>H</strong>, panneau supérieur, zone en surbrillance rouge) doit ensuite être délicatement retirée à l’aide de pinces fines (<strong>H</strong>, panneau inférieur). La bulle tympanique est OS minces et peut être facilement train de s’effriter (<strong>I</strong>) jusqu'à ce que le labyrinthe osseux de la cochlée est révélée (<strong>J</strong>, rouge entre parenthèses). Barres d’échelle sur toutes les images sont mm 1. <a href="//ecsource-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56523/56523fig1large.jpg" target="_blank">s’il vous plaît cliquez ici pour visionner une version agrandie de cette figure.</a></p>
<p class="jove_content" fo:keep-together.within-page="1"><img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56523/56523fig2.jpg"><br>
<strong>Figure 2 : Perfusion cochléaire.</strong> Une fois l’OS temporal a été isolé et la bulle tympanique a été supprimée, la cochlée peut ensuite être mises en place une perfusion à un microscope. Lorsque perfusant l’aide d’une seringue entraînée manuellement avec la tuyauterie fixée (<strong>A</strong>, ligne en pointillé), il peut être utile stabiliser la cochlée en utilisant une paire de pinces de fermeture automatique. Doucement, fixez la pince sur la partie vestibulaire de l’OS temporal (<strong>B</strong>) et reposer le forceps contre le rebord de la boîte de Pétri (<strong>A</strong>). Ce qui laisse les deux mains libres pour manipuler des instruments durant la perfusion. Une main peut alors manipuler la seringue, en contrôlant la vitesse d’écoulement du fluide, alors que l’autre main peut positionner le tube sur la fenêtre ovale à l’aide d’une deuxième paire de pinces (<strong>B</strong>). Couper l’extrémité libre du tuyau à un angle permet un positionnement plus facile, permettant la tubulure à placer correctement sans bloquer le champ de vision. Barre d’échelle est 1 mm. <a href="//ecsource-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56523/56523fig2large.jpg" target="_blank">s’il vous plaît cliquez ici pour visionner une version agrandie de cette figure.</a></p>
<p class="jove_content" fo:keep-together.within-page="1"><img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56523/56523fig3.jpg"><br>
<strong>Figure 3 : section de tissu d’une cochlée souris DECELLULARISE.</strong> Un microscope vertical d’épi-fluorescence a été utilisé à l’image des échantillons à un grossissement de 200 X (10 X oculaire et objectif 20 X). Des images ont été cousues ensemble pour créer environ un montage de 4 x 5. DAPI absence d’affichage des cellules de la coloration nucléaire est affiché dans le panneau <strong>A </strong>en utilisant un ensemble de filtre commun DAPI (350 nm excitation, émission à 470 nm) et une surexposée, image en niveaux de gris de l’autofluorescence en utilisant un filtre commun de fluorescéine isothiocyanate (FITC) ensemble (une excitation 490 nm, émission à 525 nm), qui fournit des repères anatomiques, est montré dans le groupe <strong>B</strong>. Du nombre de cellules automatisées ont été effectué dans trois régions d’intérêt, qui sont délimitées sur les panneaux <strong>A</strong> et <strong>B</strong>. Nombre total de cellules ont été calculé pour la section de tissus ensemble (<strong>1</strong>, ligne blanche continue) et la cochlée (<strong>2</strong>, ligne pointillée). Ces deux numéros ont été utilisées pour calculer le nombre de cellules dans les structures osseuses à l’extérieur de la cochlée mais à l’intérieur les bords extérieurs, osseux de la section de tissu (<strong>3</strong>, l’espace entre la ligne pointillée et la ligne pointillée). Barres d’échelle sur toutes les images sont 500 µm.<a href="//ecsource-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56523/56523fig3large.jpg" target="_blank">S’il vous plaît cliquez ici pour visionner une version agrandie de cette figure.</a><p>
<p class="jove_content" fo:keep-together.within-page="1"><img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56523/56523fig4.jpg"><br>
<strong>Figure 4 : une section près de-modiolar de la cochlée DECELLULARISE souris 1 imprégné de hWJCs.</strong> La coloration des noyaux au DAPI apparaît dans panneau <strong>Qu'a</strong> superposer sur une surexposée, autofluorescence en niveaux de gris de la couche verte, qui fournit des repères anatomiques. Automatisée de cellules chefs d’accusation ont été effectuées dans trois régions d’intérêt, qui sont définies dans le groupe <strong>A</strong>. Nombre total de cellules ont été calculé pour la section de tissus ensemble (<strong>1</strong>, ligne blanche continue) et la cochlée (<strong>2</strong>, ligne pointillée). Ces deux numéros ont été utilisées pour calculer le nombre de cellules dans les structures osseuses à l’extérieur de la cochlée mais à l’intérieur les bords extérieurs, osseux de la section de tissu (<strong>3</strong>, l’espace entre la ligne pointillée et la ligne pointillée). La coloration DAPI isolé est figure dans le groupe <strong>B</strong>, et le seuil de la souillure de DAPI utilisé lors de la quantification automatique est à panneau <strong>C</strong>. Barres d’échelle sur toutes les images sont 500 µm. <a href="//ecsource-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56523/56523fig4large.jpg" target="_blank">s’il vous plaît cliquez ici pour visionner une version agrandie de cette figure.</a></p>
<p class="jove_content" fo:keep-together.within-page="1"><img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56523/56523fig5.jpg"><br>
<strong>Figure 5 : vue de fort grossissement d’une section proche-modiolar de la cochlée DECELLULARISE souris 1 imprégné de hWJCs.</strong> La coloration des noyaux au DAPI apparaît dans panneau <strong>Qu'a</strong> superposer sur une image surexposée, de nuances de gris de l’autofluorescence de la couche verte, qui fournit des repères anatomiques. La coloration DAPI isolé est figure dans le groupe <strong>B</strong>, et l’image de seuil utilisée pendant le comptage cellulaire automatisée est à panneau <strong>C</strong>. Les espaces fluides et microstructures plus fines, logés dans la cochlée sont bien préservés pendant la DÉCELLULARISATION et phases de la culture de l’expérience de la cellule. Scala vestibulaire (SV), timbales (SM) de la Scala, Membrane basilaire (BM), Tectorial Membrane (TM), spirale Limbus (L), Canal (RC) de Rosenthal, Modiolus (M), Ligament Spiral (SL) et strie vasculaire (<strong>*</strong>). Membrane de Reissner n’était pas clairement défini dans des sections de tissu, et en conséquence les médias de la Scala n’était pas clairement défini. Barres d’échelle sur toutes les images sont 250 µm. <a href="//ecsource-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56523/56523fig5large.jpg" target="_blank">s’il vous plaît cliquez ici pour visionner une version agrandie de cette figure.</a></p>
<p class="jove_content" fo:keep-together.within-page="1"><img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/56523/56523fig6.jpg"><br>
<strong>Figure 6 : une section près de-modiolar de la cochlée DECELLULARISE souris 2 qui a été imprégnée de hWJCs.</strong> La coloration des noyaux au DAPI est montrée panneau <strong>A</strong>, à superposer sur autofluorescence en niveaux de gris de la couche verte, qui fournit des repères anatomiques. Automatisée de cellules chefs d’accusation ont été effectuées dans trois régions d’intérêt, qui sont définies dans le groupe <strong>A</strong>. Nombre total de cellules ont été calculé pour la section de tissus ensemble (<strong>1</strong>, ligne blanche continue) et la cochlée (<strong>2</strong>, ligne pointillée). Ces deux numéros ont été utilisées pour calculer le nombre de cellules dans les structures osseuses à l’extérieur de la cochlée mais à l’intérieur les bords extérieurs, osseux de la section de tissu (<strong>3</strong>, espace entre les pointillés et les lignes pointillées). La coloration DAPI isolé est figure dans le groupe <strong>B</strong>, et le seuil de la souillure de DAPI utilisé lors de la quantification automatique est à panneau <strong>C</strong>. Barres d’échelle sur toutes les images sont 500 µm. <a href="//ecsource-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56523/56523fig6large.jpg" target="_blank">s’il vous plaît cliquez ici pour visionner une version agrandie de cette figure.</a></p>
<p class="jove_content" fo:keep-together.within-page="1"><img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/56523/56523fig7.jpg"><br>
<strong>Figure 7 : vue de fort grossissement d’une section de près-modiolar de la cochlée DECELLULARISE souris 2 qui a été imprégnée de hWJCs.</strong> La coloration des noyaux au DAPI est montrée panneau <strong>A</strong>, à superposer sur autofluorescence en niveaux de gris de la couche verte, qui fournit des repères anatomiques. La coloration DAPI isolé est figure dans le groupe <strong>B</strong>, et l’image de seuil utilisée pendant le comptage cellulaire automatisée est à panneau <strong>C</strong>. Les espaces fluides et microstructures plus fines, logés dans la vis d’Archimède sont conservées pendant les phases de la culture DÉCELLULARISATION et cellulaire de l’expérience. Scala vestibulaire (SV), timbales (SM) de la Scala, Membrane basilaire (BM), Tectorial Membrane (TM), spirale Limbus (L), Canal (RC) de Rosenthal, Modiolus (M), Ligament Spiral (SL) et strie vasculaire (<strong>*</strong>). Membrane de Reissner n’était pas clairement défini dans des sections de tissu, et en conséquence les médias de la Scala n’était pas clairement défini. Barres d’échelle sur toutes les images sont 250 µm. <a href="//ecsource-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56523/56523fig7large.jpg" target="_blank">s’il vous plaît cliquez ici pour visionner une version agrandie de cette figure.</a></p>
<p class="jove_content" fo:keep-together.within-page="1"><img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/56523/56523fig8.jpg"><br>
<strong>Figure 8 : hématoxyline et éosine coloration de contrôle et traitée limaçon.</strong> L’hématoxyline et éosine coupes affichant une cochlée typique avec des cellules natives présentes colorées est montré dans le groupe <strong>A</strong>. Panneau <strong>B</strong> contient les mêmes structures que le groupe <strong>A</strong>, mais provient d’une cochlée entièrement DECELLULARISE, qui laisse derrière lui de la matrice extracellulaire. Le limaçon démontré précédemment deux imprégné de hWJCs est vus dans les séries <strong>C</strong> et <strong>D</strong>. L’anatomie cochléaire brut reste essentiellement inchangé par les DÉCELLULARISATION et les processus de culture cellulaire, bien que les populations de cellules spécifiques et les lieux est considérablement altérées. Scala vestibulaire (SV), timbales (SM) de la Scala, Membrane basilaire (BM), Tectorial Membrane (TM), spirale Limbus (L), Canal (RC) de Rosenthal, Modiolus (M), Ligament Spiral (SL) et strie vasculaire (*). Barres d’échelle sur toutes les images sont 250 µm. <a href="//ecsource-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56523/56523fig8large.jpg" target="_blank">s’il vous plaît cliquez ici pour visionner une version agrandie de cette figure.</a></p>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td><strong>A1 Cochlée + des cellules</strong></td>
			<td><strong>A2 Cochlée + des cellules</strong></td>
			<td><strong>B2 nég.</strong></td></tr></tbody></table>Contrôle cochlée
		
