Name: a step-by-step guide to mosquito electroantennography
Uploaded: 2021-03-10T14:30:00.000+00:00
Duration: 6 min 39 s
Description: Watch this Scientific Journal Video about A Step-by-Step Guide to Mosquito Electroantennography at JoVE.com

<p class="jove_content">I am grateful to Dr. Cl&#233;ment Vinauger and Dr. Jeffrey Riffell for helpful discussions. The following reagents were obtained through BEI Resources, NIAID, NIH: <em>Anopheles stephensi</em>, Strain STE2, MRA-128, contributed by Mark Q. Benedict; <em>Aedes aegypti</em>, Strain ROCK, MRA-734, contributed by David W. Severson; <em>Culex quinquefasciatus</em>, Strain JHB, Eggs, NR-43025. The author thanks Dr. Jake Tu, Dr. Nisha Duggal, Dr. James Weger and Jeffrey Marano for providing <em>Culex quinquefasciatus</em> and <em>Anopheles stephensi</em> (strain: Liston) mosquito eggs. <em>Aedes albopictus</em> and <em>Toxorhynchites rutilus septentrionalis</em> are derived from field mosquitoes collected by the author in the New River Valley area (VA, USA). This work was supported by The Department of Biochemistry and The Fralin Life Science Institute.</p>

