<p class='jove_content'>We thank Dr. Larry D. Frye for editorial help with this manuscript. This work was supported by Japan Society for the Promotion of Science Grants-in-Aid for Scientific Research 24300129 (to M.L.), 25890005 (to Y.O.) and 26640025 (to Y.T.), the National Agriculture and Food Research Organization (to Y.U.), the World Premier International Research Center Initiative (WPI) from the Ministry of Education, Culture, Sports, Science, and Technology (to Y.O., Y.T., Y.U. and M.L.) and the Nestl&#233; Nutrition Council, Japan (to M.L.).</p>

<p class="jove_content"> Déclaration éthique: procédures impliquant des sujets animaux ont été approuvés par la Commission institutionnelle Expérience animale de l&#39;Université de Tsukuba. </p><p class="jove_title"> 1. Préparation des électrodes et des câbles pour EEG / EMG Recordings </p><ol><li> Préparer EEG / EMG électrode d&#39;enregistrement selon le mode opératoire suivant. <br /> Remarque: L&#39;électrode est jetable et peut être utilisée que pour une animal. Planifiez soigneusement la configuration de câblage pour tous les connecteurs. Placer des marques sur les connecteurs pour l&#39;orientation correcte. <ol><li> Soudez chaque broche d&#39;une tête à 4 broches à un fil d&#39;acier inoxydable de 2 cm. En bref, tenir une extrémité du fil à la broche, placer un fer à souder chaud sur l&#39;articulation fil broches, et faire fondre la soudure pour assurer que juste assez de soudure fonctionne bien dans l&#39;articulation. Soyez prudent de ne pas appliquer trop de chaleur à la broche; autrement, la matière plastique autour des broches fondra. </li><li> Soudez l&#39;extrémité libre de chacun des 2 fils attachés à l&#39;en-tête de la broche à la têted&#39;une vis en acier inoxydable de 1,0 mm de diamètre. En bref, maintenir l&#39;extrémité libre du fil à fil en dessous de la tête de vis, placer un fer à souder chaud sur l&#39;articulation fil-vis, et faire fondre la soudure pour assurer que juste assez de soudure fonctionne bien dans l&#39;articulation. Les 2 fils avec vis servent électrodes d&#39;enregistrement EEG, tandis que les 2 fils sans vis servent EMG électrodes d&#39;enregistrement. </li><li> Utilisez des ciseaux pour dépouiller 1 mm de l&#39;isolant à l&#39;extrémité des électrodes EMG pour augmenter la qualité du signal EMG. </li><li> Couvrir complètement toutes les broches soudées avec de la colle époxy à l&#39;aide d&#39;une fine baguette de bois ou un cure-dent pour réduire le bruit électrique pendant les enregistrements EEG / EMG. </li></ol></li><li> Préparer un câble pour connecter l&#39;électrode avec la bague collectrice tel que décrit ci-dessous. Ce câble peut être réutilisé. <ol><li> Soudez chaque broche d&#39;un connecteur FFC / FPC 4 broches avec un fil d&#39;un câble plat 30 cm. En bref, maintenir l&#39;extrémité dénudée du fil à la broche, placer un fer à souder chaudsur le joint fil broches, et faire fondre la soudure pour assurer que juste assez de soudure fonctionne bien dans l&#39;articulation. <br /> Remarque: Choisissez la longueur du câble plat qui est approprié pour la hauteur de la cage de l&#39;animal expérimental utilisé pour les enregistrements EEG / EMG. </li><li> Solder sertir prises à une extrémité des fils sur l&#39;autre extrémité du câble plat. En bref, maintenez une prise de sertissage à l&#39;extrémité libre dénudée du fil, placer un fer à souder chaud sur le joint fil-socket, et faire fondre la soudure à veiller à ce que suffisamment de soudure tout se passe bien dans l&#39;articulation. </li><li> Insérez chaque Crimp Socket dans une position 4 sertir logement. </li><li> Couvrir complètement les douilles avec de la colle époxy sertissage en utilisant une fine baguette de bois ou un cure-dent. </li></ol></li></ol><p class="jove_title"> 2. implantation d&#39;électrodes dans le chef de la souris (Durée: env. 20 min) </p><ol><li> Stériliser tous les instruments chirurgicaux dans un stérilisateur à billes à chaud avant la chirurgie. Anesthésier une souris mâle (10 - âgé de 20 semaines, les 20 - 30 g)avec une injection intraperitoneale de pentobarbital (50 mg / kg). Après avoir vérifié que la souris est profondément anesthésié en pinçant un orteil, raser les cheveux sur la tête et le cou avec une tondeuse. </li><li> Déplacez la souris sur le cadre stéréotaxique, et fixer la tête entre les 2 bars de l&#39;oreille. Appliquez de la vaseline sur les yeux pour prévenir la sécheresse. Nettoyer la peau rasée avec de l&#39;alcool et de couper le long de la ligne médiane avec un scalpel pour exposer le crâne. Clip de la peau pour garder la zone chirurgicale ouverte. </li><li> En utilisant un couteau de carbure (taille de forage: 0,8 mm de diamètre), percer 2 trous dans le crâne, l&#39;un sur l&#39;aire corticale frontale (1 mm en avant de bregma, 1,5 mm latéralement de la ligne médiane) et l&#39;autre sur la zone pariétale ( 1 mm en avant de lambda, 1,5 mm latéralement à la ligne médiane) de l&#39;hémisphère droit, selon coordonnées stéréotaxiques de Paxinos et Franklin. <sup>23</sup> </li><li> En utilisant le tournevis d&#39;un bijoutier, en acier inoxydable de lieu d&#39;enregistrement EEG vis dans les trous et faire 2 - 2,5 tours pour chaque screw pour le positionnement péridurale sur le cortex. <br /> Note: Ne pas insérer des vis trop profondes pour éviter d&#39;endommager le tissu cérébral. Vérifier que les vis sont bien fixés sur le crâne. Ceci est important d&#39;avoir des signaux EEG stables pendant une longue période de plusieurs enregistrements (typiquement, plus de 1 mois). Vis ondulées produisent des artefacts EEG et peuvent se détacher avant la fin de l&#39;horaire expérimental. </li><li> Fixez l&#39;ensemble d&#39;électrode (cf. Section 1.1, broches tournées vers le haut) avec de la colle instantanée sur le crâne et couvrir avec du ciment dentaire. Assurez bilatéralement petits trous avec une pince dans les trapèzes (cou) muscles et insérer les fils en acier inoxydable qui servent d&#39;électrodes EMG dans les trous. Suturer la peau avec un fil de soie (d&#39;un diamètre de 0,1 mm) pour éviter l&#39;exposition du muscle. </li><li> Retirer la souris du cadre stéréotaxique. Administrer de l&#39;ampicilline (100 mg / kg) et le méloxicam (1 mg / kg) par voie intrapéritonéale à la souris pour éviter l&#39;infection bactérienne et de réduire la douleur post-chirurgicale, respectivement. Keep la souris sur un tapis de chaleur et de le surveiller jusqu&#39;à ce qu&#39;elle ait repris conscience suffisante pour maintenir décubitus sternal. Maison de la souris individuellement lors de la récupération pour éviter l&#39;élimination des électrodes par d&#39;autres animaux, et administrer meloxicam (1 mg / kg) par voie intrapéritonéale le premier jour après la chirurgie. </li></ol><p class="jove_title"> 3. Enregistrement et acquisition EEG / EMG données </p><ol><li> Après une période de récupération de 1 semaine, chaque maison souris individuellement dans une cage expérimentale placé dans une chambre d&#39;enregistrement insonorisé. Maintenir une température ambiante de 23 ± 1 ° C et contrôler automatiquement des cycles de 12 heures de lumière / 12-hr foncé (lumières allumées à 08h00, l&#39;intensité d&#39;éclairage ~ 100 lux). </li><li> Connectez l&#39;ensemble d&#39;électrodes EEG / EMG sur la tête de la souris à un câble d&#39;enregistrement. Vérifiez que le câble d&#39;enregistrement est relié à une bague collectrice (qui est conçu de telle sorte que les mouvements de la souris ne sont pas limités par la torsion du câble) et un amplificateur de signal EEG / EMG. Filtre EEG / EMG signaux (EEG, 0,5-64 Hz; EMG, 16-64 Hz), numériser à une cadence de 128 Hz d&#39;échantillonnage d&#39;un convertisseur analogique-numérique (A / D) et enfin enregistrer sur un ordinateur exécutant le logiciel d&#39;enregistrement EEG / EMG (Tableau &#39;Matériaux&#39;, 4 <sup>ème</sup> entrée à partir du bas). </li><li> Habituer la souris pour 2 - 3 jours dans la chambre d&#39;enregistrement. Si l&#39;enregistrement EEG / EMG comprend les administrations de drogue par voie intrapéritonéale, gérer doucement la souris sur chaque jour de l&#39;accoutumance au moment de l&#39;administration du médicament. </li><li> Par la suite, lancer le logiciel EEG / EMG d&#39;enregistrement (Tableau &#39;Matériaux&#39;, 4 <sup>ème</sup> entrée du bas). <ol><li> Sélectionnez l&#39;onglet «Fichier des données d&#39;informations» et cliquez sur la case à côté du nom de fichier. Entrez un nom de fichier et cliquez sur «Enregistrer». </li><li> Sélectionnez l&#39;onglet «condition d&#39;enregistrement» et sélectionnez tous les canaux EEG / EMG qui seront enregistrées. </li><li> Sélectionnez la fréquence d&#39;échantillonnage (128 Hz) dans l&#39;onglet «conditions d&#39;enregistrement». </li><li> Vérifiez si les canaux sélectionnés sont affichés correctement dans ee onglet «Canal de l&#39;information». </li><li> Sélectionnez l&#39;onglet &#39;de réglage de la minuterie &quot;et cliquez sur&quot; Monitor &quot;pour afficher EEG et EMG. </li><li> Vérifiez si les signaux EEG / EMG sont affichés correctement. </li><li> Sélectionnez l&#39;onglet &#39;de réglage de la minuterie &quot;et régler l&#39;heure de l&#39;horloge pour le début et la fin de l&#39;enregistrement dans la zone du minuteur principal. </li><li> Cliquez sur &quot;Moniteur&quot; de l&#39;onglet &quot;réglage de la minuterie» pour commencer l&#39;enregistrement. </li></ol></li><li> Enregistre des signaux EEG / EMG sous la ligne de base <em>(ie.,</em> Le comportement de sommeil / éveil d&#39;une souris de se comporter librement) et différentes conditions de traitement <em>(par</em> exemple., L&#39;administration de caféine ou un placebo) pendant plusieurs jours. Euthanasier la souris avec une injection intraperitoneale de pentobarbital (200 mg / kg) après le dernier jour d&#39;expérimentation. </li></ol><p class="jove_title"> 4. Scoring of Behavioral État Basé sur EEG / EMG données </p><ol><li> Démarrez le logiciel pour l&#39;analyse EEG / EMG (Tableau &#39;Matériaux&#39;, 4 <sup>ème</sup> entrée du bas). Ouvrez les données brutes EEG / EMG (fichier .kcd) produites dans l&#39;étape 3. Cliquez sur l&#39;onglet «sommeil» et sélectionnez le temps Epoch. Sélectionnez 10 sec. </li><li> Cliquez sur l&#39;onglet «sommeil» et sélectionnez «Multi-screening» de marquer automatiquement toutes les époques de 10 sec en 3 étapes <em>(ie.,</em> Sommeil lent et le sommeil paradoxal, et l&#39;éveil) sur la base des amplitudes des EEG et EMG et l&#39;analyse spectrale de puissance de l&#39;EEG. <sup>3</sup> </li><li> Cliquez sur &quot;l&#39;état de la FFT pour EEG». </li><li> Définissez les paramètres de l&#39;analyse spectrale de puissance [256 points de référence (correspondant à 2 secondes de l&#39;EEG), la fonction de fenêtre de Hanning, et la moyenne des 5 spectres par époque]. <sup>3 Cliquez sur</sup> «OK». </li><li> Cliquez sur «Démarrer screening» pour commencer le dépistage automatique. Ouvrez les données marqués (fichier .RAF). </li><li> Consultez les résultats de l&#39;examen automatique et, si nécessaire, corriger manuellement sur ​​la base des critères standard (voir les résultats représentatifs, <strong>figure 1B</strong> et <strong>tableau 1</strong> </strong&gt;). <sup>2, 3</sup> Brièvement, cliquez et maintenez le bouton gauche de la souris sur une époque mal marqué et faites glisser le curseur sur la chaîne des époques mal marqués. Relâchez le bouton gauche de la souris et sélectionnez l&#39;état comportemental correcte dans la fenêtre pop-up. <br /> Remarque: Parfois, les époques à la transition entre deux états vigilants sont difficiles à marquer sans ambiguïté, à un état. Dans de tels cas, l&#39;époque devrait être marqué à l&#39;état apparent et les mêmes critères doivent être appliqués à des époques similaires à travers l&#39;expérience pour assurer la reproductibilité des données. </li></ol>