		<tr>
			<td><strong>Labyrinthe osseux extérieur</strong></td>
			<td>3 758</td>
			<td>549</td>
			<td>0</td>
		</tr>
		<tr>
			<td><strong>Espaces extérieurs de cochlée d’OS</strong></td>
			<td>248</td>
			<td>586</td>
			<td>0</td>
		</tr>
		<tr>
			<td><strong>Au sein de la cochlée</strong></td>
			<td>518</td>
			<td>701</td>
			<td>0</td>
		</tr>
		<tr>
			<td><strong>Total de cellules</strong></td>
			<td>4 524</td>
			<td>1 836</td>
			<td>0</td>
		</tr>
		<tr>
			<td><strong>Section Volume (µL)</strong></td>
			<td>0.0077</td>
			<td>0,0100</td>
			<td>0,0147</td>
		</tr>
		<tr>
			<td><strong>Pour cent du Volume de la cochlée</strong></td>
			<td>0,46 %</td>
			<td>0,59 %</td>
			<td>0,87 %</td>
		</tr>
		<tr>
			<td><strong>Estimation de densité des cellules dans la cochlée</strong></td>
			<td>113 759</td>
			<td>118 732</td>
			<td>0</td>
		</tr>
	

<p class="jove_content" fo:keep-together.within-page="1"><strong>Tableau 1 : nombre de cellules.</strong> DAPI cochléaires coupes coloré ont été binariser et quantifiée à l’aide des outils de comptage automatique dans ImageJ dans 3 régions non-cumul d’intérêt (ne pas le labyrinthe osseux, OS temporal en dehors des espaces fluides cochléaires et au sein de la cochlée). Section transversale cochléaire a également mesuré ImageJ, et cette valeur est utilisée pour calculer le volume de la section en combinaison avec l’épaisseur de coupe (10 µm). En utilisant une estimation du volume total de cochléaire publié par Santi <em>et al.</em> <sup class="xref">23</sup>, le volume cochléaire relatif d’une section donnée a été calculée. La densité des cellules de la cochlée entier a été estimée en utilisant le volume cochléaire relatif d’une section donnée en combinaison avec le nombre d’éléments appariés.</p>

Watch this Scientific Journal Video about A Protocol for Decellularizing Mouse Cochleae for Inner Ear Tissue Engineering at JoVE.com

A Protocol for Decellularizing Mouse Cochleae for Inner Ear Tissue Engineering

Un protocole pour Decellularizing limaçon de souris pour l’ingénierie tissulaire oreille interne

<p class="jove_content">Le but du présent protocole doit démontrer une méthode efficace pour decellularize et détartre limaçon souris pour utilisation comme échafaudages pour applications d’ingénierie tissulaire.</p>

In mammals, mechanosensory hair cells that facilitate hearing lack the ability to regenerate, which has limited treatments for hearing loss. Current ...

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Nanyang Technological University

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Bioengineering

10.7K vues.  University of Kansas Medical Center. Le but du présent protocole doit démontrer une méthode efficace pour decellularize et détartre limaçon souris pour utilisation comme échafaudages pour applications d’ingénierie tissulaire.

Vidéo: A Protocol for Decellularizing Mouse Cochleae for Inner Ear Tissue Engineering

Evolutionary Relationships

<p>Humans have been attempting to properly classify living things since Aristotle made the first attempt during the 4<sup>th</sup> century BC. Aristotle&#8217;s system was improved upon during the Renaissance and then, subsequently, by Carolus Linnaeus in the mid 1700&#8217;s. These more formal classification and organization systems grouped species by their physical similarity to one another. For example, all vertebrates have a backbone, but invertebrates do not. Traits like the backbone are called synapomorphies, which are traits that are shared by a group of organisms, presumably because they were derived from a common ancestor. As we will explore, this method has been shown to have limitations and has more recently been amended to include genetic analysis. Still, scientists construct trees called dendrograms to create a visual representation of how species are related to one another and share common ancestors. These dendrograms can aid in our understanding of the evolutionary processes that drive these relationships. Genetic comparisons have added an important tool guiding the analysis of evolutionary relationships.</p>
<h4>Types of Dendrogram</h4>
<p>A type of dendrogram, called a cladogram, depicts the hypothetical genealogical relationships between species with the tips (or leaves) of the tree representing a species and the branches showing how species are related to each other. A slightly more complicated type of tree, called a phylogram, differs from a cladogram in that the branches leading to the species are of different lengths. The length of a branch in this type of tree represents the degree of change between species: the longer the branch, the more time since the species have diverged from a common ancestor. In both types of tree, the common ancestor of a group of species is indicated by a node, which is the point where a series of branches meet. Species that are more closely related to each other (most recently shared a common ancestor) will be located closest to the node. The two species that share a node are called a sister group<sup>1</sup>.</p>
<h4>Understanding Evolutionary Relationships using Genetic Data</h4>
<p>Historically, cladograms were constructed by comparing the morphology (physical structure) of organisms. This method is still practiced but the techniques have been modernized to include comparison of DNA (deoxyribonucleic acid) sequences between species. Using DNA for building trees has several advantages over relying solely on morphology, including being able to calculate an estimate of how long ago different species shared a common ancestor<sup>1</sup>. However, using DNA is not always feasible, especially when trees include extinct organisms. DNA is best found in soft tissues, which are not preserved during the fossilization process, and therefore it is uncommon for a DNA sample of an extinct species to be available.</p>
<p>DNA is passed on from parents to their offspring in hereditary units called genes. The nucleotide (A, G, C, and T) sequence of genes found in different species are frequently quite similar, likely due to their having come from a common ancestor. This fact allows researchers to align sequences from different species with one another to build the trees described above. Species with more similarity between their nucleotide sequences will be placed next to each other in a tree, and species with less sequence similarity will be placed further apart from each other.</p>
<p>Bioinformatics are the tools used by biologists to analyze large datasets using a combination of computer science, mathematical modeling, and statistics. One such tool is called BLAST (Basic Local Alignment Search Tool), which can be used to quickly search the entire genome of any species that is available in the NCBI (National Center for Biotechnology Information) database<sup>2</sup>. The NCBI database combines several different databases that hold different types of DNA sequence information. The process of a BLAST search includes complex computer algorithms, but basically, BLAST aligns sequences of each nucleotide base from a submitted DNA sequence (known as the <em>query</em> sequence) with sequences in the data base that most closely match it. The DNA sequences that are found will be listed in order of similarity to the sequence in question, and will therefore be from species closely related to the species containing the query gene. This comparison may or may not depict the actual evolutionary relationship between species because genes evolve at different rates. Additionally, genomes sometimes contain more than one instance of a similar sequence.</p>
<p>Comparison of DNA sequences of genes is valuable beyond consideration of evolutionary relationships. Frequently, genes are identified in model organisms, such as the fruit fly, <em>Drosophila melanogaster</em>, or the mouse<sup>3</sup>. Integral to studying a gene, the function of its product is commonly identified and analyzed. If a researcher is interested in studying that function in a different organism (humans for example), BLAST or other bioinformatic tools can be used to find candidate genes based on their similarities to the genes of known function from model organisms.</p>
<p>Human genes can also be used as the starting point to find homologs in model organisms. In fact, human disease research depends heavily on this. Once a human gene of interest is identified, mice can be genetically manipulated to have the homologous gene disrupted, or &#8220;knocked out,&#8221; creating a model of the human disease that can be studied in order to understand and treat the disease. There are many of these mouse strains currently available. For example, there is a mouse model for human Cystic Fibrosis (CF) called the <em>Cftr</em> knockout mouse and another modeling atherosclerosis, called the <em>Apoe</em> knockout<sup>3</sup>.<br />
 </p>
<h4>References:</h4>
<ol class="references">
	<li><strong>Annenberg Learner.</strong> Online Textbook: Unit 3 Evolution and Phylogenetics. <em>Rediscovering Biology. </em>[Online] 2017. [Cited: August 23, 2018.] https://www.learner.org/courses/biology/textbook/compev/compev_3.html.</li>
	<li><strong>National Center for Biotechnology Information, U.S. National Library of Medicine.</strong> BLAST: Basic Local Alignment Search Tool. [Online] https://blast.ncbi.nlm.nih.gov/Blast.cgi.</li>
	<li><strong>National Human Genome Research Institute.</strong> Background on Mouse as a Model Organism. <em>National Human Genome Research Institute. </em>[Online] December 2002. [Cited: August 23, 2018.] <a href="https://www.genome.gov/10005834/background-on-mouse-as-a-model-organism/" target="__blank">https://www.genome.gov/10005834/background-on-mouse-as-a-model-organism/</a>.</li>
</ol>