<p class="jove_title">1. Saline solution preparation</p>
<ol>
	<li>Prepare the saline in advance and store in the fridge.</li>
	<li>Follow Beyenbach and Masia<sup class="xref">26</sup> to prepare the solution.<br />
	&#8203;NOTE: Saline recipe in mM: 150.0 NaCl, 25.0 HEPES, 5.0 glucose, 3.4 KCl, 1.8 NaHCO<sub>3</sub>, 1.7 CaCl<sub>2</sub>, and 1.0 MgCl<sub>2</sub>. The pH is adjusted to 7.1 with 1 M NaOH. Do not add glucose or sucrose to the preparation at this time to increase shelf storage. Add the needed quantity to the saline right before running the EAGs (around 50 mL per experiment).</li>
</ol>
<p class="jove_title">2. Odor preparation and storage</p>
<ol>
	<li>Prepare the odorant mixtures or single compound dilutions in advance in 1.5 mL amber vials and store at -20 &#176;C to prevent compound degradation.<br />
	NOTE: Concentrations will depend on the test to be conducted. 0.1% or 1% are commonly used to determine whether a compound can be detected or not. For a dose-response curve, prepare serial dilutions of a given chemical and test them from that the lowest to the highest concentrations.</li>
	<li>Prepare the dilutions in water, ethanol, hexane, paraffin oil, or mineral oil, depending on the solubility of the tested chemical.</li>
	<li>Make sure to prepare a solvent control (a vial containing only the solvent) for the experiment.</li>
	<li>Remove the odorants from the freezer 30 min prior to starting the experiments to allow them to thaw. Vortex each vial prior to use to mix well the chemical and the solvent.</li>
	<li>Pipette 10 &#181;L of solution on to a piece of filter paper (0.5 cm x 2 cm) loaded inside a labeled glass syringe or Pasteur pipette.</li>
	<li>Load each compound or mixture in a specific Pasteur pipette or syringe to prevent contamination.<br />
	NOTE: Load 10 min before starting the experiment so the odor can diffuse in the syringe but not for longer to prevent degradation. Let the Pasteur pipette or syringe remain capped at this time to allow a good diffusion of the chemical before the experiment starts.</li>
	<li>After each EAG run, dispose the piece of the filter paper and replace it with a new one to prevent the paper from getting oversoaked and risking needle blockage. Replace the needles regularly (every 10 runs).</li>
</ol>
<p class="jove_title">3. Mosquito separation</p>
<ol>
	<li>Isolate the mosquitoes on the day of the experiments.</li>
	<li>Use mosquitoes that are at least 6 days old on the day of the experiments to increase the chances that the females are mated to enhance their response to host-related odorants.<br />
	NOTE: Adjust mosquito age at the time of the test depending on the project. Check and harmonize the physiological status (e.g., blood-fed, starved, never previously fed, etc.).</li>
	<li>Starve the mosquitoes up to 12 h (i.e., no access to sugar) to increase their motivation and sensitivity.</li>
	<li>Place the mosquito container in the fridge (4 &#176;C) until they stop flying so individuals can easily be delicately transferred to single cups with forceps.<br />
	NOTE: Species with a higher tolerance to cold can be put down using a CO<sub>2</sub> fly pad. Ensure that the mosquitoes do not stay on it for a long time to prevent desiccation, which would decrease the responsiveness of the mosquito EAG preparation.</li>
	<li>Store the cups containing single mosquitoes at room temperature before the EAGs are performed and discard any mosquitoes that may not be used during the day.</li>
</ol>
<p class="jove_title">4. Electrode holder and capillary preparation</p>
<ol>
	<li>Capillary pulling, preparation, and storage
	<ol>
		<li>Use borosilicate capillaries with filaments (I.D: 0.78 mm, O.D: 1 mm). Pull them depending on the equipment<sup class="xref">27</sup>.<br />
		NOTE: Store the pulled capillaries in a Petri dish. Place the Petri dish on to pieces of wax or unscented modeling clay to prevent them from moving and breaking.</li>
		<li>Before running the EAG experiment, gently break the tip of 2 capillaries with a pair of forceps under the microscope.<br />
		NOTE: Ensure that one is slightly bigger than the other to fit either the neck (larger capillary) or the tips of the antennae (smaller capillary). Ensure that the cut is clean with no crack present on the capillary wall. This requires patience and practice.</li>
		<li>If still intact, reuse these capillaries after rinsing with de-ionized (DI) water after the experiment is over. Remove the excess water by gently applying a cleaning wipe against the tip. Place back in storage Petri dish. If the tip is crooked, discard the capillary.</li>
	</ol>
	</li>
	<li>Electrode holder and capillary mounting
	<ol>
		<li>Label the two electrode holders as &#34;recording&#34; and &#34;reference&#34; by using pieces of lab tape of different colors. This will help guide the mounting of the mosquito head and electrodes.</li>
		<li>Ensure that the electrode holders are clear inside and that no borosilicate debris are present.</li>
		<li>Chloridization: Soak the silver wires of the electrode holders in pure bleach for about 5 min. The wires turn from shiny light grey to matte dark grey.</li>
		<li>Loosen the rubber stopper and fill the inside of the capillary with 10% saline solution using a 20 G needle.</li>
		<li>Fill the borosilicate capillary with the saline solution using a syringe. Ensure that no bubbles are present in neither the electrode holder nor the pulled capillary.<br />
		&#8203;NOTE: To reduce the chances of having bubbles in the capillary, keep pushing saline in the capillary while gently pulling the needle out and use capillaries with a filament. It is possible to load the capillaries with a solution composed of 1:3 electrode gel and saline solution. This can help prevent evaporation of the saline and can be particularly useful when learning and practicing EAGs, as the experimenter will need more time to complete the different steps.</li>
		<li>After soaking, rinse the silver wires with DI water and insert them in the two capillaries. Ensure that the tip of the wire is less than 1 mm from the tip of the capillary. Ensure that the capillary passes the rubber ring inside the electrode holder without breaking. Gently tighten the rubber stopper. Verify that no air bubbles are present.</li>
		<li>Use the capillary with the wider opening on the reference electrode holder (neck), and the smaller opening on the recording electrode holder (antennae).</li>
		<li>Leave the two mounted electrode holders on a wet cleaning wipe to prevent the tip from drying out until ready to mount the head.</li>
	</ol>
	</li>
</ol>
<p class="jove_title">5. EAG rig preparation (<strong class="xfig">Figure 1</strong>)</p>
<ol>
	<li>Ensure that the air table is up, that there is no blockage in the airline. Ensure that the tank of medical air is still full to avoid changing it in the middle of the experiment. Make sure that there are bubbles in the humidifier.</li>
	<li>Air and pulse delivery system
	<ol>
		<li>Turn on the medical air gas tank.</li>
		<li>Check the level of the two flowmeters.<br />
		&#8203;NOTE: The flowmeter controlling the main air current bathing the preparation during the whole experiment should be at 140 mL/min and the other related to the odor pulse should read 15 mL/min.</li>
	</ol>
	</li>
	<li>If doing GC-EAD, turn on the machine, gas tanks and create/load the file/method.</li>
	<li>Turn on the computers, the software applications, valve power supply, and verify the internet connection for the software application to work.
	<ol>
		<li>Software application: A short script can be written to deliver the pulse.</li>
		<li>EAG software: use any electrophysiology software.</li>
		<li>Implement the parameters in the software (e.g., amplifier, duration of recording, duration of the pulses, etc.).</li>
	</ol>
	</li>
	<li>Deliver a control pulse to verify that the valve delivering the pulses is functional.</li>
	<li>Set the power supply at 5.2 V. Verify the amplifier parameters.<br />
	NOTE: The parameters used for the data presented here are: low cut-off filter of 0.1 Hz; high cut-off filter of 500 Hz; Gain of x100.</li>
</ol>
<p class="jove_title">6. Mosquito head preparation and mounting (<strong class="xfig">Figure 2</strong>)</p>
<ol>
	<li>Place an aluminum plate on ice and place a piece of wet cleaning wipe over it.</li>
	<li>Put a small dollop of electrode gel in a corner.</li>
	<li>Place a mosquito cup on ice and let the mosquito cool down for a couple of minutes, or until it is stops flying.<br />
	NOTE: Some species are cold-resistant and might require a quick anesthesia over a CO<sub>2</sub> fly pad to go down. The least the mosquito stays on either, the better.</li>
	<li>Place the mosquito on its back and clip the tip of each antenna (only a small portion of the last segment) with micro scissors.</li>
	<li>Use forceps to drag the mosquito next to the electrode gel dollop and dip each antenna&#39;s tip gently in the gel. Avoid dipping more than just the very last segment in the electrode gel.</li>
	<li>Using forceps, pull the mosquito antennae out while maintaining them next to each other. Let them come out together from the gel. Make sure that the antennae do not touch the cleaning wipe&#39;s surface, or they might separate.</li>
	<li>Place the mosquito on its side and chop the head using micro scissors or a razor blade.<br />
	NOTE: Once the head is chopped, proceed quickly to the next steps and to the EAG rig to start the recordings. The preparation should stay responsive for about 30 min.</li>
	<li>Take the reference electrode and gently deep the tip in the gel. Keep in contact with neck tissues and let the head stick on it.</li>
	<li>Move the electrode holders under the EAG microscope and view through the microscope to place the head (i.e., reference) electrode on a micromanipulator. Ensure that the antennae are at the center.</li>
	<li>Grab the recording electrode, place it in front of the antennae tips. Move and align it as close as possible to the tips using the micromanipulator. Using the microscope, move the tip of the recording electrode towards the antennae.</li>
	<li>Connect both electrode holders to the amplifier before inserting the tips to prevent them from moving after insertion.</li>
	<li>Insert the antennae tips in the recording electrode. Ensure that they just contact the saline and the electrode gel and are visible by transparency through the capillary. The antenna goes in by the &#34;suction effect&#34;.</li>
	<li>Adjust the position of the head and tips with forceps under the microscope, if needed.</li>
	<li>Place the airline tubing close to the mosquito head preparation (distance: 1 cm).<br />
	NOTE: If the head falls off, return to the dissection station and remount the head or prepare a new one if it has been lost or if it has been more than 5 min since the head was cut off. A good connection between the capillary and the neck/antennae is essential for low noise and reliable recording. Ideally, the antennae tips will be at less than 1 mm from the wire of the recording electrode once inserted.</li>
	<li>Turn off the light source, if used.</li>
	<li>Position the vacuum line near the mosquito head preparation (distance: 20 cm) and align with the main airline.<br />
	&#8203;NOTE: The vacuum will help remove chemicals surrounding the head preparation after the stimulus, which could lead to EAG responses after the pulses were applied.</li>
</ol>
<p class="jove_title">7. Recordings</p>
<ol>
	<li>After insertion of the tips of the antennae, turn on the amplifier and the noise reducer. Observe the baseline signal and ensure that it is not noisy.<br />
	NOTE: Observe whether large oscillations in the electrical signal are present. Adjust the position of the head and antennae tips as needed until the signal is clean. Use alligator clips to ground anything that introduces noise to the Faraday cage or air table. A baseline signal of less than 0.01 mV in amplitude is ideal for detecting and discriminating minute EAG responses.</li>
	<li>Once the noise level is satisfying, insert the first odor syringe to test in the airline hole.</li>
	<li>Close the Faraday cage. Do not stay in front of the preparation, to reduce noise.</li>
	<li>Click on Record on EAG software.</li>
	<li>Deliver the pulse(s) using the software application.<br />
	NOTE: The number and duration of the pulses will vary depending on the experiments. Here, single 1 s. pulses per odorant have been used. Odorants were separated by 45 s.</li>
	<li>Note the response of the mosquito antennae in the lab notebook.<br />
	NOTE: If the odorant is detected by the mosquito antennae, a clear deflection in the signal is observed (see <strong class="xfig">Figure 3A</strong>).</li>
	<li>Proceed with the next odor or concentration. Do not forget to randomize the presentation of odorants unless a dose-response curve is performed.<br />
	NOTE: A negative control and a positive control should be used in the experiments. This will ensure that the responses observed are indeed olfactory responses and not due to mechanical or electric noise.</li>
	<li>At the end of the recording, apply a positive control to verify that the antennae are still responsive.<br />
	NOTE: Use 0.1% or 1% benzaldehyde as all mosquito species tested so far have been responsive to this compound.</li>
	<li>Proceed with the next mosquito preparation.</li>
</ol>
<p class="jove_title">8. Cleaning</p>
<ol>
	<li>Turn off the amplifier, the noise reducer, the airline, and the computer.</li>
	<li>Return the odorants to the freezer.</li>
	<li>Remove the filter papers from the glass syringes and clean with 100% ethanol if residue is visible on the walls. Let dry on a cleaning wipe overnight.</li>
	<li>Clean the electrode holders with DI water to remove any possible trace of salt. Dry by gently applying against a piece of cleaning wipe.</li>
	<li>Place mosquito remains in the freezer and dispose 24 h later.<br />
	&#8203;NOTE: If working with infected mosquitoes, follow safety requirements at your&#160;institution.</li>
</ol>
<p class="jove_title">9. Data analyses</p>
<ol>
	<li>Measure either manually or automatically the EAG responses.<br />
	NOTE: The EAG amplitude (-mV) is measured here. Average if multiple pulses have been applied for each compound. Depending on the software used, EAGs can be automatically detected and measured. However, it is essential to inspect each response individually to verify the shape of the response and assess the possible carry-over, delayed response, etc. The ideal EAG response is aligned with the pulse, shows a clear deflection, and is repeatable between mosquito preparations (<strong class="xfig">Figure 3</strong>).</li>
	<li>Present the raw data to show minimal variability, low noise signal, and clear responses (<strong class="xfig">Figure 3B</strong>).<br />
	NOTE: The data can also be normalized (e.g., Z-score). The negative control value (e.g., mineral oil) (i.e., the baseline) can be subtracted from the response and, if not, should be presented in the figures. A positive control should also be presented.</li>
	<li>Perform statistical analysis using any statistics software<sup class="xref">28</sup>.</li>
</ol>

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<p class="jove_content">The author has nothing to disclose.</p>