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<li>Xu, Q., <em>et al.</em> <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+mouse+model+mimicking+human+first+night+effect+for+the+evaluation+of+hypnotics%5BTitle%5D)+AND+%22Pharmacol.+Biochem.+Behav%22%5BJournal%5D)">A mouse model mimicking human first night effect for the evaluation of hypnotics.</a> <em>Pharmacol. Biochem. Behav</em>. <strong>116</strong>, 129-136 (2014).</li>
<li>Cho, S., <em>et al.</em> <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Marine+polyphenol+phlorotannins+promote+non-rapid+eye+movement+sleep+in+mice+via+the+benzodiazepine+site+of+the+GABAA+receptor%5BTitle%5D)+AND+%22Psychopharmacol%22%5BJournal%5D)">Marine polyphenol phlorotannins promote non-rapid eye movement sleep in mice via the benzodiazepine site of the GABAA receptor.</a> <em>Psychopharmacol</em>. <strong>231</strong> (14), 2825-2837 (2014).</li>
<li>Liu, Y. Y., <em>et al.</em> <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Piromelatine+exerts+antinociceptive+effect+via+melatonin%2C+opioid%2C+and+5HT1A+receptors+and+hypnotic+effect+via+melatonin+receptors+in+a+mouse+model+of+neuropathic+pain%5BTitle%5D)+AND+%22Psychopharmacol%22%5BJournal%5D)">Piromelatine exerts antinociceptive effect via melatonin, opioid, and 5HT1A receptors and hypnotic effect via melatonin receptors in a mouse model of neuropathic pain.</a> <em>Psychopharmacol</em>. <strong>231</strong> (20), 3973-3985 (2014).</li>
<li>Qu, W. M., <em>et al.</em> <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Lipocalin-type+prostaglandin+D+synthase+produces+prostaglandin+D2+involved+in+regulation+of+physiological+sleep%5BTitle%5D)+AND+%22Proc.+Natl.+Acad.+Sci.+USA%22%5BJournal%5D)">Lipocalin-type prostaglandin D synthase produces prostaglandin D2 involved in regulation of physiological sleep.</a> <em>Proc. Natl. Acad. Sci. USA</em>. <strong>103</strong> (47), 17949-17954 (2006).</li>
</ol>

<p class='jove_content'>The authors Yujiro Tauguchi and Sayaka Kohtoh are employees of Kissei Comtech Co., Ltd that develops SleepSign software for acquisition and automatic scoring of EEG/EMG data used in this article.</p>

<p class="jove_content"> Ce protocole décrit un set-up pour les enregistrements EEG / EMG qui permet l&#39;évaluation de sommeil et l&#39;éveil sous faible bruit, conditions de coût-efficaces et à haut débit. En raison de la petite taille de l&#39;ensemble de tête de l&#39;électrode EEG / EMG, ce système peut être combiné avec d&#39;autres implants pour des expériences intra-cérébrales, y compris optogénétique (optique de l&#39;implantation de la fibre) ou, en conjonction avec simultanée canule implantation, microperfusion de m...

<p class="jove_content"> Les progrès techniques ont souvent précipité sauts quantiques dans la compréhension des processus neurobiologiques. Par exemple, la découverte de Hans Berger en 1929 que les potentiels électriques enregistrés à partir du cuir chevelu humain ont pris la forme d&#39;ondes sinusoïdales, dont la fréquence est directement liée au niveau de l&#39;éveil du sujet, a conduit à des progrès rapides dans la compréhension de veille-sommeil la réglementation, dans les deux animaux et les humains. <sup>1</sup> A ce jour l&#39;electroencephlogram (EEG), en collaboration avec l&#39;électromyogramme (EMG), <em>ie.,</em> l&#39;activité électrique produite par les muscles squelettiques, représente les données «colonne vertébrale» de presque tous expérimentale et clinique évaluation visant à établir une corrélation entre comportement et la physiologie de l&#39;activité des neurones corticaux en se comportant animaux, y compris les humains. Dans la plupart des laboratoires de recherche du sommeil de base, ces enregistrements EEG sont effectuées en utilisant un système à base de câble <strong>(Figure 1)</strong> dans laquelle acquis dATA est soumis hors ligne le modèle et le spectre d&#39;analyse <em>[par ex.,</em> l&#39;application de la transformée de Fourier rapide (FFT)] pour déterminer l&#39;état de l&#39;objet de la vigilance en cours d&#39;enregistrement. <sup>2, 3</sup> Sommeil se compose de mouvements oculaires rapides (REM) et non-REM (NREM) sommeil. Le sommeil paradoxal est caractérisé par une basse tension rapide EEG, le mouvement des yeux aléatoire, et atonie musculaire, un état dans lequel les muscles sont effectivement paralysés. Sommeil paradoxal est également connu comme le sommeil paradoxal, car l&#39;activité cérébrale ressemble à celle de l&#39;état de veille, tandis que le corps est en grande partie déconnecté du cerveau et semble être en sommeil profond. En revanche, les neurones sont stimulés au cours du sommeil lent, mais il n&#39;y a aucun mouvement de l&#39;oeil. Le sommeil de NREM humain peut être divisé en 4 étapes, de sorte que l&#39;étape 4 est appelé sommeil profond ou le sommeil lent et qui est identifié par les grandes ondes cérébrales lentes, avec une activité delta entre 0,5 à 4 Hz dans l&#39;EEG. D&#39;autre part, une subdivision entre les phases de sommeil lent dans de plus petits animaux, comme des rats unend souris, n&#39;a pas été établie, principalement parce qu&#39;ils ne disposent pas de longues périodes consolidés de sommeil comme on le voit chez les humains. </p><p class="jove_content"> Au fil des ans, et sur la base de l&#39;EEG interprétation, plusieurs modèles de régulation veille-sommeil, à la fois sur la base de circuits et humorale, ont été proposées. La base neuronale et cellulaire du besoin de sommeil ou, alternativement, «lecteur de sommeil,&quot; toujours pas résolue, mais a été conceptualisée comme une pression homéostatique qui construit pendant la période de la veille et est dissipée par le sommeil. Une théorie est que les facteurs endogènes somnogenic accumulent pendant l&#39;éveil et que leur accumulation progressive est le fondement de sommeil pression homéostatique. Alors que la première hypothèse formelle que le sommeil est régulé par des facteurs humoraux a été crédité au travail de Rosenbaum publiée en 1892 <sup>4,</sup> il était Ishimori <sup>5, 6 et</sup> Piéron <sup>7</sup> qui indépendamment, et il ya plus de 100 ans, a démontré l&#39;existence de produits chimiques favorisant le sommeil. Les deux chercheurs ont proposé, et en effet prouvé que les substances hypnogènes ou «hypnotoxins» étaient présents dans le liquide céphalo-rachidien (LCR) de chiens privés de sommeil. <sup>8</sup> cours du siècle passé plusieurs substances hypnogènes putatifs supplémentaires impliquées dans le processus homéostatique du sommeil ont été identifiés (pour revue, voir réf. 9), y compris la prostaglandine (PG) D <sub>2,</sub> <sup>10</sup> cytokines, <sup>11</sup> adénosine, <sup>12</sup> anandamide, <sup>13</sup> et le peptide urotensine II. <sup>14</sup> </p><p class="jove_content"> Les travaux expérimentaux par Economo <sup>15, 16,</sup> Moruzzi et Magoun <sup>17,</sup> et d&#39;autres dans les résultats précoces et mi-20 <sup>e siècle</sup> produites qui a inspiré les théories sur circuit de sommeil et l&#39;éveil et, dans une certaine mesure, éclipsé la théorie humorale alors en vigueur dormir. À ce jour, plusieurs «modèles de circuits» ont été proposées, chaque informés par des données de qualité et de quantité variable (pour revue, voir réf. 18). Un modèle, Par exemple, propose que le sommeil lent est généré par l&#39;inhibition de l&#39;adénosine-médiation de la libération d&#39;acétylcholine des neurones cholinergiques dans le cerveau antérieur basal, une zone consisiting principalement du noyau de la branche horizontale de la bande diagonale de Broca et l&#39;inominata substance. <sup>19</sup> Un autre modèle populaire de la réglementation de sommeil / éveil décrit un mécanisme de commutation bascule sur la base des interactions mutuellement inhibitrices entre les neurones somnifères dans la région préoptique ventrolatérale et les neurones de sillage induisant dans le tronc de l&#39;hypothalamus et le cerveau. <sup>18, 20, 21</sup> En outre, pour la commutation dans et hors de sommeil paradoxal, une interaction réciproque inhibiteur similaire a été proposé pour les zones dans le tronc cérébral, qui est le gris ventrale périaqueducale, latéral pontique calotte, et le noyau sublaterodorsal. <sup>22</sup> Collectivement, ces modèles se sont avérés précieux heuristiques et cadres d&#39;interprétation importantes offertes pour des études en recherche sur le sommeil; cependant, un yet meilleure compréhension des mécanismes et des circuits moléculaires régulant le cycle veille-sommeil, il faudra une connaissance plus complète de ses composants. Le système d&#39;enregistrement polygraphique détaillé ci-dessous devrait aider à atteindre cet objectif. </p>