Evolutionary Relationships: Using BLAST to Test Evolutionary Hypotheses - Concept

Relations évolutives

Humans have been attempting to properly classify living things since Aristotle made the first attempt during the 4th century BC. Aristotle&#8217;s system ...

Enseignement

JoVE Lab Manual

Biologie

Evolution

Extraction

Extraction: Liquid-Liquid and Acid-Base Extraction - Procedure

Source: Lara Al Hariri and Ahmed Basabrain at the University of Massachusetts Amherst, MA, USA

	Cellulose RecoveryExpand
	Extraction and filtration can ...

Chimie

Lab Manual Sub Articles

Electrolytic Cells

<p>Source: Smaa Koraym at Johns Hopkins University, MD, USA</p>
<ol class="note-items">
	<li data-note-item="1" data-parent-collapse="">Electrolytic Cells<button class="expand">Expand</button>
	<p>In this experiment, you will be assembling an electrolytic cell to perform electroplating. An electrolytic cell contains an anode and cathode connected by a power source, which drives the nonspontaneous redox reaction in the cell. When the anode material is oxidized, the ions travel to the cathode, where they are reduced and plated.</p>
	<p>In the case of this experiment, the electrolytic cell contains a copper anode and a brass key for the cathode. Oxidation of the anode creates copper(2+) ions, which are then reduced on the brass key, forming a thin plating of solid copper.</p>
	<ul class="lab-items">
		<li data-sub-note-item="1-3">
		<div class="lab-step">Put on gloves, safety goggles, and a lab apron, which must be worn at all times.</div>
		</li>
		<li data-sub-note-item="1-4">
		<div class="lab-step">Obtain one 10-ohm resistor, one copper sheet electrode, one copper wire, and one piece of emery paper.</div>
		</li>
		<li data-sub-note-item="1-5">
		<div class="lab-step">Use the emery paper to polish the brass key and the copper sheet. Rinse the key and the copper electrode with water. The brass key will be the cathode, and the copper sheet will be the anode.</div>
		</li>
		<li data-sub-note-item="1-6">
		<div class="lab-step">Take one 50-mL beaker and a watch glass to the instructor&#39;s hood and obtain approximately 20 mL of acetone. Place the watch glass on top of the beaker to prevent evaporation and bring it back to your bench.</div>
		</li>
		<li data-sub-note-item="1-7">
		<div class="lab-step">Label the other 50-mL beaker as &#39;organic waste&#39;.</div>
		</li>
		<li data-sub-note-item="1-8">
		<div class="lab-step">Use a disposable pipette to rinse the copper electrode and key with acetone. Collect excess in the waste beaker. Place the key and copper electrode on a paper towel to dry.</div>
		</li>
		<li data-sub-note-item="1-9">
		<div class="lab-step">Measure the initial masses of the key and copper electrode and record their values in your lab notebook.
		<h4><strong>Table 1: Moles of Cu Transferred</strong></h4>
		<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
			<tbody>
				<tr>
					<td></td>
					<td style="text-align: center;"><strong>Mass of key (g) </strong></td>
					<td style="text-align: center;"><strong>Change in mass (g) </strong></td>
					<td style="text-align: center;"><strong>Moles Cu plated </strong></td>
					<td style="text-align: center;"><strong>Mass of Cu electrode (g) </strong></td>
					<td style="text-align: center;"><strong>Change in mass (g) </strong></td>
					<td style="text-align: center;"><strong>Moles Cu lost </strong></td>
				</tr>
				<tr>
					<td style="text-align: center;">0 min</td>
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					<td style="text-align: center;">5 min</td>
					<td></td>
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					<td></td>
					<td></td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">1<sup>st</sup> 30 min</td>
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					<td></td>
					<td></td>
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				</tr>
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					<td style="text-align: center;">2<sup>nd</sup> 30 min</td>
					<td></td>
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				</tr>
			</tbody>
		</table>
		<a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/53/Table_1._Moles_of_Cu_Transferred.xlsx" target="_blank">Click Here to download Table 1</a></div>
		</li>
		<li data-sub-note-item="1-10">
		<div class="lab-step">Set up the electrolytic cell. Use one electrical wire with alligator clips to connect the negative terminal on the battery to the current probe. Wrap the 10-ohm resistor around the other end of the current probe.</div>
		</li>
		<li data-sub-note-item="1-11">
		<div class="lab-step">Use another wire to connect the positive terminal on the battery to the copper electrode.</div>
		</li>
		<li data-sub-note-item="1-12">
		<div class="lab-step">Wrap the copper wire around the brass key and use an electrical wire to connect the resistor on the current probe to the copper wire on the brass key.</div>
		</li>
		<li data-sub-note-item="1-13">
		<div class="lab-step">Obtain about 200 mL of concentrated copper sulfate in a 250-mL beaker. The amount does not need to be exact. Bring the beaker back to your bench and set it on the stir plate. Carefully, place a magnetic stir bar in the beaker and turn the stir setting on to high.</div>
		</li>
		<li data-sub-note-item="1-14">
		<div class="lab-step">Connect the data acquisition system to the current probe and allow the data acquisition system to sense the probe. It should return a reading in amps. Change the rate to two scans per min and the duration to 5 min.</div>
		</li>
		<li data-sub-note-item="1-15">
		<div class="lab-step">Place the brass key in the beaker. Hang it off the side of the beaker so that the key is almost completely submerged but does not touch the magnetic stir bar. <strong>Note: </strong>The copper wire should not be in the solution.</div>
		</li>
		<li data-sub-note-item="1-16">
		<div class="lab-step">Place the copper electrode in the beaker, bending it so that it hangs over the edge of the beaker. Ensure that it is not touching the stir bar or the brass key.</div>
		</li>
		<li data-sub-note-item="1-17">
		<div class="lab-step">Start the scan and let it collect data for 5 min. This will act as a test run to make sure plating occurs on the brass key.</div>
		</li>
		<li data-sub-note-item="1-18">
		<div class="lab-step">After the scan ends, remove the key and copper electrode. The copper plating on the brass key should be evident. If it is not visible, check your circuit to make sure that the battery is connected in the correct orientation.</div>
		</li>
		<li data-sub-note-item="1-19">
		<div class="lab-step">Once you have completed a successful 5-min plating, disconnect the key and copper electrode from the wiring. Remove the copper wire from the key and rinse the key and copper electrode with water and then acetone. Lay them on a paper towel to dry.</div>
		</li>
		<li data-sub-note-item="1-20">
		<div class="lab-step">Measure the masses of the key and the copper electrode and record them in your lab notebook.</div>
		</li>
		<li data-sub-note-item="1-21">
		<div class="lab-step">Reconnect the key and copper electrode and place them back in the solution like before. Restart the run on the data acquisition system, making sure that the rate is set to two scans per min. With the duration set to 30 min, allow the electroplating process to run.</div>
		</li>
		<li data-sub-note-item="1-22">
		<div class="lab-step">After the scan is complete, remove the electrode and key from the solution, disconnect them from the wires, and rinse them with water and then acetone. Allow them to dry on a paper towel, then weigh them and record their masses in your notebook.</div>
		</li>
		<li data-sub-note-item="1-23">
		<div class="lab-step">Reassemble the setup and repeat the electroplating for a second 30-min trial. When the scan is complete, remove the key and copper electrode, rinse them like before, and record their mass.</div>
		</li>
		<li data-sub-note-item="1-24">
		<div class="lab-step">Save the recorded data as a text file on a flash drive.</div>
		</li>
		<li data-sub-note-item="1-25">
		<div class="lab-step">To clean up, turn the stir setting off and remove the stir bar from the solution using forceps. Dispose of the acetone in the organic waste container provided by your instructor. The copper sulfate solution can be reused for future labs, so pour the concentrated copper sulfate back into the carboy.</div>
		</li>
		<li data-sub-note-item="1-26">
		<div class="lab-step">Rinse all of your glassware with water and return it to the instructor.</div>
		</li>
	</ul>
	</li>
	<li data-note-item="2" data-parent-collapse="">Results<button class="expand">Expand</button>
	<ul class="lab-items">
		<li data-sub-note-item="2-1">
		<div class="lab-step">First, calculate the total charge Q that flowed through the system during the different trials. One coulomb is the quantity of electricity flowing through a wire carrying 1 ampere of current in 1 s, thus Q = I x t.</div>
		</li>
		<li data-sub-note-item="2-2">
		<div class="lab-step">Review the current probe data for the first 5-min electroplating trial.<strong> Note: </strong>If the current dropped to 0 or close to 0 at the end of the trial, then the electrons stopped flowing from the copper electrode to the key. Only use the values recorded before the drop in current occurred. If the current drop to 0 early in the plating process, repeat the trial.</div>
		</li>
		<li data-sub-note-item="2-3">
		<div class="lab-step">The current varies over time, so calculate the average current, I.
		<h4><strong>Table 2: Moles of Electrons Transferred</strong></h4>
		<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
			<tbody>
				<tr>
					<td style="text-align: center;"><strong>Trial</strong></td>
					<td style="text-align: center;"><strong>Time (s) </strong></td>
					<td style="text-align: center;"><strong>Current<sub>average</sub> (A) </strong></td>
					<td style="text-align: center;"><strong>Total charge (C) </strong></td>
					<td style="text-align: center;"><strong>Moles (e?) </strong></td>
				</tr>
				<tr>
					<td style="text-align: center;">5 min</td>
					<td></td>
					<td></td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">1<sup>st</sup> 30 min</td>
					<td></td>
					<td></td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">2<sup>nd</sup> 30 min</td>
					<td></td>
					<td></td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">Total</td>
					<td></td>
					<td></td>
					<td></td>
					<td></td>
				</tr>
			</tbody>
		</table>
		<a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/53/Table_2._Moles_of_Electrons_Transferred.xlsx" target="_blank">Click Here to download Table 2</a></div>
		</li>
		<li data-sub-note-item="2-4">
		<div class="lab-step">Use the average current over the total time to determine the total charge transferred during electroplating, Q. About 20 coulombs were transferred during the 5-min trial, and between 110 and 120 coulombs were transferred during each 30-min trial. The total charge transferred over the entire 65-min electroplating process was about 250 coulombs.</div>
		</li>
		<li data-sub-note-item="2-5">
		<div class="lab-step">Use the total charge transferred to calculate the moles of electrons plated during each trial. The moles of electrons plated equals the total charge transferred divided by the Faraday constant. The Faraday constant represents the magnitude of electric charge per mole of electrons and is determined by multiplying the charge of an electron by Avogadro&#39;s constant.</div>
		</li>
		<li data-sub-note-item="2-6">
		<div class="lab-step">Determine the moles of electrons that were transferred for the 5-min and 30-min trials.</div>
		</li>
		<li data-sub-note-item="2-7">
		<div class="lab-step">Calculate the moles of copper that were plated on the key after each trial. Similarly, determine the moles of copper lost from the copper sheet electrode.</div>
		</li>
		<li data-sub-note-item="2-8">
		<div class="lab-step">Compare the moles of copper that were lost from the electrode to the amount that was plated to the brass key. The numbers should be close.</div>
		</li>
		<li data-sub-note-item="2-9">
		<div class="lab-step">Check to see if these values match what is expected based on the number of moles of electrons that were transferred. The reduction of 1 mole of copper(2+) ions requires 2 moles of electrons to produce 1 mole of solid copper.</div>
		</li>
	</ul>
	</li>
</ol>