<p class="jove_content">Olfactory mediated behaviors are affected by many factors, including physiological (e.g., age, time of day) and environmental (e.g., temperature, relative humidity)<sup class='xref'>30</sup>. Thus, when conducting EAGs, it is essential to use insects that are in the same physiological status (i.e., monitoring for age, starving, mating)<sup class='xref'>31</sup> and to also maintain a warm and humid environment around the preparation to avoid desiccation. A temperature around 25 &#176;C is ideal and 60% to 80% humidity for the main airline. This can be easily achieved by placing a bubbler on the main airline circuit. </p>
<p class="jove_content">Moreover, it is important to consider the ecology of each species to obtain results that are relevant to the insect's biology. For example, if using a nocturnal species, consider inversing their light cycle to test their response during their subjective night. Choosing to conduct EAGs at specific moments of the day (i.e., when the insect is active) is also important. For example, if using <em>Ae. aegypti</em> mosquitoes, consider doing the experiments during the peaks of activity of this species (i.e., early in the day and later afternoon). Again, the light cycle can be easily shifted for convenience using climatic chambers or light boxes with an inversed light program using a programmable timer<sup class='xref'>32</sup>. Eilerts et al.<sup class='xref'>33</sup> and Krishnan et al.<sup class='xref'>34</sup>, have shown that the sensitivity to specific odorants varies throughout the day. Thus, a good knowledge of the insect's ecology and biology will guaranty more accurate results. </p>
<p class="jove_content">Noise (either electrical or mechanical) can be easily introduced in EAGs. For example, mechanical perturbations can be created by an AC system blowing air towards an EAG preparation. Electrical noise can be reduced with the Humbug, but, if persisting, can be tracked down by plugging elements and grounding them to the Faraday cage using alligator clips (<strong class='xfig'>Figure 3B</strong>). This applies to all the elements present around the preparation (i.e., microscope, lamp, micromanipulators). Some pieces of equipment in the Faraday cage should be unplugged before recording as they might still produce electrical noise (e.g., cold light source) or placed outside the cage. Another type of "noise" is of olfactory nature. The experimenter should avoid wearing perfume or use strongly scented shampoo or detergent. Indeed, many compounds found in these can be detected by mosquitoes (e.g., linalool, citronellol, geraniol, eugenol) and may interfere and affect the results of the experiments. Wearing a lab coat and gloves is also essential to limit unwanted contamination of the airline, syringes, and electrodes. </p>
<p class="jove_content">The presented protocol has the advantage of being easily applicable to all mosquito species, in both males and females, while extending the longevity of the preparation (&gt; 30 min) and with limited variability between preparations. This method leads to very minimal noise in the EAG signal, which allows for testing chemicals at very low concentrations. Once the dissection and mounting steps have been mastered, this technique can produce reliable data in a relatively short amount of time, and straightforward data analyses. </p>
<p class="jove_content">Electroantennography only allows the experimenter to assess whether the mosquito can detect a chemical or not. However, to determine the valence of this chemical, complementary behavioral assays, such as olfactometer assays, are critical to determining whether a specific odorant or mixture is attractive, repellent, or neutral in order to develop efficient tools for mosquito control<sup class='xref'>35</sup>. </p>

<p class="jove_content">Mosquitoes are the deadliest organisms on earth, claiming the lives of more than one million people per year and place more than half the world population at risk of exposure to the pathogens they transmit, while biting<sup class="xref">1</sup>. These insects rely on a wide range of cues (i.e., thermal, visual, mechanical, olfactory, auditory) to locate a host to feed on (both plant and animal), for mating and oviposition, as well as to avoid predators at both the larval and adult stages<sup class="xref">2</sup><sup>,</sup><sup class="xref">3</sup>. Among these senses, olfaction plays a critical role in the above mentioned behaviors, in particular for medium to long-range detection of odorant molecules<sup class="xref">2</sup><sup>,</sup><sup class="xref">3</sup>. Odors emitted by a host or an oviposition site are detected by various specific olfactory receptors (e.g., GRs, ORs, IRs) located on the mosquito palps proboscis, tarsi, and antennae<sup class="xref">2</sup><sup>,</sup><sup class="xref">3</sup>.</p>
<p class="jove_content">As olfaction is a key component of their host-seeking (plant and animal), mating and oviposition behaviors, it thus constitutes an ideal target to study to develop new tools for mosquito control<sup class="xref">4</sup>. Research on repellents (e.g., DEET, IR3535, picaridin) and baits (e.g., BG sentinel human lure) is extremely prolific<sup class="xref">5</sup>, but because of the current challenges in mosquito control (e.g., insecticide resistance, invasive species), it is essential to develop new efficient control methods informed by the mosquito biology.</p>
<p class="jove_content">Many techniques (e.g., olfactometer, landing assays, electrophysiology) have been used to assess the bioactivity of compounds or mixtures of compounds in mosquitoes. Among them, electroantennography (or electroantennograms (EAGs)) can be used to determine whether the odorants are detected by the mosquito antennae. This technique was initially developed by Schneider<sup class="xref">6</sup> and has been used in many different insect genera since then, including moths<sup class="xref">7</sup><sup>,</sup><sup class="xref">8</sup><sup>,</sup><sup class="xref">9</sup>, bumblebees<sup class="xref">10</sup><sup>,</sup><sup class="xref">11</sup>, honeybees<sup class="xref">12</sup><sup>,</sup><sup class="xref">13</sup><sup>,</sup> and fruit flies<sup class="xref">14</sup><sup>,</sup><sup class="xref">15</sup> to name a few. Electroantennography has also been employed using various protocols, including single or multiple antennae in mosquitoes<sup class="xref">16</sup><sup>,</sup><sup class="xref">17</sup><sup>,</sup><sup class="xref">18</sup><sup>,</sup><sup class="xref">19</sup><sup>,</sup><sup class="xref">20</sup><sup>,</sup><sup class="xref">21</sup><sup>,</sup><sup class="xref">22</sup><sup>,</sup><sup class="xref">23</sup><sup>,</sup><sup class="xref">24</sup><sup>,</sup><sup class="xref">25</sup>.</p>
<p class="jove_content">Mosquitoes are relatively small and delicate insects with rather thin antennae. While performing EAGs on larger insects such as moths or bumblebees is relatively easy because of their larger size and thicker antennae, conducting EAGs in mosquitoes can be challenging. In particular, maintaining a good signal-to-noise ratio and a lasting responsive preparation are two major requirements for data reproducibility and reliability.</p>
<p class="jove_content">The step-by-step guide to low noise EAGs proposed here directly offers solutions to these limitations and make this protocol applicable to several mosquito species from various genera, including <em>Aedes</em>, <em>Anopheles</em>, <em>Culex</em>, and <em>Toxorhynchites</em>, and describes the technique for both females and males. Electroantennography offers a quick yet reliable way to screen and determine bioactive compounds that can then be leveraged in bait development after valence has been determined with behavioral assays.</p>