<table><tbody><tr><td>4-pin header</td><td>Hirose</td><td>A3B-4PA-2DSA(71)</td><td></td></tr><tr><td>Ampicillin</td><td>Meiji Seika</td><td></td><td></td></tr><tr><td>Analog-to-digital converter</td><td>Contec</td><td>AD16-16U(PCIEV)</td><td></td></tr><tr><td>Caffeine</td><td>Sigma</td><td>C0750</td><td></td></tr><tr><td>Carbide cutter</td><td>Minitor</td><td>B1055</td><td></td></tr><tr><td>Crimp housing</td><td>Hirose</td><td>DF11-4DS-2C</td><td></td></tr><tr><td>Crimp socket</td><td>Hirose</td><td>DF11-30SC</td><td></td></tr><tr><td>Dental cement (Toughron Rebase)</td><td>Miki Chemical Product</td><td></td><td></td></tr><tr><td>Epoxy adhesive</td><td>Konishi</td><td>#16351</td><td></td></tr><tr><td>FFC/FPC connector</td><td>Honda Tsushin Kogyo</td><td>FFC-10BMEP1(B)</td><td></td></tr><tr><td>Flat cable</td><td>Hitachi Cable</td><td>20528-ST LF</td><td></td></tr><tr><td>Instant glue (Aron Alpha A)</td><td>Toagosei</td><td>N/A</td><td></td></tr><tr><td>Meloxicam</td><td>Boehringer Ingelheim</td><td>N/A</td><td></td></tr><tr><td>Pentobarbital</td><td>Kyoritsu Seiyaku</td><td>N/A</td><td></td></tr><tr><td>Signal amplifier</td><td>Biotex</td><td>N/A</td><td></td></tr><tr><td>Sleep recording chamber</td><td>APL</td><td>N/A</td><td></td></tr><tr><td>SleepSign software</td><td>Kissei Comtec</td><td>N/A</td><td>for EEG/EMG recording/analysis</td></tr><tr><td>Slip ring</td><td>Biotex</td><td>N/A</td><td></td></tr><tr><td>Stainless steel screw</td><td>Yamazaki</td><td>N/A</td><td>&amp;phi;1.0 &amp;times; 2.0</td></tr><tr><td>Stainless steel wire</td><td>Cooner Wire</td><td>AS633</td><td></td></tr></tbody></table>

polygraphic recording procedure for measuring sleep in mice

<p class='jove_content'>Recording of the epidural electroencephalogram (EEG) and electromyogram (EMG) in small animals, like mice and rats, has been pivotal to study the homeodynamics and circuitry of sleep-wake regulation. In many laboratories, a cable-based sleep recording system is used to monitor the EEG and EMG in freely behaving mice in combination with computer software for automatic scoring of the vigilance states on the basis of power spectrum analysis of EEG data. A description of this system is detailed herein. Steel screws are implanted over the frontal cortical area and the parietal area of 1 hemisphere for monitoring EEG signals. In addition, EMG activity is monitored by the bilateral placement of wires in both neck muscles. Non-rapid eye movement (Non-REM; NREM) sleep is characterized by large, slow brain waves with delta activity below 4 Hz in the EEG, whereas a shift from low-frequency delta activity to a rapid low-voltage EEG in the theta range between 6 and 10 Hz can be observed at the transition from NREM to REM sleep. By contrast, wakefulness is identified by low- to moderate-voltage brain waves in the EEG trace and significant EMG activity. </p>

<p class="jove_content" fo:keep-together.within-page="1"> <strong>Figure 1B</strong> illustre des exemples de l&#39;EEG souris dans les différents états de vigilance. Comme le montre le <strong>tableau 1,</strong> les époques sont classés comme si le sommeil lent EEG montre grandes ondes cérébrales, avec un rythme lent de delta inférieure à 4 Hz et l&#39;EMG ne dispose que d&#39;un signal faible ou nulle. Époques sont classés comme le sommeil paradoxal si l&#39;EEG montre rapides ondes cérébrales basse tension dans la gamme thêta e...

Watch this Scientific Journal Video about Polygraphic Recording Procedure for Measuring Sleep in Mice at JoVE.com

Polygraphic Recording Procedure for Measuring Sleep in Mice

<p class='jove_content'>The recording of electroencephalogram (EEG) and electromyogram (EMG) in freely behaving mice is a critical step to correlate behavior and physiology with sleep and wakefulness. The experimental protocol described herein provides a cable-based system for acquiring EEG and EMG recordings in mice.</p>

Procédure d&#39;enregistrement polygraphique pour la mesure du sommeil chez la souris

Recording of the epidural electroencephalogram (EEG) and electromyogram (EMG) in small animals, like mice and rats, has been pivotal to study the ...

polygraphic-recording-procedure-for-measuring-sleep-in-mice

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23.5K vues.  University of Tsukuba. The recording of electroencephalogram (EEG) and electromyogram (EMG) in freely behaving mice is a critical step to correlate behavior and physiology with sleep and wakefulness. The experimental protocol described herein provides a cable-based system for acquiring EEG and EMG recordings in mice.

Vidéo: Polygraphic Recording Procedure for Measuring Sleep in Mice

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>
					<td></td>
					<td></td>
					<td></td>
					<td></td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">5 min</td>
					<td></td>
					<td></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>
					<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>
					<td></td>
					<td></td>
				</tr>
			</tbody>
		</table>
		<a href="https://www.jove.com/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/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 ...