Electrolytic Cells: Construction and Electroplating - Procedure

Cellules électrolytiques

Source: Smaa Koraym at Johns Hopkins University, MD, USA

	Electrolytic CellsExpand
	In this experiment, you will be assembling an electrolytic cell to ...

Galvanic Cells

Galvanic Cells: Construction and Reduction Potential Measurement - Procedure

Cellules galvaniques

Source: Smaa Koraym at Johns Hopkins University, MD, USA

	Constructing a Lead-Copper Galvanic CellExpand
	In this experiment, you will construct a ...

Cell Division

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Observing the Cell Cycle in a Root Tip
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">Hypotheses: The experimental hypothesis is that in root tips slices that have been treated with nocodazole, a chemical that interferes with microtubular polymerization, all of the cells will be arrested at the same stage of the cell cycle and that in untreated onion tip slices all of the different stages of the cell cycle will be visualized. The null hypothesis is that there will be no differences in mitosis between the two groups and that different stages of the cell cycle will be visualized. 
</div></li><li data-sub-note-item="1-2"><div class="lab-step">To prepare for this experiment you will use an onion that has been placed in water for several days until new white root tips have begun to sprout. Use a razor blade to cut a 1 cm piece from the end of one of the root tips. 
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Place the slice of onion into a microcentrifuge tube labeled &#8220;N&#8221; for the nocodazole condition. 
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Then, cut a second 1 cm piece of onion root tip and place in a microcentrifuge tube labeled &#8220;C&#8221; for the control condition.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Add 2 &#181;L of the 5 mg/mL nocodazole to 1 mL of water in the microcentrifuge tube labeled N and 1 mL of water only to the tube labeled C. 
</div></li><li data-sub-note-item="1-6"><div class="lab-step">Leave the tubes containing the root tips to incubate for 12 hours and wipe down the workspace with 70% ethanol.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">When incubation is complete, set a heat block to 60 &#186;C.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">Wash the nocodazole treated tips with water and then aspirate the liquid.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">Repeat this wash two more times, for a total of three washes. 
</div></li><li data-sub-note-item="1-10"><div class="lab-step">Fill both tubes with 1 mL of one normal hydrochloric acid. Incubate the tubes in a 60 &#186;C heat block for 12 - 15 min, then discard the acid into a waste flask.
</div></li><li data-sub-note-item="1-11"><div class="lab-step">Now, add 1 mL of distilled water drop wise to the tubes at least three times to rinse the samples.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">Now add one microliter of DAPI stain to 10 mL of phosphate buffered saline (PBS).
</div></li><li data-sub-note-item="1-13"><div class="lab-step">Then place 50 &#181;L of this dilute DAPI stain into each tube, and leave them at room temperature to incubate for 12 min. 
</div></li><li data-sub-note-item="1-14"><div class="lab-step">After this, preform three distilled water washes as in step 10. 
</div></li><li data-sub-note-item="1-15"><div class="lab-step">Label one microscope slide as 'Control' and another as 'Nocodazole'. 
</div></li><li data-sub-note-item="1-16"><div class="lab-step">Transfer the stained roots onto their respective microscope slides and add a drop of water to the control slide and a drop of nocodazole to the nocodazole slide.
</div></li><li data-sub-note-item="1-17"><div class="lab-step">Use the razor blade to remove the unstained portions of each root tip and place a glass cover slip over each sample.
</div></li><li data-sub-note-item="1-18"><div class="lab-step">Carefully press down on each cover slip with a gloved fingertip, and then let the slides sit for 10 min. 
</div></li><li data-sub-note-item="1-19"><div class="lab-step">Finally, view the slides under the 40X objective of a fluorescence microscope and capture images of your root tip cell preparations.
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Understanding the Loss of Cell Cycle Control
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">NOTE: Before cells can pass through all of the stages of the cell cycle there are several checkpoints they must go through. These checkpoints are regulated by cyclin and cyclin-dependent kinase proteins, or CDKs. If the cyclins and CDKs determine that previous cellular events have not been adequately completed they will arrest the cells at this step and prevent them from proliferating. In cancer cells one or several of these molecular restraints for regulating cell division are lost and this allows the damaged cells to escape from cell proliferation control. These abnormal cells develop into tumors which are masses of uncontrolled proliferating cells that interfere with the normal functions of the body. Notice how before it divides this cancer cell rounds up and refracts light very strongly under the light microscope. This is because not all cancer cells successfully divide into daughter cells after each round of mitosis. This will mean that they often contain an excess of organelles and DNA and this increase in cellular content means they will refract light more intensely than normal cells do. 
</div></li></ul></li><li data-note-item="3" data-parent-collapse="">Results
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="3-1"><div class="lab-step">Examine your collected images and record in Table 1 the different cell structures and stages that could be identified in the onion tip slice that was treated with nocodazole. <a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/10/Table_1_Cell_division.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 1</a>
</div></li><li data-sub-note-item="3-2"><div class="lab-step">If there are any cells undergoing mitosis, note what stages of cell division you see.
</div></li><li data-sub-note-item="3-3"><div class="lab-step">Record the percentage of cells in each stage of mitosis in Table 1. 
</div></li><li data-sub-note-item="3-4"><div class="lab-step">Next, examine the images of the untreated onion tip slice, and record what cell structures can you identify. 
</div></li><li data-sub-note-item="3-5"><div class="lab-step">If there are any cells undergoing mitosis, consider what stages of cell division you see, and record the percentages of cells in each stage of mitosis in the Table 1. 
</div></li></ul></li></ol>

Cell Division: Observing Cell Cycle in a Root Tip - Procedure

Division cellulaire

Observing the Cell Cycle in a Root Tip
ExpandHypotheses: The experimental hypothesis is that in root tips slices that have been treated with nocodazole, a ...