<table><tbody><tr><td>Air table Clean Bench</td><td>TMC</td><td>https://www.techmfg.com/products/labtables/cleanbench63series/accessoriess</td><td>Noise reducer</td></tr><tr><td>Analog-to-digital board</td><td>National Instruments</td><td>BNC-2090A</td><td></td></tr><tr><td>Benchtop Flowbuddy Complete</td><td>Genesee Scientific</td><td>59-122BC</td><td>To anesthesize mosquitoes</td></tr><tr><td>Borosillicate glass capillary</td><td>Sutter Instrument</td><td>B100-78-10</td><td>To make the recording and references capillaries</td></tr><tr><td>Chemicals</td><td>Sigma Aldrich</td><td>Benzaldehyde: 418099-100 mL; Butyric acid: B103500-100mL; 1-Hexanol: 471402-100mL; Mineral oil: M8410-1L</td><td>Chemicals used for the experiments presented here</td></tr><tr><td>CO2</td><td>Airgas or Praxair</td><td>N/A</td><td>To anesthesize mosquitoes</td></tr><tr><td>Cold Light Source</td><td>Volpi</td><td>NCL-150</td><td></td></tr><tr><td>Disposable syringes</td><td>BD</td><td>1 mL (309628)&amp;nbsp; / 3 mL (309657)</td><td></td></tr><tr><td>Electrode cables</td><td>World Precision Instruments</td><td>5371</td><td></td></tr><tr><td>Electrode gel salt free</td><td>Parkerlabs</td><td>12-08-Spectra-360</td><td></td></tr><tr><td>Faraday cage</td><td>TMC</td><td>https://www.techmfg.com/products/electric-and-magnetic-field-cancellation/faradaycages</td><td>Noise reducer</td></tr><tr><td>Flowmeters</td><td>Bel-art</td><td>65 mm (H40406-0010) / 150 mm (H40407-0075)</td><td>One of each</td></tr><tr><td>GCMS vials and caps</td><td>Thermo-fisher scientific</td><td>2-SVWKA8-CPK</td><td>To prepare odorant dilutions</td></tr><tr><td>Glass syringes (Fortuna)</td><td>Sigma Aldrich</td><td>Z314307</td><td>For odor delivery to the EAG prep</td></tr><tr><td>Humbug</td><td>Quest Scientific</td><td>http://www.quest-sci.com/</td><td>Noise reducer</td></tr><tr><td>2 mm Jack Holder, Narrow, 90 deg., With Wire</td><td>A-M Systems</td><td>675748</td><td>Electrode holder</td></tr><tr><td>Magnetic bases</td><td>Kanetec</td><td>MB-FX</td><td>x 2</td></tr><tr><td>MATLAB + Toolboxes</td><td>Mathworks</td><td>https://www.mathworks.com/products/matlab.html</td><td>For delivering the pulses</td></tr><tr><td>Medical air</td><td>Airgas or Praxair</td><td>N/A</td><td>For main airline</td></tr><tr><td>Microscope</td><td>Nikkon</td><td>SMZ-800N</td><td></td></tr><tr><td>Micromanipulators Three-Axis Coarse/Fine Compact Micromanipulator</td><td>Narishige</td><td>MHW-3</td><td>x 2</td></tr><tr><td>Microelectrode amplifier with headstage</td><td>A-M Systems</td><td>Model 1800</td><td></td></tr><tr><td>Mosquito rearing supplies</td><td>Bioquip</td><td>https://www.bioquip.com/Search/WebCatalog.asp</td><td></td></tr><tr><td>Needles</td><td>BD</td><td>25G (305127) / 21G (305165)</td><td></td></tr><tr><td>Pasteur pipettes</td><td>Fisher Scientific</td><td>13-678-6A</td><td>For odor delivery to the EAG prep</td></tr><tr><td>PTFE Tubing of different diameters</td><td>Mc Master Carr</td><td>N/A</td><td>To connect solenoid valve, flowmeter, airline ect.</td></tr><tr><td>30V/5A DC Power Supply</td><td>Dr. Meter</td><td>PS-305DM</td><td></td></tr><tr><td>R version 3.5.1</td><td>R project</td><td>https://www.r-project.org/</td><td>For data analyses</td></tr><tr><td>Relay for solenoid valve</td><td>N/A</td><td>Custom made</td><td></td></tr><tr><td>Silver wire 0.01&amp;rdquo;</td><td>A-M Systems</td><td>782500</td><td></td></tr><tr><td>Solenoid valve (3-way)</td><td>The Lee Company</td><td>LHDA0533115H</td><td></td></tr><tr><td>WinEDR software</td><td>Strathclyde Electrophysiology Software</td><td>WinEDR V3.9.1</td><td>For EAG recording</td></tr><tr><td>Whatman paper</td><td>Cole Parmer</td><td>UX-06648-03</td><td>To load chemical in glass syringe / Pasteur pipette</td></tr></tbody></table>

a step-by-step guide to mosquito electroantennography

<p class="jove_content">Female mosquitoes are the deadliest animals on earth, claiming the lives of more than 1 million people every year due to pathogens they transmit when acquiring a blood-meal. To locate a host to feed on, mosquitoes rely on a wide range of sensory cues, including visual, mechanical, thermal, and olfactory. The study details a technique, electroantennography (EAG), that allows researchers to assess whether the mosquitoes can detect individual chemicals and blends of chemicals in a concentration-dependent manner. When coupled with gas-chromatography (GC-EAG), this technique allows to expose the antennae to a full headspace/complex mixture and determines which chemicals present in the sample of interest, the mosquito can detect. This is applicable to host body odors as well as plant floral bouquets or other ecologically relevant odors (e.g., oviposition sites odorants). Here, we described a protocol that permits long durations of preparation responsiveness time and is applicable to both female and male mosquitoes from multiple genera, including <em>Aedes</em>, <em>Culex</em>, <em>Anopheles</em>, and <em>Toxorhynchites</em> mosquitoes. As olfaction plays a major part in mosquito-host interactions and mosquito biology in general, EAGs and GC-EAG can reveal compounds of interest for the development of new disease vector control strategies (e.g., baits). Complemented with behavioral assays, the valence (e.g., attractant, repellent) of each chemical can be determined.</p>