Enseignement

JoVE Lab Manual

Chimie

Lab Manual Sub Articles

Synthesis of Luminol

<p>Source: Lara Al Hariri and Ahmed Basabrain at the University of Massachusetts Amherst, MA, USA</p>
<ol class="note-items">
	<li data-note-item="1" data-parent-collapse="">Synthesis of 3-Nitrophthalhydrazide<button class="expand">Expand</button>
	<p>In this lab, you&#39;ll synthesize 3-aminophthalhydrazide, which is also called luminol, in a 2-step process. The first step is a condensation reaction between 3-nitrophthalic acid and hydrazine.</p>
	<p>During this step, the carboxylic acid groups are substituted with NH groups to produce 3-nitrophthalhydrazide and two water molecules. You must work in a fume hood throughout this lab because the reactions produce toxic vapors and gases.</p>
	<ul class="lab-items">
		<li data-sub-note-item="1-3">
		<div class="lab-step">Before you get started, put on a lab coat, splash-proof safety glasses, and nitrile gloves. <strong>Note:</strong> Hydrazine is highly toxic and flammable, so avoid touching it and keep anything containing hydrazine closed or covered when you transport it between fume hoods.</div>
		</li>
		<li data-sub-note-item="1-4">
		<div class="lab-step">Now measure 10 mL of deionized water and pour it into a 25-mL Erlenmeyer flask. Clamp the flask on a hot plate and place a thermometer in it.</div>
		</li>
		<li data-sub-note-item="1-5">
		<div class="lab-step">Then, start heating the water. You&#39;ll want the temperature of the water to be 80 &#176;C, so set the hot plate setting to slightly higher.</div>
		</li>
		<li data-sub-note-item="1-6">
		<div class="lab-step">While it heats, set up a Bunsen burner under a clamp fixed on a lab stand.</div>
		</li>
		<li data-sub-note-item="1-7">
		<div class="lab-step">Obtain a 25-mL test tube and adjust the clamp so that it will hold the test tube about 5 &#8211; 7 cm above the flame. Then, remove the test tube and place it in a 400-mL beaker.</div>
		</li>
		<li data-sub-note-item="1-8">
		<div class="lab-step">Once the water reaches 80 &#176;C, adjust the heat setting to keep the temperature stable.</div>
		</li>
		<li data-sub-note-item="1-9">
		<div class="lab-step">Then, weigh 0.6 g of 3-nitrophthalic acid in a tared weighing boat. Pour the 3-nitrophthalic acid into your test tube. Then, bring the test tube and a test tube stopper to the solvent hood.</div>
		</li>
		<li data-sub-note-item="1-10">
		<div class="lab-step">Use the provided volumetric pipette to carefully measure 0.9 mL of the 10% v/v hydrazine solution and transfer it to the test tube. Remember to cap the bottle of hydrazine solution and stopper your test tube before you return to your hood.</div>
		</li>
		<li data-sub-note-item="1-11">
		<div class="lab-step">Clamp the test tube over the Bunsen burner and remove the stopper. Then, pour ~2 mL of triethylene glycol into a graduated cylinder and set it out of the way in your hood.</div>
		</li>
		<li data-sub-note-item="1-12">
		<div class="lab-step">Now, add 2 or 3 boiling chips to the test tube. Place a high-temperature thermometer in the test tube and ignite the Bunsen burner. Heat the mixture with a medium flame until the 3-nitrophthalic acid dissolves.</div>
		</li>
		<li data-sub-note-item="1-13">
		<div class="lab-step">Then, add the triethylene glycol and increase the heat of the flame. <strong>Note:</strong> Once the water from the hydrazine solution has boiled off, the temperature will continue increasing.</div>
		</li>
		<li data-sub-note-item="1-14">
		<div class="lab-step">When the reaction mixture reaches 210 &#176;C, turn down the heat and keep the mixture between 210 &#176;C and 220 &#176;C for 3 &#8211; 4 min.</div>
		</li>
		<li data-sub-note-item="1-15">
		<div class="lab-step">Then, extinguish the flame and wait for the mixture to cool to 100 &#176;C.</div>
		</li>
		<li data-sub-note-item="1-16">
		<div class="lab-step">Turn off the hotplate, remove the thermometer, and unclamp the flask. Use tongs to hold the flask and measure 8 mL of hot water with the graduated cylinder. Pour this hot water into the test tube and remove the thermometer.</div>
		</li>
		<li data-sub-note-item="1-17">
		<div class="lab-step">Obtain a piece of plastic paraffin film and cover the mouth of the test tube. Then, use tongs to transfer the test tube to the holding beaker.</div>
		</li>
		<li data-sub-note-item="1-18">
		<div class="lab-step">At a lab sink, hold the test tube under running tap water to cool it, being careful not to let water into it.</div>
		</li>
		<li data-sub-note-item="1-19">
		<div class="lab-step">Then, dry the outside of the tube and bring it back to your hood. Clamp the tube upright and remove the paraffin film. Let the yellow 3-nitrophthalhydrazide precipitate in the tube for 15 min.</div>
		</li>
		<li data-sub-note-item="1-20">
		<div class="lab-step">While you wait, add ~10 mL of deionized water to a 25-mL Erlenmeyer flask. Prepare an ice bath in a 250-mL beaker and start chilling the water in it.</div>
		</li>
		<li data-sub-note-item="1-21">
		<div class="lab-step">Then, set up for vacuum filtration using a 250-mL filter flask, a B&#252;chner funnel, and a piece of circular filter paper.</div>
		</li>
		<li data-sub-note-item="1-22">
		<div class="lab-step">Once the product has precipitated for at least 15 min, obtain a Pasteur pipette and wet the filter paper with a few drops of cold water.</div>
		</li>
		<li data-sub-note-item="1-23">
		<div class="lab-step">Then, turn on the vacuum and pour the contents of the test tube into the B&#252;chner funnel. Rinse the remaining solid into the funnel with 1 &#8211; 2 mL of cold water.</div>
		</li>
		<li data-sub-note-item="1-24">
		<div class="lab-step">Remove the boiling chips with tweezers and rinse them with a few drops of cold water to finish collecting the product.</div>
		</li>
		<li data-sub-note-item="1-25">
		<div class="lab-step">Turn off the vacuum pump once no more liquid is dripping into the flask from the funnel.</div>
		</li>
	</ul>
	</li>
	<li data-note-item="2" data-parent-collapse="">Synthesis of 3-Aminopthalhydrazide<button class="expand">Expand</button>
	<p>In the second step of the luminol synthesis, you&#39;ll reduce the nitro group of 3-nitrophthalhydrazide using sodium dithionite in a basic solution of NaOH. This will produce 3-aminophthalhydrazide dianion.</p>
	<p>After the reduction, you&#39;ll add acetic acid to protonate the dianion, forming luminol. Sodium dithionite is highly reactive and degrades quickly, so you&#39;ll use a slight excess of it. After the reduction, you&#39;ll protonate the luminol dianion to decrease its solubility in water. Lastly, you&#39;ll precipitate and collect luminol as a yellow solid.</p>
	<ul class="lab-items">
		<li data-sub-note-item="2-3">
		<div class="lab-step">Place a clean 25-mL test tube in a beaker and use a clean spatula to transfer your 3-nitrophthalhydrazide to the test tube.</div>
		</li>
		<li data-sub-note-item="2-4">
		<div class="lab-step">Set the B&#252;chner funnel aside and clamp the test tube over the Bunsen burner. Leave the filter flask in place for later.</div>
		</li>
		<li data-sub-note-item="2-5">
		<div class="lab-step">Now, use a 10-mL graduated cylinder to obtain 3 mL of approximately 3 M NaOH. Pour the sodium hydroxide solution into the test tube and stir the mixture with a glass rod until the 3-nitrophthalhydrazide dissolves.</div>
		</li>
		<li data-sub-note-item="2-6">
		<div class="lab-step">Now, weigh 2 g of sodium dithionite in a tared weighing boat and add it to the test tube. Obtain 3 mL of deionized water and pour it into the tube.</div>
		</li>
		<li data-sub-note-item="2-7">
		<div class="lab-step">Stir the mixture well to dissolve as much of the solid as you can. Then, add a few clean boiling chips to the test tube and ignite the Bunsen burner.</div>
		</li>
		<li data-sub-note-item="2-8">
		<div class="lab-step">Heat the mixture to boiling over a medium flame and then let it boil for 5 min.</div>
		</li>
		<li data-sub-note-item="2-9">
		<div class="lab-step">While you wait, measure 1.5 mL of glacial acetic acid and bring it to your hood.</div>
		</li>
		<li data-sub-note-item="2-10">
		<div class="lab-step">Once the reaction mixture has boiled for 5 min, shut off the Bunsen burner and add the acetic acid to the tube. Let the mixture cool to room temperature, which usually takes 10 &#8211;15 minutes. Meanwhile, prepare an ice bath in a 600-mL beaker.</div>
		</li>
		<li data-sub-note-item="2-11">
		<div class="lab-step">Once the mixture has cooled, place it in the ice bath and wait 15 min for the luminol to precipitate.</div>
		</li>
		<li data-sub-note-item="2-12">
		<div class="lab-step">Reassemble the vacuum filtration setup using a clean B&#252;chner funnel and a new piece of filter paper, and start cooling another 10 mL of deionized water in an ice bath.</div>
		</li>
		<li data-sub-note-item="2-13">
		<div class="lab-step">Once the luminol has been in the ice bath for at least 15 min, wet the filter paper with cold water. Then, collect the solid luminol by vacuum filtration.</div>
		</li>
		<li data-sub-note-item="2-14">
		<div class="lab-step">Retrieve the boiling chips and rinse the luminol with more cold water. Once liquid stops dripping from the funnel, turn off the vacuum and transfer the luminol to a 50-mL beaker.</div>
		</li>
		<li data-sub-note-item="2-15">
		<div class="lab-step">Now, prepare the combined filtrate for disposal. First, label a 250-mL beaker &#8216;hydrazine and dithionite waste&#8217;. Transfer the filtrate to the beaker and rinse the flask with deionized water to remove residual hydrazine and dithionite.</div>
		</li>
		<li data-sub-note-item="2-16">
		<div class="lab-step">Next, dilute the filtrate with 10 mL of deionized water. Then, measure 10 mL of 1 M sodium carbonate and add it to the combined filtrates.</div>
		</li>
		<li data-sub-note-item="2-17">
		<div class="lab-step">Bring a 50-mL graduated cylinder to the waste hood. Measure 40 mL of 10% sodium hypochlorite and pour it into the filtrate mixture. Place the beaker on the hot plate, add a stir bar, and secure a thermometer in the beaker.</div>
		</li>
		<li data-sub-note-item="2-18">
		<div class="lab-step">Heat the mixture to 50 &#176;C while stirring. The mixture must stay at 50 &#176;C for 1 h to oxidize leftover hydrazine and dithionite, so make sure that the temperature is stable and set a timer for 1 h before starting the last part of the lab.</div>
		</li>
	</ul>
	</li>
	<li data-note-item="3" data-parent-collapse="">Chemiluminescence and&#160;Fluorescence<button class="expand">Expand</button>
	<p>For the last part of the lab, you&#39;ll mix some of the luminol you made with dimethyl sulfoxide over solid potassium hydroxide. The hydroxyl ions deprotonate the two amine groups. Then, ambient oxygen oxidizes the luminol dianion to 3-aminophthalate, or 3-APA, in an excited state. This unstable complex will quickly relax to the ground state, releasing energy as visible light. Since the excited 3-APA was the product of a chemical reaction, the emitted light is called chemiluminescence.</p>
	<p>Once you have seen the chemiluminescence from the oxidation reaction, you&#39;ll add fluorescein, a fluorescent molecule, to the mixture. Some excited 3-APA will transfer energy directly to fluorescein rather than emitting light, giving you a solution with two different light-emitting compounds.</p>
	<ul class="lab-items">
		<li data-sub-note-item="3-3">
		<div class="lab-step">Now, measure about 0.2 g of the luminol you made and place it in a 15-mL Erlenmeyer flask. It&#39;s OK if the solid is still damp.</div>
		</li>
		<li data-sub-note-item="3-4">
		<div class="lab-step">Obtain 5 &#8211; 7 pellets of potassium hydroxide and pour them into the flask. Then, bring the flask, a stopper, and a 10-mL graduated cylinder to the solvent hood.</div>
		</li>
		<li data-sub-note-item="3-5">
		<div class="lab-step">Measure 2 mL of DMSO, add it to the flask, and stopper it. Put the graduated cylinder in your fume hood and bring the flask to a dark room.</div>
		</li>
		<li data-sub-note-item="3-6">
		<div class="lab-step">Shake the stoppered flask vigorously until you see chemiluminescence and then put the flask down. The light fades quickly because both the oxidation and relaxation are fast processes.</div>
		</li>
		<li data-sub-note-item="3-7">
		<div class="lab-step">Shake the flask and let the light fade 6 &#8211; 10 more times without opening the flask. The light will be progressively dimmer as the oxygen in the flask is consumed. <strong>Note:</strong> To make it brighter, leave the flask open in your fume hood for a minute to replenish the oxygen.</div>
		</li>
		<li data-sub-note-item="3-8">
		<div class="lab-step">Close the flask, return to the dark room, and shake the flask again to see the effect.</div>
		</li>
		<li data-sub-note-item="3-9">
		<div class="lab-step">Now, measure 0.2 g of fluorescein dye and add it to your flask. Stopper the flask and return to the dark room. Shake the flask vigorously until the mixture is glowing brightly and then set it down to observe the difference in color from luminol alone.</div>
		</li>
		<li data-sub-note-item="3-10">
		<div class="lab-step">When you&#39;re finished, check on the filtrate. Once it has been at 50 &#176;C for at least 1 h, turn off the hotplate and wait for the beaker to cool to room temperature.</div>
		</li>
		<li data-sub-note-item="3-11">
		<div class="lab-step">Then, add about 10 mL of tap water to the filtrate and flush it down the drain.</div>
		</li>
		<li data-sub-note-item="3-12">
		<div class="lab-step">Dispose of the remaining chemical and glass waste in the appropriate containers, clean your glassware and tools, and put away your lab equipment.</div>
		</li>
	</ul>
	</li>
	<li data-note-item="4" data-parent-collapse="">Results<button class="expand">Expand</button>
	<p>Under your oxidation reaction conditions, excited 3-APA emits blue-green light in a broad range around 500 nm when it relaxes, and fluorescein is excited by absorbing light at 480 &#8211; 490 nm and 515 &#8211; 525 nm.</p>
	<p>When you add fluorescein to the luminol oxidation reaction mixture, excited 3-APA can transfer energy that fluorescein would absorb as light directly to a nearby fluorescein molecule in a special interaction called nonradiative energy transfer. The resulting excited fluorescein emits yellow-green light when it relaxes.</p>
	<p>Excited 3-APA is highly unstable, so if no fluorescein is close enough for nonradiative energy transfer, it will relax by emitting light as usual. Thus, a spectrum of your final glowing mixture would show contributions from both blue-green and yellow-green light.</p>
	<ul class="lab-items">
	</ul>
	</li>
</ol>

Synthesis of Luminol and Chemiluminescence - Procedure

Synthèse du Luminol

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

	Synthesis of 3-NitrophthalhydrazideExpand
	In this lab, ...