Processus cellulaires

Cell Structure

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Visualizing Onion and Human Cells
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">In this experiment you will prepare different types of cells from a plant and animal, onion and human respectively, and then visualize them under a microscope. HYPOTHESES: The experimental hypothesis is that the cells will appear different in overall shape and membrane structure, but there will be some shared similar structures, like nuclei, in both cells. The null hypothesis of the experiment is that there will be no difference in structure between the two cell types.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">First, to prepare an animal cell slide, start by pouring 30 mL of distilled water into a beaker.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Then, use a glass dropper to place 2 - 3 drops of the water onto the center of a clean microscope slide.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Next, take a clean toothpick and scrape the inside of your cheek to gather cells.  CAUTION: Do not scrape your cheeks too aggressively. The scraping should not cause discomfort or bleeding. 
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Put the end of the toothpick with the cheek cells onto the wetted area of the slide and mix the water and cheek cells together.
</div></li><li data-sub-note-item="1-6"><div class="lab-step">Next, add two drops of methylene blue staining solution to the wetted area of the slide and mix it in with a clean toothpick.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Let the mixture sit at room temperature for 3 minutes.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">After 3 minutes have passed, use a glass dropper to add a drop of glycerol to the mixture.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">Carefully, place a cover slip over the mix on the slide by first standing one end of the cover slip vertically and aligning it along to one side of the mixture. Then, carefully lower the other end of the cover slip down until the mixture is completely covered.
</div></li><li data-sub-note-item="1-10"><div class="lab-step">Finally, remove any excess liquid with blotting paper.
</div></li><li data-sub-note-item="1-11"><div class="lab-step">Next, prepare a slide of plant cells by placing five drops of distilled water into a clean watch glass.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">Then, with forceps, take a thin strip of onion and place it into the water.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">Apply 5 drops of safranin solution to another watch glass.
</div></li><li data-sub-note-item="1-14"><div class="lab-step">Now using a pair of forceps transfer the piece of onion from the distilled water into the safranin.
</div></li><li data-sub-note-item="1-15"><div class="lab-step">Allow the onion to soak for 30 seconds.
</div></li><li data-sub-note-item="1-16"><div class="lab-step">Then, again with forceps, move the onion back into the distilled water.
</div></li><li data-sub-note-item="1-17"><div class="lab-step">Now, place 3 drops of glycerol onto the center of a glass microscope slide.
</div></li><li data-sub-note-item="1-18"><div class="lab-step">Using the forceps again, transfer the onion into the glycerol on the slide.
</div></li><li data-sub-note-item="1-19"><div class="lab-step">Gently place a cover slip over the onion on the slide and blot away any excess liquid with a piece of blotting paper.
</div></li><li data-sub-note-item="1-20"><div class="lab-step">Next, you will visualize the slides you have prepared using a compound microscope. First, turn on the scope and set the objective lens to the lowest level of magnification.  NOTE: A compound microscope typically has two levels of magnification.
</div></li><li data-sub-note-item="1-21"><div class="lab-step">Place the cheek tissue slide onto the stage of the microscope and bring the cells into focus by adjusting the stage and using the coarse adjustment knob.
</div></li><li data-sub-note-item="1-22"><div class="lab-step">When the cells are in focus, observe the individual cells and draw what you see. Repeat these observations at medium and high magnifications adjusting the focus with the fine adjustment knob. IMPORTANT: Do NOT use the coarse adjustment knob at high magnifications. 
</div></li><li data-sub-note-item="1-23"><div class="lab-step">Next, rotate the nose piece so that the oil lens is almost in place.
</div></li><li data-sub-note-item="1-24"><div class="lab-step">Place a drop of immersion oil on top of the cover slip of the cheek cell slide.
</div></li><li data-sub-note-item="1-25"><div class="lab-step">Use the oil lens to view the cheek cells and record your observations.
</div></li><li data-sub-note-item="1-26"><div class="lab-step">Repeat these steps to view the onion plant cell, moving again from low to medium, high, and oil lenses (steps 20 &#8211; 24).
</div></li><li data-sub-note-item="1-27"><div class="lab-step">Record all of your observations and note the differences you see between animal and plant cells.
</div></li><li data-sub-note-item="1-28"><div class="lab-step">When you are finished, place the cheek cell slide and toothpicks into a diluted bleach solution. Dispose of the cover slips into a glass waste container.
</div></li><li data-sub-note-item="1-29"><div class="lab-step">Finally, turn off the microscope and clean the lenses with lens cleaning paper.
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Results
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">Calculate the total magnification for each of the viewing levels. Remember to account for both the eyepiece magnification and the objective lens magnification. NOTE: Eye piece: 10X magnification, Objective lens: 4X, 10X, and 40X magnification
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Then, identify the structural differences between plant and animal cells using the observations made under the microscope.
</div></li><li data-sub-note-item="2-3"><div class="lab-step">Label the nuclei in both cell types and then highlight or note any differences that you see between them.
</div></li></ul></li></ol>

Cell Structure: Visualizing Onion and Human Cells - Procedure

Structure cellulaire

Visualizing Onion and Human Cells
ExpandIn this experiment you will prepare different types of cells from a plant and animal, onion and human ...

Les bases

Genetics of Organisms

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">REF and WEF Fly Genetics
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">NOTE: At your station, you should have four vials of flies. They will be labeled as male or female. They will also be labeled with the notation REF for red-eyed flies or WEF for white-eyed flies. The female vial should also be labeled virgin as these flies should be unmated for the purpose of this experiment.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">Make sure you have around five flies in each vial. NOTE: You will perform crosses using these flies to determine if red or white eyes is dominant and if the trait is sex-linked. You should also have two fly anesthesia wands, a paint brush, magnifying glass, fly anesthetic, some empty fly vials, and a sheet of white paper.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Draw a line in the middle of the paper to make a male side and a female side.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Dip the fly anesthesia wand into the solution and insert it into the red-eyed female vial until the flies stop moving.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Then, dip the same wand into the white-eyed male vial until these flies are also anesthetized.
</div></li><li data-sub-note-item="1-6"><div class="lab-step">Tap the flies from the two vials onto the sheet - Females on one side and males on the other.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Check to make sure that all the flies from the female vial are female and likewise, confirm that the male flies are male.  NOTE: Males are slightly smaller and have a darker solid pigment on the end of their abdomen. Females are larger and have stripes rather than a solid spot.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">If there are any male flies in the female vial, obtain a new vial as these females likely have sperm from males from their own strain and are unusable.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">Now, move the five red-eyed females and five white-eyed males into a new vial using the paintbrush.
</div></li><li data-sub-note-item="1-10"><div class="lab-step">Label this vial to indicate that this is a cross between red-eyed females and white-eyed males. HYPOTHESES: Use Punnett squares to formulate hypotheses for this experiment. Consider what you will see in a few days if the white allele is dominant and there is no effect of parental sex. What if red is dominant? Under the potential offspring genotype in each box, write the expected phenotype.
</div></li><li data-sub-note-item="1-11"><div class="lab-step">To set up the reverse cross, again dip the fly anesthesia wand into the solution and insert it this time into the white-eyed female vial and then the red-eyed male vial until all the flies have stopped moving.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">Tap the flies onto a sheet of white paper and check to make sure that all the flies from the female vial are female and likewise with the male flies.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">Now, move the five white-eyed females and five red-eyed males into a new vial using the paintbrush.
</div></li><li data-sub-note-item="1-14"><div class="lab-step">Label this vial to indicate that this is a cross between white-eyed females and red-eyed males.
</div></li><li data-sub-note-item="1-15"><div class="lab-step">Finally, place the two tubes containing the new crosses into a rack for incubation at room temperature. NOTE: After about three days, larvae should be moving around in the vial.
</div></li><li data-sub-note-item="1-16"><div class="lab-step">Remove the 10 adult flies and dispose of them into the morgue tube containing 70% ethanol.
</div></li><li data-sub-note-item="1-17"><div class="lab-step">Now, put the tubes back in the rack and let the larvae mature until they reach the adult stage in up to two weeks.
</div></li><li data-sub-note-item="1-18"><div class="lab-step">Once all of the larvae from these crosses have matured to adulthood, anesthetize the flies in one of the tubes using the wand.
</div></li><li data-sub-note-item="1-19"><div class="lab-step">Then, on a piece of white paper, sort the flies by sex and phenotype and then record these data.
</div></li><li data-sub-note-item="1-20"><div class="lab-step">Now, repeat this process and tally the sexes and phenotypes for the second cross (steps 1 &#8211; 18).
</div></li><li data-sub-note-item="1-21"><div class="lab-step">Finally, once you have counted all of your flies, dispose of them in the morgue vial.
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Results
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">To begin investigating your results, first compare the data you collected with the hypothesis you made using the Punnett squares.
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Did your data match the simple Punnett square hypothesis where red or white is dominant and there is no effect of sex? Compare your data with other lab groups.
</div></li><li data-sub-note-item="2-3"><div class="lab-step">If your data does not match what your Punnett squares predicted, does this indicate that the gene may be sex-linked?
</div></li><li data-sub-note-item="2-4"><div class="lab-step">If the red gene were dominant and sex-linked, when we cross red-eyed females with white-eyed males, all of the offspring would have red eyes. Assuming all of the females were homozygous for the color allele, does this match your data for the first cross?
</div></li><li data-sub-note-item="2-5"><div class="lab-step">If white were dominant and the gene were sex-linked, we would see the exact opposite. So all of the flies would have white eyes. From your data, can you conclude from your first cross whether red or white is dominant?
</div></li><li data-sub-note-item="2-6"><div class="lab-step">When the reciprocal cross of white-eyed females and red-eyed males is performed, if the red phenotype is dominant, all of the females would be red and all of the males would be white. This would also confirm that the gene is specifically x-linked because white-eyed females could then only be produced in crosses with white-eyed males. Did your second cross produce any white-eyed females?
</div></li><li data-sub-note-item="2-7"><div class="lab-step">Finally, if the gene were x-linked and white was dominant, crossing white-eyed females with red-eyed males would result in all of the offspring having white eyes. Does this hypothesis fit with your second cross?
</div></li><li data-sub-note-item="2-8"><div class="lab-step">Look over your data and conclude which of the hypotheses best fits the patterns you found in your crosses. If your data does not fit with any of these, hypothesize why this might be the case.
</div></li></ul></li></ol>