<p class="jove_content">Electroantennography is a powerful tool to determine whether a chemical or blend of chemicals is detected by an insect antenna. It can also be used to determine the detection threshold for a given chemical using a gradual increase of concentration (i.e., dose curve response, <strong class="xfig">Figure 4B</strong>). Moreover, it is useful to test the effects of repellent on the response to host-related odors<sup class="xref">29</sup>.</p>
<p class="jove_content">Positive and negative controls should always be used in EAGs. Here, benzaldehyde was used as a positive control (<strong class="xfig">Figure 3B</strong>, <strong>3C</strong>, <strong>4A</strong>). This compound has been found to elicit an antennal response in all mosquito species tested so far<sup class="xref">24</sup><sup>,</sup><sup class="xref">25</sup><sup>,</sup><sup class="xref">29</sup>. A negative control should also be used and can consist of the solvent used for diluting the chemicals (e.g., mineral or paraffin oils, hexane, etc.) and should not elicit a response (<strong class="xfig">Figure 3B</strong>, <strong>3C</strong>, <strong>4A</strong>).</p>
<p class="jove_content">Indeed, when conducting EAGs, a deflection should not be noted when applying the control (<strong class="xfig">Figure 3B</strong>, <strong>3C</strong>, <strong>4A</strong>). If a response is observed, either the syringe, the solvent control, and/or the odor line is likely contaminated. If it is the case, a new solution should be prepared, the syringe cleaned with 100% ethanol and dried and/or the airline decontaminated by rinsing with 100% ethanol and dried. If the chosen control elicits a response (e.g., ethanol), the value obtained in -mV for the control should be subtracted from the value obtained for ethanol and tested chemical combined to assess the impact of the tested chemical on the antennae.</p>
<p class="jove_content">Mosquito species vary in their ability to respond to various compounds as well as in the magnitude of their response. For example, <em>Toxorhynchites</em> mosquitoes produce very large EAGs in comparison to <em>Ae. aegypti</em>, <em>An. Stephensi</em> and <em>Cx. quinquefasciatus</em> (<strong class="xfig">Figure 3C</strong>, <strong class="xfig">Figure 4A</strong>).</p>
<p class="jove_content">In EAGs, the second pulse and the following usually lead to smaller EAG responses. The presentation of one odorant can also affect the response to the following, so it is important to randomize the odorant order and multiple assays to efficiently test a panel of odorants (unless a dose-response curve is performed). Moreover, separating pulses (e.g., 5 s) and odorants (e.g., 45 s) presentation will help to optimize the EAG responses.</p>
<p class="jove_content">The volatility of the tested chemicals varies and can affect the olfactory response and potentially lead to a delayed response if the tested chemical has very low volatility. Chemical volatility and solubility should be known before conducting EAGs to optimize the assay. The solvent used to prepare the dilutions should also be carefully selected (e.g., ethanol, hexane, mineral, or paraffin oil). Moreover, concentrations should be chosen wisely and should ideally be ecologically relevant. A 1% or 0.1% concentration is often used but is relatively high and not necessarily representative of what the insects can experience in nature. Yet, it is useful to screen compounds with relatively high concentrations in some cases (e.g., for bait development). Repellents can be tested at their commercially available concentration (e.g., DEET is typically sold at a 40% concentration).</p>
<p class="jove_content" fo:keep-together.within-page="1">If coupled with gas chromatography (i.e., GC-EADs)<sup class="xref">25</sup>, the compounds eliciting a response can be identified with a GC-MS and then tested individually at various concentrations or in mixtures with EAGs. It is worth mentioning that the valence of the tested chemicals cannot be determined with EAGs. Only a complementary behavioral experiment (e.g., olfactometer, feeding assay) can assess whether the chemical detected by the antennae is attractive, repellent, or neutral to the mosquito. Finally, EAG experiments are only showing responses of the peripheral nervous system.</p>
<p class="jove_content biglegend" fo:keep-together.within-page="1"><img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62042/62042fig1v2.jpg" /><br />
<strong class="xfig">Figure 1</strong><strong>: Electroantennogram setup comprised of:</strong> <strong>A</strong>) Microscope: the microscope used should allow the experimenter to clearly see the preparation to allow the mosquito antennae tips to be inserted in the recording electrodes. <strong>B</strong>) Cold light lamp: the lamp should be turned off when the recordings begin. <strong>C</strong>) Vacuum line: this reduces the risk of accumulation of the odorants around the mosquito head preparation, which could result in antennal responses decoupled from actual stimulation. <strong>D</strong>) Micromanipulators (x2): these will allow for very fine electrode holders movements, which is required for inserting the mosquito antennae in the capillary of the recording electrode. <strong>E</strong>) Recording electrode holder. <strong>F</strong>) Reference electrode holder. <strong>G</strong>) Head stage: both electrodes are plugged in the head stage which is then connected to the amplifier. <strong>H</strong>) Main airline: a constant clean airflow bathed the mosquito head. The flow rate is regulated by a flowmeter. <strong>I</strong>) Syringe for odor delivery connected to the solenoid valve and flowmeter; <strong>J</strong>) Air table: the air table will reduce noise. <strong>K</strong>) Faraday cage: The Faraday cage will prevent electric noise. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/62042/62042fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a></p>
<p class="jove_content biglegend" fo:keep-together.within-page="1"><img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62042/62042fig2v2.jpg" /><br />
<strong class="xfig">Figure 2</strong><strong>: Step-by-step <em>Aedes</em> <em>albopictus</em> mosquito head preparation for EAG recordings.</strong> <strong>A</strong>) Female mosquito on its back on an icy plate to verify that both antennae are intact. <strong>B</strong>) Last segment of the antennae excision with micro scissors. <strong>C</strong>) Antennae are dipped in electrode gel. <strong>D</strong>) The antennae stick together after pulling them out. Only one segment of each antenna should be in the electrode gel. <strong>E</strong>) Mosquito head excision. <strong>F</strong>) Head mounted on the reference electrode. It should be stable enough to be moved to the EAG rig. <strong>A&#39;-F&#39;</strong>. Same steps as presented above for male EAGs. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/62042/62042fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a></p>
<p class="jove_content biglegend" fo:keep-together.within-page="1"><img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62042/62042fig3v2.jpg" /><br />
<strong class="xfig">Figure 3</strong><strong>: Schematic of the mosquito EAG and raw EAG traces. A)</strong> EAG schematic (left) and characteristics of the EAG response (right). (Left) The mosquito head is mounted between a reference electrode and a recording electrode connected to an amplifier. The antennae are bathed in a constant airflow in which odorant stimuli are pulsed. Detection of a chemical leads to a deflection (in mV) in the signal. (right) The chemical detection leads to cell depolarization (DPR) followed by cell repolarization (RPR) until return to baseline.&#160;The odorant pulse is represented by the gray rectangle.&#160;The red line indicates the amplitude of the EAG response.&#160;<strong>B</strong>) Screenshot of the WinEDR software highlighting a whole EAG recording trace of a <em>Culex quinquefasciatus </em>female mosquito. Top: unfiltered (i.e., raw) signal. Middle: 1 s odor pulses are indicated by numbers. Bottom: filtered (i.e., 1.5 Hz low pass) signal to 3 odorants and a control (mineral oil). Note the deflections in response to 1% 1-hexanol (1), 1% benzaldehyde (2), and 1% butyric acid (3). Note the absence of response to the negative control, mineral oil (4). <strong>C)</strong> From left to right: Representative EAG responses (in mV) to 1% benzaldehyde (top) and a mineral oil control (bottom) in females <em>Aedes aegypti</em>, <em>Anopheles stephensi, Culex quinquefasciatus</em>, and<em> Toxorhynchites rutilus septentrionalis</em>. The one-second pulse is represented by the colored rectangle above the EAG trace. Note the large deflection in response to benzaldehyde and the lack of response to the mineral oil. Also, note the different scale in <em>Toxorhynchites rutilus septentrionalis</em>. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/62042/62042fig3v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a></p>
<p class="jove_content biglegend" fo:keep-together.within-page="1"><img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62042/62042fig4v2.jpg" /><br />
<strong class="xfig">Figure 4</strong><strong>:&#160;Example representation of EAG results and their statistical analyses. A.</strong> Average EAG responses of <em>Culex quinquefasciatus</em> (N = 8), <em>Anopheles stephensi</em> (N = 10), <em>Aedes aegypti</em> (N = 8) and <em>Toxorhynchites rutilus septentrionalis</em> (N = 7) females to 1% 1-hexanol (green), 1% butyric acid (orange), 1% benzaldehyde (yellow) and mineral oil (blue). <strong>B.</strong> <em>Culex quinquefasciatus</em> females EAG dose-response curve for 1-hexanol (left) (N = 9) and benzaldehyde (right) (N = 8). Bars represent standard error of the mean. Letters above error bars indicate statistical differences (Pairwise Wilcoxon rank sum test with a Bonferroni correction).&#160;<a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/62042/62042fig4v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a></p>

Watch this Scientific Journal Video about A Step-by-Step Guide to Mosquito Electroantennography at JoVE.com

A Step-by-Step Guide to Mosquito Electroantennography

<p class="jove_content">The present article details a step-by-step protocol for successful and low noise electroantennograms in several genera of mosquitoes, including both females and males. </p>

Female mosquitoes are the deadliest animals on earth, claiming the lives of more than 1 million people every year due to pathogens they transmit when ...

a-step-by-step-guide-to-mosquito-electroantennography

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4.6K Views.  Virginia Polytechnic Institute and State University. The present article details a step-by-step protocol for successful and low noise electroantennograms in several genera of mosquitoes, including both females and males. 

Video: A Step-by-Step Guide to Mosquito Electroantennography

Galvanic Cells

Galvanic Cells: Construction and Reduction Potential Measurement - Procedure

Ogniwa galwaniczne

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

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

Education

JoVE Lab Manual

Chemistry

Lab Manual Sub Articles

Electrolytic Cells

<p>Source: Smaa Koraym at Johns Hopkins University, MD, USA</p>
<ol class="note-items">
	<li data-note-item="1" data-parent-collapse="">Electrolytic Cells<button class="expand">Expand</button>
	<p>In this experiment, you will be assembling an electrolytic cell to perform electroplating. An electrolytic cell contains an anode and cathode connected by a power source, which drives the nonspontaneous redox reaction in the cell. When the anode material is oxidized, the ions travel to the cathode, where they are reduced and plated.</p>
	<p>In the case of this experiment, the electrolytic cell contains a copper anode and a brass key for the cathode. Oxidation of the anode creates copper(2+) ions, which are then reduced on the brass key, forming a thin plating of solid copper.</p>
	<ul class="lab-items">
		<li data-sub-note-item="1-3">
		<div class="lab-step">Put on gloves, safety goggles, and a lab apron, which must be worn at all times.</div>
		</li>
		<li data-sub-note-item="1-4">
		<div class="lab-step">Obtain one 10-ohm resistor, one copper sheet electrode, one copper wire, and one piece of emery paper.</div>
		</li>
		<li data-sub-note-item="1-5">
		<div class="lab-step">Use the emery paper to polish the brass key and the copper sheet. Rinse the key and the copper electrode with water. The brass key will be the cathode, and the copper sheet will be the anode.</div>
		</li>
		<li data-sub-note-item="1-6">
		<div class="lab-step">Take one 50-mL beaker and a watch glass to the instructor&#39;s hood and obtain approximately 20 mL of acetone. Place the watch glass on top of the beaker to prevent evaporation and bring it back to your bench.</div>
		</li>
		<li data-sub-note-item="1-7">
		<div class="lab-step">Label the other 50-mL beaker as &#39;organic waste&#39;.</div>
		</li>
		<li data-sub-note-item="1-8">
		<div class="lab-step">Use a disposable pipette to rinse the copper electrode and key with acetone. Collect excess in the waste beaker. Place the key and copper electrode on a paper towel to dry.</div>
		</li>
		<li data-sub-note-item="1-9">
		<div class="lab-step">Measure the initial masses of the key and copper electrode and record their values in your lab notebook.
		<h4><strong>Table 1: Moles of Cu Transferred</strong></h4>
		<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
			<tbody>
				<tr>
					<td></td>
					<td style="text-align: center;"><strong>Mass of key (g) </strong></td>
					<td style="text-align: center;"><strong>Change in mass (g) </strong></td>
					<td style="text-align: center;"><strong>Moles Cu plated </strong></td>
					<td style="text-align: center;"><strong>Mass of Cu electrode (g) </strong></td>
					<td style="text-align: center;"><strong>Change in mass (g) </strong></td>
					<td style="text-align: center;"><strong>Moles Cu lost </strong></td>
				</tr>
				<tr>
					<td style="text-align: center;">0 min</td>
					<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.bdigitaluss.remotexs.co/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.bdigitaluss.remotexs.co/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

Ogniwa elektrolityczne

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

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

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.bdigitaluss.remotexs.co/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.bdigitaluss.remotexs.co/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.bdigitaluss.remotexs.co/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

Zachowanie zwierząt

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 ...