Buffers

<p>Source: Smaa Koraym at Johns Hopkins University, MD, USA</p>
<ol class="note-items">
	<li data-note-item="1" data-parent-collapse=""><strong>Preparing 50 mM NaH<sub>2</sub>PO<sub>4</sub> Buffer, pH 7</strong><button class="expand">Expand</button>
	<p>In the first part of this experiment, you will prepare a sodium phosphate solution buffered at pH 7.0. Monosodium phosphate is a weak acid with the conjugate base, disodium phosphate. Unadjusted monosodium phosphate solutions usually have a pH of about 4 - 6.</p>
	<p>Buffers are most effective close to their pKa, which is 6.8 to 7.2 for monosodium phosphate. So, you will use NaOH to push the equilibrium towards the conjugate base without altering the overall composition of the buffer equilibrium.</p>
	<ul class="lab-items">
		<li data-sub-note-item="1-3">
		<div class="lab-step">Before starting the experiment, calculate the mass of monosodium phosphate that you will need to make 200 mL of a 50 mM solution.
		<h4><strong>Table 1: Preparation of 50 mM NaH<sub>2</sub>PO<sub>4</sub> Buffer, pH 7.0</strong></h4>
		<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always" height="247" width="358">
			<tbody>
				<tr>
					<td colspan="2"><strong>Molar mass of NaH<sub>2</sub>PO<sub>4</sub> = 119.98 g/mol</strong></td>
				</tr>
				<tr>
					<td>Solution volume (mL)</td>
					<td style="text-align: center;">200</td>
				</tr>
				<tr>
					<td>Molarity (mM)</td>
					<td style="text-align: center;">50</td>
				</tr>
				<tr>
					<td>Moles of NaH<sub>2</sub>PO<sub>4</sub> (mol)</td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td>Mass needed (mg)</td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td>Initial volume of 1 M NaOH (mL)</td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td>Final volume of 1 M NaOH (mL)</td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td>Volume of NaOH used (mL)</td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td>Volume of DI water needed (mL)</td>
					<td style="text-align: center;"></td>
				</tr>
			</tbody>
		</table>
		<a href="https://www.jove.com/CDNSource/lm/labs/45/Table_1._Preparation_of_50_mM_NaH2PO4_Buffer,_pH_7.0.xlsx" target="_blank">Click Here to download Table 1</a></div>
		</li>
		<li data-sub-note-item="1-4">
		<div class="lab-step">To begin, put on the necessary personal protective equipment, including a lab coat, splash-resistant safety glasses, and gloves. <strong>Note: </strong>This section of the lab uses NaOH, which is corrosive and toxic. Use caution when pouring and transporting NaOH.</div>
		</li>
		<li data-sub-note-item="1-5">
		<div class="lab-step">Fill a 250-mL plastic wash bottle with deionized water.</div>
		</li>
		<li data-sub-note-item="1-6">
		<div class="lab-step">Label two 100-mL beakers, one for neutral aqueous waste and one for basic aqueous waste.</div>
		</li>
		<li data-sub-note-item="1-7">
		<div class="lab-step">Calibrate your pH meter using the provided buffers. Store the probe in its storage solution when you are done.</div>
		</li>
		<li data-sub-note-item="1-8">
		<div class="lab-step">Tare weighing paper, and use a clean spatula to measure out the required amount of monosodium phosphate according to your calculations. Record the exact amount of monosodium phosphate in your lab notebook. <strong>Note: </strong>Monosodium phosphate absorbs moisture from the air, so work quickly to obtain an accurate measurement and close the container tightly when you are done with it.</div>
		</li>
		<li data-sub-note-item="1-9">
		<div class="lab-step">Place the monosodium phosphate in a 400-mL beaker and clean the spatula with a laboratory wipe. Throw out the weighing paper and the laboratory wipe before returning to your fume hood.</div>
		</li>
		<li data-sub-note-item="1-10">
		<div class="lab-step">Use a graduated cylinder to measure out 175 mL of deionized water. Pour the water into the beaker of monosodium phosphate and add a magnetic stir bar.</div>
		</li>
		<li data-sub-note-item="1-11">
		<div class="lab-step">Stir the solution until the salt has dissolved completely and the solution appears homogeneous. This usually takes 2 - 3 min.</div>
		</li>
		<li data-sub-note-item="1-12">
		<div class="lab-step">Measure 15 mL of deionized water and pour it into the beaker. Continue stirring the solution until it appears homogeneous again (about 1 &#8211; 2 min). Turn off the stir motor.</div>
		</li>
		<li data-sub-note-item="1-13">
		<div class="lab-step">Rinse the pH probe with deionized water, and then clamp it in the solution with the sensor above the stir bar.</div>
		</li>
		<li data-sub-note-item="1-14">
		<div class="lab-step">Bring a 10-mL graduated cylinder and watch glass to the dispensing hood and measure out 10 mL of 1 M NaOH. Note the exact volume in the graduated cylinder.</div>
		</li>
		<li data-sub-note-item="1-15">
		<div class="lab-step">Cover your NaOH with the watch glass, and carefully bring it to your fume hood.</div>
		</li>
		<li data-sub-note-item="1-16">
		<div class="lab-step">Resume stirring the monosodium phosphate solution. While monitoring the pH reading, use a disposable pipette to slowly add NaOH to the stirring solution in a drop-wise manner. Once the buffer pH reaches 7.0, return any NaOH still in the pipette to the graduated cylinder.</div>
		</li>
		<li data-sub-note-item="1-17">
		<div class="lab-step">Calculate the volume of NaOH that you added to the buffer. Subtract that volume from 10 mL to determine how much deionized water you must add to your buffer to reach a total solution volume of 200 mL.</div>
		</li>
		<li data-sub-note-item="1-18">
		<div class="lab-step">Measure the deionized water with another 10-mL graduated cylinder and add it to the stirring solution to finish making the buffer.</div>
		</li>
		<li data-sub-note-item="1-19">
		<div class="lab-step">Rinse the pH sensor with deionized water, cover it with the storage solution filled cap, and unplug or turn off the probe.</div>
		</li>
		<li data-sub-note-item="1-20">
		<div class="lab-step">Label a 250-mL polyethylene bottle as &#39;50 mM monosodium phosphate buffer, pH 7.0&#39;. Retrieve the magnetic stir bar from the solution.</div>
		</li>
		<li data-sub-note-item="1-21">
		<div class="lab-step">Use a funnel to pour the buffer solution into the labeled bottle and cap it tightly.</div>
		</li>
	</ul>
	</li>
	<li data-note-item="2" data-parent-collapse=""><strong>Absorption Spectroscopy of Free and Bound Neutral Red</strong><button class="expand">Expand</button>
	<p>Buffers can be used to evaluate compounds at specific pH values. In this section, you will record the absorbance spectrum of the indicator neutral red in various buffers. The protonated form of neutral red is red, and the deprotonated form is yellow-orange, meaning that they absorb green and blue-violet light, respectively. Thus, the acidic and basic forms have distinct absorption wavelengths. Protein binding alters the properties of neutral red, changing both its absorbance and its pKa.</p>
	<p>After measuring the absorbance spectrum of free neutral red at several pH values, you will add riboflavin-binding protein, or RP, to the solutions and measure the absorbances again. Absorbance intensity is related to concentration, so you will use the spectra to determine the pKa of free and bound neutral red after the lab.</p>
	<ul class="lab-items">
		<li data-sub-note-item="2-3">
		<div class="lab-step">Draw the following table in your lab notebook. Number the cuvettes 1 through 10 and list the nine buffer pH values that you will use. The 10th cuvette will be a deionized water blank. <strong> Note: </strong>Always hold cuvettes by the textured sides and wipe the transparent sides just before putting the cuvette in the spectrophotometer. Remember to align the transparent sides with the beam of light in the spectrophotometer.
		<h4><strong>Table 2: Absorbance of Free and Bound Neutral Red</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>Cuvette #</strong></td>
					<td style="text-align: center;"><strong>Buffer pH </strong></td>
					<td style="text-align: center;"><strong>Abs. at &#955;<sub>max </sub>(Free NRH<sup>+</sup>)</strong></td>
					<td style="text-align: center;"><strong>Abs. at &#955;<sub>max</sub> (Bound NRH)</strong></td>
				</tr>
				<tr>
					<td style="text-align: center;">1</td>
					<td style="text-align: center;">5.0</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td style="text-align: center;">2</td>
					<td style="text-align: center;">5.5</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">3</td>
					<td style="text-align: center;">6.0</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">4</td>
					<td style="text-align: center;">6.5</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">5</td>
					<td style="text-align: center;">7.0</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">6</td>
					<td style="text-align: center;">7.5</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">7</td>
					<td style="text-align: center;">8.0</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">8</td>
					<td style="text-align: center;">8.5</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">9</td>
					<td style="text-align: center;">11</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td style="text-align: center;">10</td>
					<td style="text-align: center;">blank</td>
					<td></td>
					<td></td>
				</tr>
			</tbody>
		</table>
		<a href="https://www.jove.com/CDNSource/lm/labs/45/Table_2._Absorbance_of_Free_and_Bound_Neutral_Red.xlsx" target="_blank">Click Here to download Table 2</a></div>
		</li>
		<li data-sub-note-item="2-4">
		<div class="lab-step">Obtain ten 1.5-mL cuvettes and caps and label the caps 1 through 10 to match the table in your lab notebook.</div>
		</li>
		<li data-sub-note-item="2-5">
		<div class="lab-step">Label a 25-mL beaker as &#8216;DIH2O&#8217; and fill it with deionized water.</div>
		</li>
		<li data-sub-note-item="2-6">
		<div class="lab-step">Flush your neutral aqueous waste down the drain and relabel the beaker as &#39;aqueous buffer waste&#39;.</div>
		</li>
		<li data-sub-note-item="2-7">
		<div class="lab-step">Label a 400-mL beaker for used micropipette tips.</div>
		</li>
		<li data-sub-note-item="2-8">
		<div class="lab-step">Attach a tip to a 1-mL micropipette and use it to dispense 1000 &#181;L of deionized water into cuvette 10. This is your solvent blank.</div>
		</li>
		<li data-sub-note-item="2-9">
		<div class="lab-step">Place 925 &#181;L of your pH 7.0 monosodium phosphate buffer in cuvette 5.</div>
		</li>
		<li data-sub-note-item="2-10">
		<div class="lab-step">Bring the remaining empty cuvettes to the buffer table. Guided by the table in your lab notebook, dispense 925 &#181;L of each buffer into the appropriate cuvette using the labeled micropipettes. Be careful not to use the same pipette tip for different buffers.</div>
		</li>
		<li data-sub-note-item="2-11">
		<div class="lab-step">Once you have finished, bring the buffers back to your workbench. There, set a 200-&#181;L micropipette to dispense 75 &#181;L, and attach a tip to the micropipette.</div>
		</li>
		<li data-sub-note-item="2-12">
		<div class="lab-step">Dispense 75 &#181;L of neutral red into each of the nine buffer cuvettes. Invert each cuvette several times to thoroughly mix the solutions. <strong>Note: </strong>Replace the pipette tip if it touches a buffer.</div>
		</li>
		<li data-sub-note-item="2-13">
		<div class="lab-step">Once you have mixed neutral red with every buffer, take a picture, or write down the colors of the solutions.</div>
		</li>
		<li data-sub-note-item="2-14">
		<div class="lab-step">Turn on a hand-held spectrophotometer and allow the light source to warm up. Once it is ready, create a new experiment to measure absorbance versus wavelength for the free neutral red.</div>
		</li>
		<li data-sub-note-item="2-15">
		<div class="lab-step">Insert the cuvette of deionized water and acquire a spectrum of the deionized water. Set it as a solvent background or blank.</div>
		</li>
		<li data-sub-note-item="2-16">
		<div class="lab-step">Then, remove the blank and insert the cuvette of neutral red in the lowest pH buffer, which will have the highest concentration of protonated neutral red.</div>
		</li>
		<li data-sub-note-item="2-17">
		<div class="lab-step">Acquire a spectrum, which should show one strong peak. Identify the wavelength corresponding to the highest point of this peak, or the maximum. Record this wavelength in your lab notebook as lambda max for protonated free neutral red.</div>
		</li>
		<li data-sub-note-item="2-18">
		<div class="lab-step">Save the data and remove the cuvette. Record the absorbance in your notebook, then collect spectra for cuvettes 2 - 9 using the same procedure.</div>
		</li>
		<li data-sub-note-item="2-19">
		<div class="lab-step">Record the wavelength of the isosbestic point in your lab notebook. That is where all the spectra cross at a single wavelength. If a spectrum does not cross at that point, empty and clean the cuvette, prepare a fresh sample, and try again.</div>
		</li>
		<li data-sub-note-item="2-20">
		<div class="lab-step">Once you finish collecting spectra for all nine cuvettes, save and export the data.</div>
		</li>
		<li data-sub-note-item="2-21">
		<div class="lab-step">Create a new experiment to measure absorbance versus wavelength for the bound neutral red.</div>
		</li>
		<li data-sub-note-item="2-22">
		<div class="lab-step">Fit a new tip to the 200-&#181;L micropipette and add 75 &#181;L of riboflavin-binding protein solution to each cuvette, including the blank. Eject the tip, secure the cuvette caps, and invert the cuvette several times to mix the solutions.</div>
		</li>
		<li data-sub-note-item="2-23">
		<div class="lab-step">Insert cuvette 10 into the spectrophotometer and set it as the solvent blank.</div>
		</li>
		<li data-sub-note-item="2-24">
		<div class="lab-step">Acquire a spectrum of the lowest pH sample, and identify the wavelength corresponding to the maximum of the most intense peak. Record it in your lab notebook as the lambda max of protonated-bound neutral red.</div>
		</li>
		<li data-sub-note-item="2-25">
		<div class="lab-step">Acquire spectra for cuvettes 2 - 9 in the same way as you did for the free neutral red samples. Record the intensities at lambda max for protonated-bound neutral red in your table, along with the wavelength at the isosbestic point.</div>
		</li>
		<li data-sub-note-item="2-26">
		<div class="lab-step">When you are done, save your data, export it for later analysis, and shut down the spectrophotometer.</div>
		</li>
		<li data-sub-note-item="2-27">
		<div class="lab-step">Put away all equipment and dispose of your used pipette tips in an approved waste container or trash bin.</div>
		</li>
		<li data-sub-note-item="2-28">
		<div class="lab-step">Empty the cuvettes into the aqueous buffer waste beaker and rinse the cuvettes into the beaker with deionized water.</div>
		</li>
		<li data-sub-note-item="2-29">
		<div class="lab-step">Empty the excess NaOH into the basic waste and rinse the graduated cylinder with water.</div>
		</li>
		<li data-sub-note-item="2-30">
		<div class="lab-step">Flush the basic aqueous waste down the drain with copious tap water, along with the other aqueous waste.</div>
		</li>
		<li data-sub-note-item="2-31">
		<div class="lab-step">Wash your glassware, cuvette caps, and stir bar per your lab standard procedures. Clean your work surfaces with a damp paper towel and throw out used paper towels and lab wipes in the lab trash.</div>
		</li>
	</ul>
	</li>
	<li data-note-item="3" data-parent-collapse=""><strong>Results</strong><button class="expand">Expand</button>
	<p>Now, let&#39;s analyze our absorbance data to determine the pKa&#39;s of neutral red.</p>
	<h4><strong>Table 3: Determination of Neutral Red pKa&#39;s</strong></h4>
	<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
		<tbody>
			<tr>
				<td></td>
				<td>Free NRH<sup>+</sup></td>
				<td>Bound NRH<sup>+</sup></td>
			</tr>
			<tr>
				<td>&#160;&#955;<sub>max</sub> (nm)</td>
				<td></td>
				<td></td>
			</tr>
			<tr>
				<td>&#160;&#916;A (nm)</td>
				<td></td>
				<td></td>
			</tr>
			<tr>
				<td>&#160;Midpoint (nm)</td>
				<td></td>
				<td></td>
			</tr>
			<tr>
				<td>&#160;pKa</td>
				<td></td>
				<td></td>
			</tr>
		</tbody>
	</table>
	<p><a href="https://www.jove.com/CDNSource/lm/labs/45/Table_3._Determination_of_Neutral_Red_pKa.xlsx" target="_blank">Click Here to download Table 3</a></p>
	<ul class="lab-items">
		<li data-sub-note-item="3-2">
		<div class="lab-step">Plot both sets of intensity data with absorbance intensity at lambda max on the y-axis and pH on the x-axis, with the points joined by smooth lines.</div>
		</li>
		<li data-sub-note-item="3-3">
		<div class="lab-step">Calculate the difference between the starting and ending absorbance intensities for each data series.</div>
		</li>
		<li data-sub-note-item="3-4">
		<div class="lab-step">Determine the absorbance midway between the starting and ending absorbances for each series, or the absorbance midpoints.</div>
		</li>
		<li data-sub-note-item="3-5">
		<div class="lab-step">Find where the midpoint occurs on each line and identify the pH value at that point. These pH values are the pKa&#39;s of free and bound neutral red.</div>
		</li>
		<li data-sub-note-item="3-6">
		<div class="lab-step">Here, we see an increase in pKa of about one between free and bound neutral red, indicating that bound-protonated neutral red is a weaker acid than free protonated neutral red by an order of magnitude.</div>
		</li>
	</ul>
	</li>
</ol>