Genetics of Organisms: REF and WEF Fly Genetics - Procedure

Génétique des organismes

REF and WEF Fly Genetics
ExpandNOTE: At your station, you should have four vials of flies. They will be labeled as male or female. They will also be ...

Génétique

Microbial and Fungal Diversity

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Examining Microbial Colony Structures
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">To begin, collect your allocated culture plates and then remove the Parafilm and lid from one of them.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">Place the uncovered plate under a dissecting microscope and then observe the individual communities within each quadrant of the plate.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">As the colonies are identified, fill out the observed colony properties in the appropriate column in the table.  <a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/28/Table_1_Microbial_diversity.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 1</a>
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Then, repeat the observations for each of the other two bacterial species.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">To perform Gram staining of your three bacterial species, first label the ends of three microscope slides with the initials of each bacteria species, your initials, and the date. HYPOTHESES: In this experiment, the experimental hypothesis might be that it is possible to elucidate different bacterial species and shapes using Gram staining, and that Gram-positive bacteria will stain purple with crystal violet whereas Gram-negative bacteria will appear pink due to the safranin counterstain. The null hypothesis could be that all of the bacterial types will retain both stains equally, and staining will not help elucidate their species or shape.
</div></li><li data-sub-note-item="1-6"><div class="lab-step">To begin the first stain, attach a clip to the slide for ease of handling. For each bacterial smear, remember to be sure to use a cleaned and flame-sterilized inoculating loop.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Place two loopfuls of sterile distilled water onto the center of the slide.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">After sterilizing the loop again, cool the loop by touching an uninoculated portion of the agar several times.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">Then, remove a small amount of culture from an isolated colony and mix it with the water on the slide.
</div></li><li data-sub-note-item="1-10"><div class="lab-step">Allow the slide to dry at room temperature and heat-fix the smear by quickly passing it through a flame.
</div></li><li data-sub-note-item="1-11"><div class="lab-step">Once dry, pipette several drops of crystal violet onto the slide and let it sit for two to three minutes.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">Carefully rinse the slide with distilled water by aiming the direct flow towards the top of the slide, allowing the water to gently flow down. Do not aim the water flow directly at the smear.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">Next, cover the slide with Gram&#39;s iodine, leave it to sit again for two minutes, and then gently rinse with distilled water.
</div></li><li data-sub-note-item="1-14"><div class="lab-step">Decolorize the slide with 95% ethanol until stain no longer washes away.
</div></li><li data-sub-note-item="1-15"><div class="lab-step">Immediately, rinse the slide with distilled water. This will limit over-decolorizing.
</div></li><li data-sub-note-item="1-16"><div class="lab-step">Then, add several drops of safranin counterstain and leave for 30 s. This will stain any Gram-negative cells present.
</div></li><li data-sub-note-item="1-17"><div class="lab-step">Gently rinse the slide with distilled water and blot dry with absorbent paper.
</div></li><li data-sub-note-item="1-18"><div class="lab-step">After staining the other two slides in the same manner (steps 6 &#8211; 17), observe each with a microscope.
</div></li><li data-sub-note-item="1-19"><div class="lab-step">Use a low power objective first to make coarse adjustments. When you find an ideal section, move on to the higher magnifications.
</div></li><li data-sub-note-item="1-20"><div class="lab-step">After further imaging and adjusting the smear with a medium power objective, add immersion oil directly to the smear. NOTE: The oil is needed for high power objectives that will provide the best micrographs.
</div></li><li data-sub-note-item="1-21"><div class="lab-step">Now, use colored pencils to draw the observed bacteria in the appropriate magnification circles and fill in the observed characteristics in the appropriate column, noting the bacteria&#39;s shape and whether it is Gram-positive, identified by darker purple staining, or Gram-negative, identified with pink staining. <a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/28/Table_2_Microbial_diversity.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 2</a>
</div></li><li data-sub-note-item="1-22"><div class="lab-step">Finally, repeat the observation, drawing, and characterization for the other two stained bacterial slides.
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Examining Fungal Structures
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">Remove the <em>S. cerevisiae</em> plate from the incubator and place a drop of water onto a microscope slide.
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Using a sterilized inoculating loop, smear a film of yeast into the drop of water and place a cover slip over the slide.
</div></li><li data-sub-note-item="2-3"><div class="lab-step">Place the slide under the microscope at the lowest magnification, increasing the magnification to the oil immersion setting to see the individual yeast cells.
</div></li><li data-sub-note-item="2-4"><div class="lab-step">Observe the general shape and structure of the yeast and identify any cells that are undergoing asexual reproduction by budding.
</div></li><li data-sub-note-item="2-5"><div class="lab-step">Now, draw the cells in the appropriate magnification circles.
</div></li><li data-sub-note-item="2-6"><div class="lab-step">When you are done observing the yeast, replace the <em>S. cerevisiae</em> slide with a prepared Ascomycetes apothecium slide and set the microscope to the lowest magnification.
</div></li><li data-sub-note-item="2-7"><div class="lab-step">Using colored pencils, draw the structures at low and high magnifications, including the asci and ascospores.
</div></li><li data-sub-note-item="2-8"><div class="lab-step">Finally, place a button mushroom, <em>Agaricus bisporus</em>, under the dissecting microscope and draw the features, including the pileus, or cap, the gills, and the stipe, or stalk.
</div></li></ul></li><li data-note-item="3" data-parent-collapse="">Results
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="3-1"><div class="lab-step">Using the coloring of the bacteria recorded in the table, determine which species are Gram-positive and which are Gram-negative.
</div></li><li data-sub-note-item="3-2"><div class="lab-step">Then, look at the shapes recorded in the table to determine which bacterial species are bacillus, coccus, or spirillum.
</div></li><li data-sub-note-item="3-3"><div class="lab-step">Next compare your illustrations of the yeast cells to your illustrations of the bacterial cells. NOTE: Yeast from the Ascomycetes phylum, such as <em>S. cerevisiae</em>, can reproduce asexually, which can be observed through the budding of the cells under a microscope.
</div></li><li data-sub-note-item="3-4"><div class="lab-step">Record any observations of yeast cells budding.  NOTE: Ascomycetes can also reproduce sexually via mitosis and haploid spore formation.
</div></li><li data-sub-note-item="3-5"><div class="lab-step">Record any observations of spore or asci formation you observed in the table.
</div></li><li data-sub-note-item="3-6"><div class="lab-step">NOTE: Fungus from the Basidiomycota phylum exhibit clearly observable gills under the pileus.
</div></li><li data-sub-note-item="3-7"><div class="lab-step">Record any fungal species that demonstrated gills under the pileus.
</div></li></ul></li></ol>

Microbial Colony and Fungal Diversity - Procedure

Diversité microbienne et fongique

Examining Microbial Colony Structures
ExpandTo begin, collect your allocated culture plates and then remove the Parafilm and lid from one of them.
Place ...