Ecology

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.bdigitaluss.remotexs.co/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.bdigitaluss.remotexs.co/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.bdigitaluss.remotexs.co/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

Wymieranie

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

Species Distribution and Biogeography

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Simulating Colonization and Extinction on Islands
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">Form groups of three. One member of the group will simulate colonization by throwing balls into the groups of cups representing the islands. Another will simulate extinction by rolling dice to randomly remove balls from the cups. The third group member will record the data.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">Take time to familiarize yourself with the four island stations. NOTE: There are two large islands with 12 cups and two small islands with 6 cups. Two of the stations will have tape lines that are 2 m away from the cups. These are the far islands. The other two stations will have tape lines that are 1 m away from the cups. These are the near islands. Thus, the stations are far large and near large and far small and near small. HYPOTHESES: In this experiment, one alternate hypothesis might be that the larger islands with more cups will support a greater number of species than the smaller islands. The null hypothesis might be that all islands will support this same number of species regardless of size. Another alternate hypothesis might be that islands closer to the mainland will have a higher colonization rate than islands farther from the mainland. The null hypothesis might be that all islands will have the same colonization rate regardless of their distance from the mainland. Additionally, in all trials we may hypothesize that the colonization rate will be expected to decrease as more species inhabit an island whereas the extinction rate will increase as more species are found on the island. The null hypothesis might be that the colonization rate and extinction rates are expected to stay the same across all numbers of species on the island.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">First, choose one of the four stations as a starting place.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">From the line of tape at this location, the colonization simulator must throw the ball to the island and try to get the ball into a cup. NOTE: One round consists of five throws, each throw representing a colonization attempt. If the ball lands in a cup that does not already have a ball in it, this is a successful colonization attempt and the ball should be left in the cup.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">The data collector should place a tally on the colonizers box for the round in the recording sheet found at the station. <a href="https://www-jove-com.bdigitaluss.remotexs.co/CDNSource/lm/labs/29/Tables_1-4_Island_biogeography.xlsx" style="display:block;margin:20px;" target="_blank">Click Here to download Tables 1-4</a>
</div></li><li data-sub-note-item="1-6"><div class="lab-step">If the ball does not land in a cup, retrieve it. This is an unsuccessful colonization attempt.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">If a ball lands in a cup that already has a ball in it, retrieve it. This is also an unsuccessful colonization attempt.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">Add the number of species at the beginning of the round with the tally of colonizers.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">The data collector should record this number in the column entitled &#8220;Number of species after colonization&#8221; in the recording sheet found at the station.
</div></li><li data-sub-note-item="1-10"><div class="lab-step">The extinction simulator should now roll a die. If the trial is occurring at a large island site, choose one of the two colored dice at random with eyes closed.
</div></li><li data-sub-note-item="1-11"><div class="lab-step">Remove the ball if there is one present from the cup that is of the same color as the die and labeled with the number rolled by that die.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">The data collector should count the removal as an extinction in the station recording table. NOTE: In a trial at the small island station, follow the same procedure except that it is only necessary to match the number rolled by the die to the cup. If there is no ball in the cup, no extinction is recorded. For both the large and small islands, repeat the extinction roll for a total of two rolls if there are 3 - 8 species present on the island and three rolls if there are 9 - 12 species on the island. If there are fewer than three species on the island, do not roll the die again.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">This is the end of one round. After this, count the total number of species currently in the cups at this station and then tally this number into the &#8220;Number of species at beginning of round&#8221; column on the station recording sheet for the next round.
</div></li><li data-sub-note-item="1-14"><div class="lab-step">Groups should switch stations after five rounds at one table to control for the throwing ability of the colonization simulators. Repeat the trials and rotation through the stations until 20 rounds have been recorded in total for all groups combined at all four stations.
</div></li><li data-sub-note-item="1-15"><div class="lab-step">Count the tally marks for the number of species at the beginning of the round, colonizers, total number of species after colonization, and extinctions for each round at each island station.
</div></li><li data-sub-note-item="1-16"><div class="lab-step">Look at your data recording sheet. How many times were there zero species at the beginning of a round? How many times was there only one species at the beginning of a round? Repeat this calculation for 0 through 12 species present at the beginning of a round.
</div></li><li data-sub-note-item="1-17"><div class="lab-step">Then, tally these values in the corresponding rows of the &#8220;Rounds with this many species&#8221; column in the appropriate colonization rate calculation table. <a href="https://www-jove-com.bdigitaluss.remotexs.co/CDNSource/lm/labs/29/Tables_5-8_Island_biogeography.xlsx" style="display:block;margin:20px;" target="_blank">Click Here to download Tables 5-8</a>
</div></li><li data-sub-note-item="1-18"><div class="lab-step">In the colonization attempts column, multiply these values by five to represent the total number of Ping-Pong balls thrown.
</div></li><li data-sub-note-item="1-19"><div class="lab-step">Look at your data recording sheet once more and calculate the total number of colonizers for each value in the number of species at beginning of round column.
</div></li><li data-sub-note-item="1-20"><div class="lab-step">Record these values in the &#8220;Successful colonization attempts&#8221; column of the appropriate colonization rate calculation table.
</div></li><li data-sub-note-item="1-21"><div class="lab-step">Divide the successful colonization attempts by the colonization attempts for each initial species value and record this in the &#8220;Colonization rate&#8221; column of the appropriate colonization rate calculation table.
</div></li><li data-sub-note-item="1-22"><div class="lab-step">Repeat the process for determining the colonization rate for all of the island trials. 
</div></li><li data-sub-note-item="1-23"><div class="lab-step">To calculate the extinction rate for each of the trials, return to the data recording sheet for the near large island trial. Count the number of times that the number of species after colonization matched each number from 1 to 12, as previously described.
</div></li><li data-sub-note-item="1-24"><div class="lab-step">Record these values in the &#8220;Rounds with this many species&#8221; column in the appropriate extinction rate calculation table. <a href="https://www-jove-com.bdigitaluss.remotexs.co/CDNSource/lm/labs/29/Tables_9-12_Island_biogeography.xlsx" style="display:block;margin:20px;" target="_blank">Click Here to download Tables 9-12</a>
</div></li><li data-sub-note-item="1-25"><div class="lab-step">To calculate extinction attempts, multiply the value in the &#8220;Rounds with this many species&#8221; cells by 1 if the species after colonization value is 0 &#8211; 2, by 2 if the value is 3 &#8211; 8, and by 3 if the value is 9 or above. NOTE: These values correspond with the number of times the die was rolled during the extinction simulation.
</div></li><li data-sub-note-item="1-26"><div class="lab-step">Then, using the data contained in the data recording sheet, count the number of extinctions that occurred for each value in the &#8220;Number of species after colonization&#8221; column.
</div></li><li data-sub-note-item="1-27"><div class="lab-step">Record these values in the &#8220;Extinctions&#8221; column of the appropriate extinction rate calculation table.
</div></li><li data-sub-note-item="1-28"><div class="lab-step">Record these values in the &#8220;Extinctions&#8221; column of the appropriate extinction rate calculation table.
</div></li><li data-sub-note-item="1-29"><div class="lab-step">Repeat the process for calculating the extinction rate for all of the island trials. 
</div></li><li data-sub-note-item="1-30"><div class="lab-step">After this, plot the colonization and extinction rates versus the number of species for each of the island stations.
</div></li><li data-sub-note-item="1-31"><div class="lab-step">Draw the best-fit lines for the colonization and extinction rates in each of the four graphs.
</div></li><li data-sub-note-item="1-32"><div class="lab-step">Compare the colonization rate between the near and far islands. Determine the distance that had a greater dynamic equilibrium of species and develop a hypothesis to explain your observation. 
</div></li><li data-sub-note-item="1-33"><div class="lab-step">Next, compare the extinction rates between the large and small islands. Determine the size that had a greater dynamic equilibrium number of species.
</div></li><li data-sub-note-item="1-34"><div class="lab-step">Observe the slopes of the colonization rates in the plots. Determine if they are positive or negative. 
</div></li><li data-sub-note-item="1-35"><div class="lab-step">Observe the slopes of the extinction rates in the plots. Determine if they are positive or negative. 
</div></li></ul></li></ol>