Buffers: Common Ion Effect, Henderson-Hasselbalch Equation, and Buffer Capacity - Procedure

Tampons

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

	Preparing 50 mM NaH2PO4 Buffer, pH 7Expand
	In the first part of this experiment, you will ...

Scientific Measurement and Lab Skills

Two-Dimensional Graphs, Curve Fitting, and Standard Deviation - Procedure

Mesure scientifique et compétences de laboratoire

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

	Measuring Diameter and CircumferenceExpand
	Error can be introduced in any experiment, even ...

Optimal Foraging

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Simulating Foraging with Artificial Habitats and Prey
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">NOTE: The foragers will hunt for prey represented by pinto beans in four buckets of rice with varying prey densities. Without knowing what these densities are, foragers must obtain as many prey items as possible in as little time as possible. 
</div></li><li data-sub-note-item="1-2"><div class="lab-step">HYPOTHESES: In this experiment, the experimental hypothesis could be that foragers will catch the most prey in the higher prey density bucket and also spend the most time foraging there.  The null hypothesis would then be that foragers will catch an equal number of prey in each bucket and that foragers will spend an equal amount of time foraging in each patch. 
</div></li><li data-sub-note-item="1-3"><div class="lab-step">To begin the foraging activity, record the names and role of each participant at the top of the student data collection sheet.  NOTE: Before starting the experiment, the forager is allowed to become familiar with the foraging process &#8211; the following steps are a practice round. 
</div></li><li data-sub-note-item="1-4"><div class="lab-step">The forager should kneel or sit in front of the average density bucket. 
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Once the timer says start, the forager may begin looking for the pinto beans representing the prey. The forager can spend as much time as they like foraging in this bucket. 
</div></li><li data-sub-note-item="1-6"><div class="lab-step">Once a prey item is located, the forager must signal that one has been found and place the prey in the cup next to the bucket.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Once they believe they have acquired most of the prey in the patch, they should tell the timer to stop the time and the practice round is over. 
</div></li><li data-sub-note-item="1-8"><div class="lab-step">To begin the simulation proper, the forager should sit or kneel in front of bucket A. 
</div></li><li data-sub-note-item="1-9"><div class="lab-step">The timer must be ready to start the stopwatch. 
</div></li><li data-sub-note-item="1-10"><div class="lab-step">The recorder should be ready to record the time each prey item is located, in addition to the time that the forager arrived at and left each bucket. 
</div></li><li data-sub-note-item="1-11"><div class="lab-step">When all parties are ready, the timer should start the stopwatch and say &#8220;start&#8221;,  and the forager should begin foraging. NOTE: The stopwatch will run through the rest of the round for buckets B and C.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">As the forager signals finding pinto beans and places them in the cup next to the bucket, the timer should tell the recorder the time.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">The recorder should then write the time in the student data collection sheet.  <a href="https://www.jove.com/CDNSource/lm/labs/34/Table_1_Optimal_foraging.xlsx" style="display:block;margin:20px;" target="_blank">Click Here to download Table 1</a>
</div></li><li data-sub-note-item="1-14"><div class="lab-step">Once the forager has decided to move on to the next bucket, they should tell the timer &#8220;done&#8221;, and begin moving to the next patch. 
</div></li><li data-sub-note-item="1-15"><div class="lab-step">The recorder should record the time the forager left and the time they arrived at bucket B in the student data collection sheet.
</div></li><li data-sub-note-item="1-16"><div class="lab-step">Repeat the same procedure for buckets B and C. Once the forager has completed the activity, it is time for the next forager to begin using the same procedure. 
</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">Before performing any data analysis, review the data collection sheet and compile your individual data for captured prey, foraging time, and travel time for each bucket in your student calculations table. <a href="https://www.jove.com/CDNSource/lm/labs/34/Table_2_Optimal_foraging.xlsx" style="display:block;margin:20px;" target="_blank">Click Here to download Table 2</a>
</div></li><li data-sub-note-item="2-2"><div class="lab-step">After this, calculate your individual capture rate for each bucket by dividing the number of prey captured by the sum of the foraging time and travel time and record these values in your table. Individual capture rate =  Number of prey captured / (foraging time + travel time)
</div></li><li data-sub-note-item="2-3"><div class="lab-step">To calculate your Giving-Up-Time or a GUT for each bucket, subtract the time that the last prey item was acquired from the time you left the bucket.  GUT = time forager left bucket  -  time last prey item acquired
</div></li><li data-sub-note-item="2-4"><div class="lab-step">Next, calculate the mean GUT for your foraging group and record the values in your table.
</div></li><li data-sub-note-item="2-5"><div class="lab-step">Finally, calculate your scaled GUT for each bucket by dividing the bucket GUT by your group&#39;s mean GUT. Scaled GUT =  Bucket GUT / Group mean GUT NOTE: The scaled GUT will allow class data to be pooled later on without differences in individual foragers skewing the data. 
</div></li><li data-sub-note-item="2-6"><div class="lab-step">The instructor should now compile the data into the classroom spreadsheet.  <a href="https://www.jove.com/CDNSource/lm/labs/34/Table_3_Optimal_foraging.xlsx" style="display:block;margin:20px;" target="_blank">Click Here to download Table 3</a>
</div></li><li data-sub-note-item="2-7"><div class="lab-step">Plot a bar graph of the average number of prey collected in each bucket. On the x-axis, A should represent the low prey density bucket, B should represent the medium prey density bucket, and C the high prey density bucket. Plot the y-axis as the number of prey items caught. 
</div></li><li data-sub-note-item="2-8"><div class="lab-step">Next, plot the graph of the class average scaled GUT, where the x-axis represents the buckets with varying levels of prey density and the y-axis shows the values for the average scaled GUT. 
</div></li><li data-sub-note-item="2-9"><div class="lab-step">Plot a third graph of the average time spent foraging in each bucket. 
</div></li><li data-sub-note-item="2-10"><div class="lab-step">Now review the Marginal Value Theorem (MVT) from the concepts video and examine the three graphs that you generated from your class data. Take note of whether the graphs conform to the predictions of MVT. 
</div></li></ul></li></ol>

Optimal Foraging: Foraging with Artificial Habitats and Prey - Procedure

Alimentation optimale

Simulating Foraging with Artificial Habitats and Prey
ExpandNOTE: The foragers will hunt for prey represented by pinto beans in four buckets of rice with ...