Community Diversity

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Measuring Biodiversity in Areas with Different Disturbance Regimes
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">NOTE: In this experiment, you will compare the biodiversity in a less disturbed area against a more highly disturbed area. HYPOTHESES: The experimental hypothesis could be that a less disturbed area will contain a greater diversity of species than a highly disturbed area. The null hypothesis could be that there will be no difference in diversity between less disturbed and highly disturbed areas. 
</div></li><li data-sub-note-item="1-2"><div class="lab-step">To begin, divide into groups of 2-4 and collect a hula hoop, ball, measuring tape, set of printed tables, 10 utility flags, and an ID guide from the front of the class.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Additionally, collect a shovel and 10 collecting bottles for the soil sampling exercise.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Once in the field, your instructor will show you the two habitats you will study - choose one habitat to begin.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Place one flag in each of four corners to form a rectangular sampling area.
</div></li><li data-sub-note-item="1-6"><div class="lab-step">Measure the length and width of the sampling area in meters, and then multiply the two numbers to determine the area.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Standing in the middle of the sampling area, throw a tennis ball into the air, and then Mark the location where it lands with a flag.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">Repeat this random throw four additional times to determine all five of your random sampling points.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">Keeping your flag at the center, place a hula hoop around it. The area within the hula hoop is a quadrat.
</div></li><li data-sub-note-item="1-10"><div class="lab-step">Identify all of the different species in the quadrat and record them as species A, B, and so on in Table 1, provided by your instructor.
</div></li><li data-sub-note-item="1-11"><div class="lab-step">Once all of the plants have been recorded, use a shovel to collect one cup of soil from each of the sites and put the samples bottles labeled with your names and the site name and quadrat number.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">Repeat the plant identification and soil sampling for the remaining four quadrats in the first habitat, recording your observations in Table 1.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">Once you have completed recording in the first habitat, move on to the second field, again marking out a rectangular sampling area.
</div></li><li data-sub-note-item="1-14"><div class="lab-step">Repeat the random placement of quadrats using the tennis ball, and then record the species present in each quadrat in Table 2. 
</div></li><li data-sub-note-item="1-15"><div class="lab-step">Finally, collect one cup of soil from each of the disturbed habitats&#39; quadrats, for a total of five cups from the disturbed and five cups from the undisturbed habitats.
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Measuring Biodiversity Along a Transect
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">NOTE: The next part of the study will test for edge effects, or a change along a gradient from where one ecosystem type meets another to the core habitat of the ecosystem. HYPOTHESES: The experimental hypothesis is that there will be a greater diversity of species closer to the edge of the ecosystem than in the center, or core, of the ecosystem. The null hypothesis is that there is no change in diversity along the transect from edge to core habitat.
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Place a flag at the perimeter of the field where it meets another habitat, like a forest or parking lot.
</div></li><li data-sub-note-item="2-3"><div class="lab-step">Extend a tape measure towards the center of the field, perpendicular to the edge of the field. This will be the transect.
</div></li><li data-sub-note-item="2-4"><div class="lab-step">Place a flag at the zero meter of the transect (the edge of the other habitat) then place additional flags at 10 m, 20 m, 30 m, and 40 m into the core of the habitat for a total of five flagged locations.
</div></li><li data-sub-note-item="2-5"><div class="lab-step">Starting at the 0 m point along the transect, place the hula hoop at the flagged location. Record the species present in the quadrat in Table 4.
</div></li><li data-sub-note-item="2-6"><div class="lab-step">Now repeat the quadrant placement and species recording at the 10 m, 20 m, 30 m, and 40 m marks.
</div></li><li data-sub-note-item="2-7"><div class="lab-step">Next, place two more transects from different starting points at the perimeter of the field extending into the core area.
</div></li><li data-sub-note-item="2-8"><div class="lab-step">Mark the 10-meter points along these transects, and again use these as markers to place quadrats. Record the species found in each of these quadrats in Table 4.
</div></li></ul></li><li data-note-item="3" data-parent-collapse="">Processing the Soil Samples
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="3-1"><div class="lab-step">HYPOTHESES: The experimental hypothesis is that the two landscapes under different disturbance regimes will differ in their soil composition and pH. The null hypothesis is that there will be no difference in the soil composition or pH between the two sites. 
</div></li><li data-sub-note-item="3-2"><div class="lab-step">To begin, remove soil from one bottle of the first site sampled and fill the pH color compartment of the soil sample kit to the soil fill line.
</div></li><li data-sub-note-item="3-3"><div class="lab-step">Pour the pH test capsule into the test chamber, then fill the capsule with distilled water until it reaches the water fill line.
</div></li><li data-sub-note-item="3-4"><div class="lab-step">Place the cap on the compartment and shake it.
</div></li><li data-sub-note-item="3-5"><div class="lab-step">Allow the color to develop for about a minute, and then compare the soil to the color chart to determine the pH.
</div></li><li data-sub-note-item="3-6"><div class="lab-step">Repeat the pH analysis for the second sampled site, filling out Table 3 for the recorded pH.
</div></li><li data-sub-note-item="3-7"><div class="lab-step">To process the soil samples for nitrogen, phosphorus, and potassium, remove 20 mL of soil from the bottle of the first site sampled.
</div></li><li data-sub-note-item="3-8"><div class="lab-step">Place the soil in a clean container with 100 mL of distilled water.
</div></li><li data-sub-note-item="3-9"><div class="lab-step">Repeat this process for the second site, labeling each container, and then allow them to sit for one hour.
</div></li><li data-sub-note-item="3-10"><div class="lab-step">After one hour, use a dropper to fill each of the nitrogen, phosphorous, and potassium comparators to the fill line with solution from the first sampled site.
</div></li><li data-sub-note-item="3-11"><div class="lab-step">Add the appropriate capsule to each of the comparators for each chemical test.
</div></li><li data-sub-note-item="3-12"><div class="lab-step">Cap and shake the comparators thoroughly, and then compare the solution in the test chamber to the color chart, recording the results as the appropriate number on the scale into Table 3.
</div></li><li data-sub-note-item="3-13"><div class="lab-step">Repeat the nitrogen, phosphorous, and potassium tests for each of the sampled sites. NOTE: Rinse the comparator thoroughly between samples
</div></li></ul></li><li data-note-item="4" data-parent-collapse="">Results
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="4-1"><div class="lab-step">To calculate the Shannon-Weiner Diversity for each of the observed sites, first open a new Excel spreadsheet.
</div></li><li data-sub-note-item="4-2"><div class="lab-step">Label cell A1 'species name', and cell B1 'total count'.
</div></li><li data-sub-note-item="4-3"><div class="lab-step">Starting from the second row and in column A, fill in the sheet for each species found in the first site, using the data recorded in Table 1.
</div></li><li data-sub-note-item="4-4"><div class="lab-step">Calculate the total number of each species found in the entire sampling effort and record this in the B column of the Excel spreadsheet.
</div></li><li data-sub-note-item="4-5"><div class="lab-step">Now, label cell G1 'total count of all species', and in cell H1, write a formula, or highlight the appropriate cells to sum all of the count values entered in column B: = SUM(B2: BX)
</div></li><li data-sub-note-item="4-6"><div class="lab-step">Label cell C1 'proportion of each species' and in cell C2, type the formula for the proportion of the total of each species at the site. = B2/H$1
</div></li><li data-sub-note-item="4-7"><div class="lab-step">Drag this formula down for each row of data represented in column B.
</div></li><li data-sub-note-item="4-8"><div class="lab-step">Label cell D1 'natural log proportion of species' and in cell D2, type the formula for the natural log of the proportion of species at the site. =LN(C2)
</div></li><li data-sub-note-item="4-9"><div class="lab-step">Drag this formula down for each row of data represented in column C.
</div></li><li data-sub-note-item="4-10"><div class="lab-step">Add the formula into cell E1 for the proportion of species multiplied by the natural log of the proportion of species: ps* ln ps
</div></li><li data-sub-note-item="4-11"><div class="lab-step">In cell E2, type the formula multiplying the proportion of species by the natural log of the proportion of species at each site, and then drag this formula down for each row of data represented in column D. = C2 * D2 
</div></li><li data-sub-note-item="4-12"><div class="lab-step">Label cell G2 as 'sum of ps * ln ps', and then, in cell H2, type a formula summing all of the values in column E. = SUM(E3 - EX) 
</div></li><li data-sub-note-item="4-13"><div class="lab-step">Label cell G3 'Shannon-Weiner Index', and in cell H3, enter the formula for the Shannon-Weiner Diversity Index. = (EXP - 1 * H2) NOTE: This is the calculated Shannon-Weiner Diversity Index for the first sampled site, and will be used for comparison between the two sites.
</div></li><li data-sub-note-item="4-14"><div class="lab-step">Repeat the data analysis in a new sheet using the data collected for the second site, which was recorded in Table 2. NOTE: Areas with greater index values are considered to have greater species diversity. Note whether you see differences between your sites.
</div></li><li data-sub-note-item="4-15"><div class="lab-step">Repeat the Shannon-Weiner Diversity calculations using a total count of all the species at each distance along the transect, then create a table of all of these values in a new Excel sheet.
</div></li><li data-sub-note-item="4-16"><div class="lab-step">Click on the Data tab in Excel, and then on the 'Data Analysis' button.
</div></li><li data-sub-note-item="4-17"><div class="lab-step">Select 'Regression' and click 'Okay'.
</div></li><li data-sub-note-item="4-18"><div class="lab-step">For the Input Y Range, highlight the Shannon-Weiner Diversity values, and for the Input X range, highlight the distance values.
</div></li><li data-sub-note-item="4-19"><div class="lab-step">Select an output location for the regression results and click 'Okay'.
</div></li><li data-sub-note-item="4-20"><div class="lab-step">Check the P-value box in the intercept row to see if there is a significant correlation between distance from edge to core habitat and biodiversity. NOTE: A value of less than 0.05 rejects the null hypothesis of no change in diversity along the transect, while a value greater than 0.05 does not reject the null hypothesis.
</div></li><li data-sub-note-item="4-21"><div class="lab-step">Next, to perform t-tests for determining soil composition differences between sites, open a new Excel spreadsheet.
</div></li><li data-sub-note-item="4-22"><div class="lab-step">In cell A1, type 'Site 1:pH', and in cell B1, type 'Site 2:pH'.
</div></li><li data-sub-note-item="4-23"><div class="lab-step">Fill in cells A2 through A6 and cells B2 through B6 with the soil pH collected in the disturbed and undisturbed sites, respectively.
</div></li><li data-sub-note-item="4-24"><div class="lab-step">Click the Data tab in Excel and then select 'Data Analysis'.
</div></li><li data-sub-note-item="4-25"><div class="lab-step">Now scroll down and select 't-Test: Two-Sample Assuming Equal Variances' and select 'Okay'.
</div></li><li data-sub-note-item="4-26"><div class="lab-step">Click the drop down next to Variable 1 Range and highlight cells A2 through A6 in the spreadsheet, then click the dropdown next to Variable 2 Range and highlight cells B2 through B6 in the spreadsheet.
</div></li><li data-sub-note-item="4-27"><div class="lab-step">In the box for hypothesized mean difference, type zero as our null hypothesis. This means that there is no difference between the two disturbance regimes.
</div></li><li data-sub-note-item="4-28"><div class="lab-step">Leave the Labels box unchecked and set the alpha value to 0.05.
</div></li><li data-sub-note-item="4-29"><div class="lab-step">Select 'Output Range', and in the box, click an empty space in the spreadsheet.
</div></li><li data-sub-note-item="4-30"><div class="lab-step">Click 'Okay'. A table of t-Test results should be outputted in the spreadsheet.
</div></li><li data-sub-note-item="4-31"><div class="lab-step">The relevant data are the means for each of the sites and the alpha value. If this second value is less than our alpha value of 0.05, we can reject the null hypothesis that there is no difference in pH between the two sites, and the hypothesis that disturbed versus undisturbed sites differ in their pH value would gain support. If this value is greater than 0.05, however, we may not reject our null hypothesis.
</div></li><li data-sub-note-item="4-32"><div class="lab-step">Repeat this analysis for each of the other three soil variables, creating a new spreadsheet for each of the phosphorus, potassium, and nitrogen elements. 
</div></li><li data-sub-note-item="4-33"><div class="lab-step">Note whether your pH and nutrient level results were different in each condition, and discuss with your group how this might affect the community and diversity in the different sites.
</div></li></ul></li></ol>