Rozmieszczenie gatunków i biogeografia

Simulating Colonization and Extinction on Islands
ExpandForm groups of three. One member of the group will simulate colonization by throwing balls into ...

Energy Dynamics

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Caterpillar Energy Dynamics
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">NOTE: In this lab you will grow caterpillars in conditions with and without food to examine the transfer of biomass in a typical trophic system.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">Weigh and record the wet mass of enough leaves of cabbage to line the bottom of a plastic tub.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Then, collect two batches of five caterpillars from their cabbage habitat, and weigh and record the wet mass of each batch separately in individual weigh boats. Make sure to tare the balance before each measurement. HYPOTHESES: The experimental hypothesis is that only a fraction of energy will be transferred between the trophic levels of the cabbage and the caterpillars and a significant amount will be lost as waste and heat energy. The null hypothesis is that there will be no significant difference between the energy input from the cabbage, and the obtained energy stored as biomass in the caterpillars.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Place the weighed cabbage and five caterpillars in a plastic tub lined with damp paper towels, leaving the lid slightly cracked open for ventilation, but not wide enough for the animals to escape.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Place the other five caterpillars in a second clean container prepared the same way, but with no cabbage.
</div></li><li data-sub-note-item="1-6"><div class="lab-step">After three days, transfer the caterpillars to new weigh boats, and weigh and record the final wet mass of both groups of caterpillars and the cabbage.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Then place the caterpillar waste, or frass, and cabbage into two separate dishes, and put both dishes into a 37 &#176;C incubator to dry for 48 hours.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">Euthanize the weighed caterpillars by putting them into a minus 20 &#176;C freezer for 24 hours.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">Then, transfer the dishes to a 37 &#176;C incubator for 48 hours.
</div></li><li data-sub-note-item="1-10"><div class="lab-step">After drying, weigh and record the dry mass of the uneaten cabbage, the two groups of caterpillars, and the frass, and use these data to calculate the percent biomass of the caterpillars.
</div></li><li data-sub-note-item="1-11"><div class="lab-step">Finally, discard all of the dried organisms and plates into the hazardous waste bin, and wipe down your workspace with 70% ethanol.
</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">Take a look at the data you collected. Ensure the initial wet mass, the final wet mass, and the final dry mass columns are accurately filled out in your table. <a href="https://www-jove-com.bdigitaluss.remotexs.co/CDNSource/lm/labs/13/Table_1_Energy_Dynamics.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 1</a>
</div></li><li data-sub-note-item="2-2"><div class="lab-step">NOTE: Remember an object&#39;s biomass is equal to its dry mass. Therefore, percent biomass for each group is the final dry mass divided by the final wet mass times 100. For example, obtain the percent biomass of the cabbage by dividing the final dry mass by the final wet mass and multiplying by 100. This means that approximately 15% of the cabbage&#39;s total weight is biomass.
</div></li><li data-sub-note-item="2-3"><div class="lab-step">After this, estimate the initial biomass of the cabbage and caterpillars by multiplying the percent biomass by the initial wet mass, and put the values in this new table. NOTE: Remember, biomass equals dry mass. Therefore, the final dry mass is equal to the final biomass of each group.
</div></li><li data-sub-note-item="2-4"><div class="lab-step">To calculate the approximate initial and final energy in the form of kilocalories within the cabbage, caterpillars, and frass, multiply the biomass values recorded in your table by the conversions, and record the calculated values in the table. <a href="https://www-jove-com.bdigitaluss.remotexs.co/CDNSource/lm/labs/13/Table_2_Energy_Dynamics.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 2</a>
</div></li><li data-sub-note-item="2-5"><div class="lab-step">After this, subtract the initial energy from the final energy to obtain the change in energy for each group.
</div></li><li data-sub-note-item="2-6"><div class="lab-step">NOTE: The negative sign denotes loss in energy, and positive indicates gain.
</div></li><li data-sub-note-item="2-7"><div class="lab-step">Using your data table, calculate the energy in kilocalories that the fed caterpillars lost to respiration. The formula is the energy gained from cabbage, which is the positive of the energy lost by cabbage, minus the biomass gained from cabbage, minus the energy lost via frass. <a href="https://www-jove-com.bdigitaluss.remotexs.co/CDNSource/lm/labs/13/Table_3_Energy_Dynamics.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 3</a>
</div></li><li data-sub-note-item="2-8"><div class="lab-step">Then, repeat the same calculation for the unfed group of caterpillars. Make another column in the table and note this down.
</div></li><li data-sub-note-item="2-9"><div class="lab-step">Finally, add up all of the energy gained via biomass and cabbage consumption, and the energy lost via frass and respiration for each group of caterpillars. NOTE: For both groups of caterpillars the net energy is zero, meaning that this experiment obeyed the first law of thermodynamics.
</div></li></ul></li></ol>

Studying Energy Dynamics and Productivity - Procedure

Dynamika energii

Caterpillar Energy Dynamics
ExpandNOTE: In this lab you will grow caterpillars in conditions with and without food to examine the transfer of biomass in a ...