Biologie

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/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/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 ...

Processus cellulaires

Animal Behavior

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Making Choice Chambers
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">Before beginning the experiments, use one of the Sharpies from the lab table to trace a line around the circumference of the bottom of both bottles. One line should be 1 cm from the base of the bottle and the other should be 1.5 cm from the base of the bottle.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">Use a razor blade to make a small cut in each bottle at the mark. 
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Then, insert the scissors and carefully cut along the line until the bottoms of the bottles are removed completely.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Fit one bottle bottom into the other.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Tape the bottles together to make a secure chamber.
</div></li><li data-sub-note-item="1-6"><div class="lab-step">Use the razor blade and scissors to poke a hole into the center of the chamber. 
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Take a piece of tape and fold the edge onto itself to make a tab and place the tape over the hole.  Note: The tab helps with easy removal and reattachment of the tape to the chamber.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">Use the Sharpie to label one side of the chamber A and the other side B.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">Upon receiving the colony of <em>Drosophila</em>, perform a count to make sure there are 40 living individuals in the vial.
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Geotaxis
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">To perform a geotaxis assay, first attach a clean, empty bottle cap to each end of the choice chamber and turn the fly vial upside down onto its lid.
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Open the tape flap in the center of the choice chamber.
</div></li><li data-sub-note-item="2-3"><div class="lab-step">Insert the funnel into the hole and tap all of the flies into the chamber. HYPOTHESES: The alternate hypothesis is that more flies will be moving against gravity, or up, regardless of the orientation of the choice chamber. Alternatively, if the flies are positively geotactic, then more flies will move down, or with gravity. The null hypothesis is that the flies will have no preference with regard to gravity and will move randomly within the chamber. 
</div></li><li data-sub-note-item="2-4"><div class="lab-step">Once the <em>Drosophila</em> are in the chamber, close the tape flap, rotate the chamber so that it stands vertically with side A on the bottom, and start the timer.
</div></li><li data-sub-note-item="2-5"><div class="lab-step">Immediately, count the number of <em>Drosophila</em> that are moving up and the number that are moving down. Record these data for time point 0 in the table. 
</div></li><li data-sub-note-item="2-6"><div class="lab-step">After 30 seconds, rotate the chamber so that side B is on the bottom and count the number of <em>Drosophila</em> that are moving up or down. Record these data for time point 30 in the table.
</div></li><li data-sub-note-item="2-7"><div class="lab-step">After 60 seconds, rotate the chamber so that side A is on the bottom again and record the number of moving flies for time point 60.  <a href="https://www.jove.com/CDNSource/lm/labs/15/Table_1_Animal_behavior.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 1</a>
</div></li><li data-sub-note-item="2-8"><div class="lab-step">Hold the chamber over the fly vial and carefully open the cap to return all of the <em>Drosophila</em> back to their empty vial. 
</div></li></ul></li><li data-note-item="3" data-parent-collapse="">Phototaxis
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="3-1"><div class="lab-step">To perform a phototaxis analysis, first cover side A of the choice chamber with tin foil, taking care that the foil reaches to the exact center between the two sides. 
</div></li><li data-sub-note-item="3-2"><div class="lab-step">Ensure the caps are secured on both ends of the chamber.  HYPOTHESES: In this exercise, the alternative hypothesis is that more flies will be found in the light side of the choice chamber, if the flues are attracted to light, or on the dark side if they are repelled by light. The null hypothesis is that <em>Drosophila</em> will have no preference with regard to either stimulus. Therefore, equal numbers of flies will be found on either side of the choice chamber.
</div></li><li data-sub-note-item="3-3"><div class="lab-step">Open the tape flap in the center of the choice chamber, insert the funnel into the hole, and tap all 40 flies into the chamber as previously described.
</div></li><li data-sub-note-item="3-4"><div class="lab-step">Set the timer to sound an alarm at 10 minutes.
</div></li><li data-sub-note-item="3-5"><div class="lab-step">When the alarm sounds, stop the timer, and count the number of flies in the open side B of the chamber.
</div></li><li data-sub-note-item="3-6"><div class="lab-step">Subtract that number from the total number of flies added to the choice chamber to calculate the number of flies in the covered side A.
</div></li><li data-sub-note-item="3-7"><div class="lab-step">Record these data in the table for time point 10.  <a href="https://www.jove.com/CDNSource/lm/labs/15/Table_2_Animal_behavior.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 2</a>
</div></li><li data-sub-note-item="3-8"><div class="lab-step">Carefully, return all of the flies back to their empty vial.
</div></li></ul></li><li data-note-item="4" data-parent-collapse="">Chemotaxis
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="4-1"><div class="lab-step">HYPOTHESES: In this exercise, the alternative hypothesis is that more flies will be found on the side of the chamber with the apple cider vinegar if the flies are attracted to the odor. Or, if the flies are repelled by the vinegar, then more of them will be found on the side of the chamber presenting the odorless water. The null hypothesis is that <em>Drosophila</em> will have no preference with regard to either stimulus and will move randomly in the chamber.  Use an individual disposable plastic pipette to dispense two drops of the apple cider vinegar onto one cotton ball for side A of the chamber. 
</div></li><li data-sub-note-item="4-2"><div class="lab-step">Use an individual disposable plastic pipette to dispense two drops of the distilled water onto a cotton ball for side B of the chamber. 
</div></li><li data-sub-note-item="4-3"><div class="lab-step">Fold two pieces of tape over themselves to create individual pieces of double-sided tape and use the tape to attach each cotton ball to the inside of the single bottle cap.
</div></li><li data-sub-note-item="4-4"><div class="lab-step">Screw the bottle caps with the cotton balls onto their corresponding sides of the chamber.
</div></li><li data-sub-note-item="4-5"><div class="lab-step">Transfer all of the flies into the chamber and set the timer to sound an alarm after 10 minutes. 
</div></li><li data-sub-note-item="4-6"><div class="lab-step">When the alarm sounds, stop the timer and record the number of flies in each side of the chamber. <a href="https://www.jove.com/CDNSource/lm/labs/15/Table_3_Animal_behavior.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 3</a>
</div></li><li data-sub-note-item="4-7"><div class="lab-step">At the end of the final experiment, funnel the flies into a morgue tube containing 70% ethanol. 
</div></li><li data-sub-note-item="4-8"><div class="lab-step">Dismantle the choice chambers for recycling.  NOTE: Since all of the chemicals used in these experiments are lab safe, the liquids can be disposed of down the sink, the chambers rinsed, and the solids can be thrown away in the trash. 
</div></li></ul></li><li data-note-item="5" data-parent-collapse="">Results
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="5-1"><div class="lab-step">Gather all of the data collected for the geotaxis, phototaxis, and chemotaxis experiments.
</div></li><li data-sub-note-item="5-2"><div class="lab-step">Plot these values as bar graphs for different conditions and different time points.
</div></li><li data-sub-note-item="5-3"><div class="lab-step">Use these data to draw inferences about the behavior of <em>drosophila melanogaster</em>. 
</div></li></ul></li></ol>

Animal Behavior: Studying Taxis Behavior of Drosophila - Procedure

Comportement animal

Making Choice Chambers
ExpandBefore beginning the experiments, use one of the Sharpies from the lab table to trace a line around the circumference of the ...

Extinction

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Diversity and Extinction during the Paleogene Epoch
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">Begin this procedure by reading the extinction reading supplement that has been printed out for you. <br /><strong>NOTE:</strong> Alternatively, you may download the reading supplement at the link below.&nbsp; <a href="https://www.jove.com/CDNSource/lm/labs/24/Lab_20_Reading Supplement.docx" style="display:block;margin:20px;" target="_blank">Click here to download the extinction reading supplement</a>
</div></li><li data-sub-note-item="1-2"><div class="lab-step">Using the information contained in the extinction reading supplements pertaining to the taxa of interest and the Chicxulub impact event, make predictions for each taxon listed in Table 1. Consider whether diversity in each taxon increased, decreased, or stayed the same immediately following the Cretaceous-Paleogene boundary and later on in the Paleogene epoch. <a href="https://www.jove.com/CDNSource/lm/labs/24/Table_1_Extinction.xlsx" style="display:block;margin:20px;" target="_blank">Click Here to download Table 1</a>
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Then, fill out the reasons for each of your predictions in the Contributing Factors column. HYPOTHESES: The experimental hypothesis is that many of the different taxa will show changes in diversity after the Cretaceous-Paleogene event, and again at the end of the Paleogene, with some increasing in diversity, while others decrease. The null hypothesis is that there will be no changes in diversity between taxa following the Cretaceous-Paleogene event, or at the end of the Paleogene.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Gather diversity data by going to the Paleobiology Database: <a href="http://www.paleobiodb.org" target="_blank">www.paleobiodb.org</a>.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">In the center of the screen, click the yellow box labeled Download Data. Under the italicized heading, &#8220;What do you want to download?&#8221;, select Diversity over time on the left side and Comma-separated values on the right side.
</div></li><li data-sub-note-item="1-6"><div class="lab-step">Scroll down to the heading &#8220;Select by taxonomy&#8221;.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">In the first box labeled 'Taxon or taxa to include', type in the name of the taxon that data is being gathered on.  NOTE: Ensure the spelling is correct.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">Now, go to the dropdown menu right below that box labeled &#8220;Taxonomic resolution&#8221;, and select genera.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">Under the next heading, Select by time, type Maastrichtian into the first box and Chattian in the box directly below. Make sure the dropdown menu labeled Time rule is set to major and the Temporal resolution is set to stage.
</div></li><li data-sub-note-item="1-10"><div class="lab-step">Select &#8220;Assume extant taxa range through to present&#8221;. Scroll back up to the middle of the page and click download. A save window will open.
</div></li><li data-sub-note-item="1-11"><div class="lab-step">Save the .csv file to a safe and accessible place and rename it to make it easy to identify.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">Open the file and scroll down to the main dataset under Records.  NOTE: The data are arranged in reverse time order by geological stage. The second column, interval_name, gives the accepted name of the stage in question. The next two columns give the appropriate dates of the stage in millions of years before the present. The final column, number of occurrences, gives a total count of the genera extant during that geological stage as determined from occurrence of fossils.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">To more easily work with this data, copy and paste each column into a new sheet. Be sure to label this sheet with the taxon in question, the data and time the data were obtained, and the data source.
</div></li><li data-sub-note-item="1-14"><div class="lab-step">Repeat this process for each of the taxa in Table 1 (steps 4 &#8211; 12). NOTE: Data from multiple taxa can be kept on the same sheet, so long as they are kept distinct and labeled appropriately.
</div></li><li data-sub-note-item="1-15"><div class="lab-step">To create line plots for each of the taxa showing the change in diversity over time, select the max_ma column and the number of occurrences column simultaneously.
</div></li><li data-sub-note-item="1-16"><div class="lab-step">In Excel, select Insert from the top toolbar and find the section labeled Charts. Select the bottom center icon, Insert Scatter Chart.
</div></li><li data-sub-note-item="1-17"><div class="lab-step">Then, select the Scatter with Straight Lines and Markers and a graph should appear. NOTE: Since the axis of this graph is in units of millions of years ago, the axis is in the reverse direction of time when put in numerical order.
</div></li><li data-sub-note-item="1-18"><div class="lab-step">To fix this, click on the green plus sign in the top-right corner of the graph and hover over Axes. This should reveal a small black arrow. Click the arrow to open up a small menu and select More Options.
</div></li><li data-sub-note-item="1-19"><div class="lab-step">This will open the Axis Options menu. Under Axis Options, click the box next to Values in reverse order. NOTE: It is also possible to change the axis scales from this menu, which can be used to eliminate the excess time that is not being analyzed in this lab. Keep any changes consistent between different graphs.
</div></li><li data-sub-note-item="1-20"><div class="lab-step">Double-click on the title at the top of the graph and change it to the taxon being represented.
</div></li><li data-sub-note-item="1-21"><div class="lab-step">Make a separate graph for each of the taxa in Table 1. Keep them organized and in one location. A separate sheet for graphs is recommended.
</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">For each graph, identify the location of the Cretaceous-Paleogene boundary and draw a vertical line through the graph at this point.
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Using the information from the graphs, fill out Table 2. In some cases, it may be helpful to go back to the original dataset and compare actual values, particularly when they are small. <a href="https://www.jove.com/CDNSource/lm/labs/24/Table_2_Extinction.xlsx" style="display:block;margin:20px;" target="_blank">Click Here to download Table 2</a>
</div></li><li data-sub-note-item="2-3"><div class="lab-step">Compare these values to the predictions made in Table 1 and discuss if the original contributing factors are still likely candidates.
</div></li><li data-sub-note-item="2-4"><div class="lab-step">Using the dataset, calculate the percentage of total taxa that went extinct over the Cretaceous-Paleogene boundary using this equation.
</div></li><li data-sub-note-item="2-5"><div class="lab-step">There is an event in the middle of the Paleogene known as the Paleocene-Eocene Thermal Maximum about 55.5 million years ago. This represents a high point in temperature during the Paleogene epoch at approximately 8 &#176;C higher than today&#39;s average temperature. Does this point seem to have an effect on diversity for any of the taxa? If yes, then which ones?
</div></li></ul></li></ol>