Community Diversity: Measuring Biodiversity in Different Areas - Procedure

Diversité communautaire

Measuring Biodiversity in Areas with Different Disturbance Regimes
ExpandNOTE: In this experiment, you will compare the biodiversity in a less disturbed ...

Ecologie

Cellular Respiration

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Quantifying Respiration using Microrespirometers
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">NOTE: In this experiment, you will measure the rate of cellular respiration for germinating seeds by measuring the rate of exchange for oxygen. As oxygen is consumed to provide energy, germinating seeds release carbon dioxide. This carbon dioxide is absorbed by potassium carbonate and thus the overall gaseous pressure of the respirometer will be reduced. HYPOTHESES: The experimental hypothesis is that germinating seeds will show a greater rate of respiration than control glass beads. Additionally, that at higher temperatures, the rate of cellular respiration in the seeds will increase. The null hypotheses are that both glass beads and germinating seeds will display a similar rate of cellular respiration and that temperature has no effect on respiration.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">To set up the control, first remove the plunger from a prepared microrespirometer and label it with a &#8220;CR&#8221; or &#8220;CH&#8221; respectively, depending on whether it is the control room temperature or heated treatment.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Add the glass microbeads to each of the microrespirometers up to the 0.5 mL mark, and push the plunger in the syringe to the 1 mL mark.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Place 3 - 4 washers onto the flared end of each microrespirometer to weigh them down while in the water bath.  NOTE: The microrespirometer will sit capillary tube end up.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Set the control devices to the side.
</div></li><li data-sub-note-item="1-6"><div class="lab-step">To set up the experimental seated respirometers, remove the plunger from a prepared microrespirometer and label it with &#8220;ER&#8221; or &#8220;EH&#8221; respectively, depending on whether it is the experimental room temperature or heated treatment.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Measure 0.5 mL of germinating seeds into the tuberculin syringe of the microrespirometer.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">Then, push the syringe to the 1 mL mark.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">Place 3 - 4 washers onto the flared end of each microrespirometer to weigh them down while in the water bath.
</div></li><li data-sub-note-item="1-10"><div class="lab-step">To use the microrespirometers, carefully press the &#8220;CR&#8221; and &#8220;ER&#8221; microrespirometers into the room temperature water bath. The entire syringe should be submerged, but the capillary tubes should be entirely exposed. Remove water from the bath until this position is achieved. Be sure the top of the capillary tube is open.
</div></li><li data-sub-note-item="1-11"><div class="lab-step">Perform the same procedure (step 9) for the &#8220;CH&#8221; and &#8220;EH&#8221; microrespirometers to set up the higher temperature experimental group.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">Wait 5 minutes before moving on to allow the temperature in the microrespirometers to equalize with the water bath.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">Add a single drop of manometer fluid to the top of each of the four capillary tubes which seals the microrespirometer chambers.  NOTE: If the chambers are working properly, the manometer fluid will be sucked into the capillary chamber.
</div></li><li data-sub-note-item="1-14"><div class="lab-step">Use the plunger set in the bath to suck the manometer fluid into the capillary. Stop when the fluid is about halfway down the capillary tube. IMPORTANT: The microrespirometers should not be disturbed once they are added to the bath. Do not bump the table or insert or remove anything to or from the water bath once the chambers have been sealed and equalized.
</div></li><li data-sub-note-item="1-15"><div class="lab-step">Mark the bottom edge of the manometer fluid that is closer to the chamber. This represents time point zero.
</div></li><li data-sub-note-item="1-16"><div class="lab-step">Set a timer to sound in 5 minutes.
</div></li><li data-sub-note-item="1-17"><div class="lab-step">After 5 minutes, create a new mark where the manometer fluid is on each microrespirometer. Repeat every 5 minutes until 25 minutes has passed or until the manometer fluid has traveled the entire length of the capillary tube.
</div></li><li data-sub-note-item="1-18"><div class="lab-step">Then, remove the respirometers from the water baths.
</div></li><li data-sub-note-item="1-19"><div class="lab-step">Measure the distance between each of the marks and record them in Table 1 next to the correct time point. <a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/8/Table_1_Cellular_respiration.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 1</a>  
</div></li><li data-sub-note-item="1-20"><div class="lab-step">During clean up, manometers should either be deconstructed to be reused or disposed of in the trash.
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Results
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">To analyze the data you collected, first plot the data points with the x-axis as time and the y-axis as the distance the manometer moved, representing oxygen consumption.
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Calculate the slope of each line using this equation.
</div></li><li data-sub-note-item="2-3"><div class="lab-step">Record these values for each microrespirometer in Table 2. NOTE: This value represents the amount of cellular respiration that took place over the course of the experiment.  <a href="https://www-jove-com.remotexs.ntu.edu.sg/CDNSource/lm/labs/8/Table_2_Cellular_respiration.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 2</a>
</div></li><li data-sub-note-item="2-4"><div class="lab-step">Plot the respiration rates as a bar graph.
</div></li><li data-sub-note-item="2-5"><div class="lab-step">Examine both of the graphs. Consider how your results fit with the hypotheses. Note any observed differences in respiration rate between the control beads and the germinating seeds. Determine if increasing the temperature had any effect on the respiration rate.
</div></li></ul></li></ol>

Cellular Respiration: Quantification using Microrespirometers - Procedure

Respiration cellulaire

Quantifying Respiration using Microrespirometers
ExpandNOTE: In this experiment, you will measure the rate of cellular respiration for germinating seeds ...