Extinction: Diversity and Extinction during the Paleogene Epoch - Procedure

Diversity and Extinction during the Paleogene Epoch
ExpandBegin this procedure by reading the extinction reading supplement that has been printed out for ...

Measuring Biodiversity

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Quadrats and Transects
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">In this activity, you will be using quadrats along transect lines to take biodiversity measurements of the plant species in three distinct habitats near your classroom. Begin by splitting into groups. Four works best.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">Select one person in your group to be the data recorder. The other three will be the data collectors.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Next, take one quadrat, or hula hoop, per group and place it on one side of one transect, or rope, next to a knot to allow identification of the plant species encircled by your quadrat. HYPOTHESES: For this experiment, the alternative hypothesis is that the alpha diversity will be smaller than gamma, meaning that there will be differences in the species that are present in each different habitat. The null hypothesis for this activity is that the alpha and gamma will be equal, meaning that there will be no species diversity difference between the communities sampled.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Each data collector must survey and identify the different plant species or any morphologically distinct varieties within your group&#39;s quadrat, giving the name of each plant type that you identify to the data recorder. It may seem like there are only a couple of species present at first, but look closer. It may take several minutes to find them all. 
</div></li><li data-sub-note-item="1-5"><div class="lab-step">In addition to recording the species name, the data recorder must also write a short description in the table, so that each species can be identified again in subsequent quadrats. <a href="https://www.jove.com/CDNSource/lm/labs/25/Table_1_Measuring_biodiversity.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 1</a>
</div></li><li data-sub-note-item="1-6"><div class="lab-step">Once all of the species within a quadrat have been identified, pick up your hoop and move on to the next knot along the same transect.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Continue surveying and moving to the next knot until you have completed the entire rope transect.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">Once you have completed the rope, move to a transect in a different habitat, and begin identifying the different species within each quadrat along the new transect, until each group has identified plants along three lengths of rope, one in each habitat, for a total of 15 quadrats surveyed.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">When all of the species data have been collected, carry the transects and quadrats back to the classroom.
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Species-Accumulation Curves and Richness/Evenness 
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">In the next activity, you will be calculating and comparing the species richness and evenness in two distinct populations, represented here by two opaque bags containing beads of various colors. For this exercise, start by taking one paper bag of beads per group.
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Have one person reach into the bag and randomly pull out three beads without looking.
</div></li><li data-sub-note-item="2-3"><div class="lab-step">Write down the bead colors, as well as the total number of colors pulled from the bag in the table. <a href="https://www.jove.com/CDNSource/lm/labs/25/Table_2_Measuring_biodiversity.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 2</a>HYPOTHESES: In this activity, the alternative hypothesis states that species evenness in the two communities will be different, while the richness will be the same. The null hypothesis might be that species richness and evenness will be the same in both communities.
</div></li><li data-sub-note-item="2-4"><div class="lab-step">Place the beads back in the bag after their colors are recorded.
</div></li><li data-sub-note-item="2-5"><div class="lab-step">Then, shake the bag and have the next person in the group draw another set of three beads.
</div></li><li data-sub-note-item="2-6"><div class="lab-step">Record the colors and numbers of beads that they pulled in the table.
</div></li><li data-sub-note-item="2-7"><div class="lab-step">Record the colors in the second row and the cumulative number of colors drawn in the last column.
</div></li><li data-sub-note-item="2-8"><div class="lab-step">Continue to take turns drawing three beads from the bag until 20 samples have been drawn.
</div></li><li data-sub-note-item="2-9"><div class="lab-step">Next, switch bags with a group with a bag from the other community, and repeat the exercise of drawing 20 samples, recording the color data in the appropriate community column of the table.
</div></li><li data-sub-note-item="2-10"><div class="lab-step">When all of the data have been recorded, return the beads to the bag and return the bags to your instructor.
</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">To calculate the alpha diversity of the transect and quadrat data collected by your group, use the data your data recorder entered for each species and enter it into your diversity table. <a href="https://www.jove.com/CDNSource/lm/labs/25/Table_3_Measuring_biodiversity.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 3</a>
</div></li><li data-sub-note-item="3-2"><div class="lab-step">Then, fill out the alpha diversity for each habitat into the appropriate column in the table.
</div></li><li data-sub-note-item="3-3"><div class="lab-step">Use the species data from your quadrat and transect table and enter the beta diversity data into the appropriate data cells within the diversity table. NOTE: The beta diversity is the number of species that are unique between two habitats. In this video, there are three species in common between the two sites, but a total of six unique species, giving a beta diversity of six between site A and B.
</div></li><li data-sub-note-item="3-4"><div class="lab-step">To calculate the gamma diversity for your activity, use the data from your quadrat and transect table and enter the gamma diversity data into the appropriate data cell. NOTE: The gamma diversity is the number of species in all of the habitats within a single study. It is not the sum of all of the alpha diversities, because species that are common between habitats should be counted only once.
</div></li><li data-sub-note-item="3-5"><div class="lab-step">To construct species accumulation curves for both of the bead sampling communities, use your bead color selection data to plot the species by sampling effort. That is the number of total colors by sample in the graph for each bead bag community.
</div></li><li data-sub-note-item="3-6"><div class="lab-step">Then, estimate the asymptote of the plotted data by appearance. Predict the number of species you would expect to find in each community. Note any differences in appearance between the two curves. If it was easier to estimate the number of species in one of the communities, make a hypothesis to explain this observation.
</div></li><li data-sub-note-item="3-7"><div class="lab-step">To construct rank abundance curves for both of the bead sampling communities, use the color bead data to fill in the table for both community A and community B, listing the colors that were identified during the sampling in the color column, and adding the total number of beads of each color that was sampled in the abundance column. <a href="https://www.jove.com/CDNSource/lm/labs/25/Table_4_Measuring_biodiversity.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 4</a>
</div></li><li data-sub-note-item="3-8"><div class="lab-step">Now, assign a rank to each color in the rank column, designating the color of the highest abundance the highest rank, rank one, and the color with the lowest abundance the lowest rank, rank 10. If there are multiple colors with the same number, just assign them consecutive ranks.
</div></li><li data-sub-note-item="3-9"><div class="lab-step">Plot individual rank abundance curves for the number of individuals by species ranked for the most to least common in both community A and community B. Compare the steepness of the two rank curves. Compare the richness and evenness between the two communities. Identify the community that you consider more biodiverse. 
</div></li></ul></li></ol>

Mesurer la biodiversité

Quadrats and Transects
ExpandIn this activity, you will be using quadrats along transect lines to take biodiversity measurements of the plant species in ...

Scientific Method

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Observation and Background Research
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">NOTE: The first two steps in the scientific process involve studying and observing natural phenomena of interest.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">Observe the leaves and branches of the individual plant species. Each species comes from a different environment.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Compare each branch to one another and record your observations about any qualitative similarities or differences. <a target="__blank" style="display:block;padding:20px" href="https://www.jove.com/CDNSource/lm/labs/1/Table_A_Lab_1.xlsx">Click Here to download Table A</a>
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Use leaf clippers to cut the leaves from the branches.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Make additional qualitative observations about the leaves from each species.
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Hypothesis Formulation
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">Think of some general questions that you might have after observing the different plant species.
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Using these general questions, create a null and an experimental hypothesis.
</div></li></ul></li><li data-note-item="3" data-parent-collapse="">Experimentation and Data Collection
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="3-1"><div class="lab-step">Weigh and record the mass of each leaf. <a target="__blank" style="padding:20px;display:block;" href="https://www.jove.com/CDNSource/lm/labs/1/Table_B_Lab_1.xlsx">Click Here to download Table B</a>
</div></li><li data-sub-note-item="3-2"><div class="lab-step">Measure and record the length of each leaf from the petiole to the tip in the table.
</div></li><li data-sub-note-item="3-3"><div class="lab-step">Next, carefully trace each leaf outline onto a blank piece of computer paper.
</div></li><li data-sub-note-item="3-4"><div class="lab-step">Then, cut out and weigh and 4 x 4 cm square of the same paper, weigh the cutout tracing, and record these data in the table.
</div></li></ul></li><li data-note-item="4" data-parent-collapse="">Data Analysis and Conclusion
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="4-1"><div class="lab-step">To calculate the leaf surface area, divide the surface area of the 4 x 4 cm paper square by the mass of the paper square, and multiply the resulting number by the mass of the leaf tracing.
</div></li><li data-sub-note-item="4-2"><div class="lab-step">Next, create a bar graph with your five plant species listed on the x-axis, and the surface area in square centimeters indicated on the y-axis.
</div></li><li data-sub-note-item="4-3"><div class="lab-step">Draw a trend line through the graph if a relationship is evident.
</div></li><li data-sub-note-item="4-4"><div class="lab-step">Using your qualitative observations and quantitative measurements, draw conclusions about the data you collected.
</div></li></ul></li></ol>

Procédure d'enregistrement polygraphique pour la mesure du sommeil chez la souris

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