Sexual Selection and Mate Choice

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Sexual Selection Class Simulation
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">Note: In this activity, you and your classmates will portray a sexually reproducing population consisting of an equal number of male and female animals. You will simulate several different mating scenarios to determine how relative quality, attractiveness, sex, and parental commitment affect mate choice and reproductive success. The goal of this activity is to maximize your progeny. 
</div></li><li data-sub-note-item="1-2"><div class="lab-step">Hypotheses: There are several overall hypotheses for female and male behavior for this activity. The main experimental hypothesis for female behavior is that females will mate with males of high quality more frequently than those of low quality, yielding a higher reproductive success. The null hypothesis is that females will not demonstrate preference towards high- or low-quality males in any of the four rounds of mating. For males, one experimental hypothesis is that males with a higher visible score will have greater reproductive success. Another could be that males will be more selective for females in scenarios in which they join females to care for offspring as opposed to scenarios in which they exhibit no parental commitment. The null hypothesis for male behavior might be that males will not demonstrate preference towards high- or low-quality females in any of the four rounds of mating. 
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Activity Overview: To begin, each student will be assigned a sex as well as a relative quality and attractiveness score. Remember - the goal is to maximize your reproductive success in each round. You do this by mating with partners of the highest possible quality which is rated on a scale of one to three with three being the highest possible score. Attractiveness scores, which follow the previously described scoring paradigm, correlate with quality but do not exactly predict the score. Though useful, they are not a perfect way to choose the best mate.  If you are assigned a male ID holder, you will have 16 gametes. However, if you are assigned a female ID holder, you will only have four. A successful mating occurs when one sperm and one egg are paper-clipped together and deposited at the fertilization area. 
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Note - Rules of the activity: 
<ul>
	<li>Assigned male and female identities for these experiments are not representative of students&#39; personal gender identities. </li>
	<li>Running is not permitted. </li>
	<li>Talking is not permitted.</li>
	<li>Hidden values cannot be shared but individuals can attempt to hide their visible scores. </li>
	<li>Respect the space around each individual and remain an arm&#39;s length away. </li>
	<li>Offspring will only be counted towards an individual score if both gametes are paper-clipped together and dropped off in the designated fertilization area before the end of the two-minute round. </li>
</ul>
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Round 1 &#8211; Hidden Quality Score
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">To begin the first simulation, take the clip-on ID holder you were randomly assigned and make sure it contains four gametes if you are assigned female or sixteen gametes if male. Males should also have a strip of white paper and two paperclips. 
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Next, males should paperclip the white strip of paper over their quality score to hide it, but keep the attractiveness score visible. 
</div></li><li data-sub-note-item="2-3"><div class="lab-step">Record your ID number for round one in table two.  <a href="https://www-jove-com.bdigitaluss.remotexs.co/CDNSource/lm/labs/35/Table_2_sexual_selection.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">Next, clip on the ID holder and move to the designated mating area. 
</div></li><li data-sub-note-item="2-5"><div class="lab-step">When the instructor says begin, you may commence the mating simulation. You will have exactly two minutes to produce as many offspring as possible. Males may freely give their sperm to females, and females may choose to use these sperm immediately after which they would walk to the fertilization area and paperclip it to one of their eggs, or they may choose to wait until they receive other sperm to do so. 
</div></li><li data-sub-note-item="2-6"><div class="lab-step">Once the gametes have been deposited in the fertilization area, females may resume mating. 
</div></li><li data-sub-note-item="2-7"><div class="lab-step">When the two minutes have elapsed, all matings must stop and any gametes not in the fertilization area at this time will not be counted. 
</div></li><li data-sub-note-item="2-8"><div class="lab-step">The instructor should now tally all of the matings and record the numbers of the individuals who produced each offspring.
</div></li><li data-sub-note-item="2-9"><div class="lab-step">Now, find all the gametes that came with your ID holder and repackage them for the next round. 
</div></li></ul></li><li data-note-item="3" data-parent-collapse="">Round 2 &#8211; Hidden Attractiveness Score
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="3-1"><div class="lab-step">For the second round, obtain a new randomly assigned ID holder and again ensure it contains the proper number of gametes. 
</div></li><li data-sub-note-item="3-2"><div class="lab-step">Next, males should use the paperclip and strip of white paper to hide their attractiveness score (instead of the quality score, which was covered in round one).
</div></li><li data-sub-note-item="3-3"><div class="lab-step">Record the ID number of your clip in table two under round two.  Note: The mating simulation process for round two will be the same as round one.
</div></li><li data-sub-note-item="3-4"><div class="lab-step">Move to the designated mating area and with for the instructor to signal the start of the round. As before, males may freely give away their sperm while females will choose to use it now or wait for different sperm. When a female decides to fertilize an egg, she will walk to the fertilization area and paperclip the gametes together. 
</div></li><li data-sub-note-item="3-5"><div class="lab-step">Proceed with the simulation for two minutes, and at the signal from the instructor all mating activity should stop. 
</div></li><li data-sub-note-item="3-6"><div class="lab-step">The instructor should now tally all of the matings from this second round.
</div></li><li data-sub-note-item="3-7"><div class="lab-step">Now, find and return the gametes to your ID holder, and then return the holder to the instructor.
</div></li></ul></li><li data-note-item="4" data-parent-collapse="">Round 3 &#8211; Visible Quality and Attractiveness Scores
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="4-1"><div class="lab-step">For the third simulation, you should again obtain a new clip-on ID holder and check that it contains the appropriate number of gametes. Record the ID number of your clip in table two.
</div></li><li data-sub-note-item="4-2"><div class="lab-step">For this round, leave both attractiveness and quality scores visible on male IDs. Note: The mating simulation process for round three will be the same as the previous two rounds.
</div></li><li data-sub-note-item="4-3"><div class="lab-step">Move to the mating area and wait for the signal to begin the simulation.
</div></li><li data-sub-note-item="4-4"><div class="lab-step">Once the instructor starts the timer and gives the signal, commence mating. As before, males are allowed to freely give away their sperm to females and females may use the sperm immediately or wait for different sperm. Again, fertilized eggs only count if placed in the fertilization area. 
</div></li><li data-sub-note-item="4-5"><div class="lab-step">After the two-minute time limit, the instructor should tally all matings. 
</div></li><li data-sub-note-item="4-6"><div class="lab-step">Now, find and return the gametes to your ID holder, and then return the holder to the instructor.
</div></li></ul></li><li data-note-item="5" data-parent-collapse="">Round 4 &#8211; Increased Male Parental Care
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="5-1"><div class="lab-step">In the final round, males will again leave their attractiveness and quality scores visible. 
</div></li><li data-sub-note-item="5-2"><div class="lab-step">Obtain an ID holder and ensure it contains the appropriate number of gametes. 
</div></li><li data-sub-note-item="5-3"><div class="lab-step">Clip on the ID holder and move to the mating area. Note: As before, males may freely give away their sperm to females. However, in this round, once a female chooses to mate with a male, both the male and female will walk to the fertilization area to paperclip their gametes together. After placing the paper-clipped gametes on the table, both male and female actors may return to the mating area. 
</div></li><li data-sub-note-item="5-4"><div class="lab-step">The instructor should signal the start of the round and begin the timer &#8211; at this point participants can begin the simulation.
</div></li><li data-sub-note-item="5-5"><div class="lab-step">Once the two-minute time limit is up, all matings must stop. 
</div></li><li data-sub-note-item="5-6"><div class="lab-step">The instructor should tally all mating events.
</div></li><li data-sub-note-item="5-7"><div class="lab-step">Find and return the gametes to your ID holder. Give your ID holder to the instructor and notify them if any gametes or supplies are missing. 
</div></li></ul></li><li data-note-item="6" data-parent-collapse="">Results
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="6-1"><div class="lab-step">To analyze the data generated during the mating activities, first split up into groups of four. 
</div></li><li data-sub-note-item="6-2"><div class="lab-step">Collect the class data from the instructor for ONE of the four rounds of mating. Note: In this activity, there were several types of females and males distinguished by different letters. These letter types correspond to specific combinations of quality and attractiveness scores. In the compiled data, the average number of matings and reproductive success has already been sorted by letter type for you by the instructor. 
</div></li><li data-sub-note-item="6-3"><div class="lab-step">Use these data to generate four graphs for the round you were assigned. First plot two graphs for the average number of matings against female and male type. 
</div></li><li data-sub-note-item="6-4"><div class="lab-step">Then plot the reproductive success for males and females against male or female type, giving a total of four graphs.
</div></li><li data-sub-note-item="6-5"><div class="lab-step">Once these graphs are complete, each group should present their data to the class, making sure to state the original hypothesis for your assigned round and whether or not the hypothesis was upheld.
</div></li><li data-sub-note-item="6-6"><div class="lab-step">After each group presents, discuss the overall results of the four rounds together. 
</div></li><li data-sub-note-item="6-7"><div class="lab-step">Note the affect that hiding the quality score had on the overall quality of mates chosen.
</div></li><li data-sub-note-item="6-8"><div class="lab-step">Note how hiding or revealing the attractiveness score affected the results.
</div></li><li data-sub-note-item="6-9"><div class="lab-step">Discuss the effect of male parental investment on the number and overall quality of mating pairs.
</div></li></ul></li></ol>

Dobór płciowy i wybór partnera

Sexual Selection Class Simulation
ExpandNote: In this activity, you and your classmates will portray a sexually reproducing population consisting of an ...

Artificial Selection

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">Observing Artificial Selection in Plants 
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">To begin the experiment, collect five two-week old grown plants from the generation one population that appear healthy. Note: Plants should appear green, not wilted, and not have damaged leaves. Experimental Hypothesis: The experimental hypothesis for this lab might be that the number of lead trichomes varies naturally, but the average number will increase over generations if only plants with the highest number of trichomes are interbred or decrease if those with the least are interbred. Null Hypothesis: The null hypothesis might be that the average number of trichomes will not change over subsequent generations after artificial selection.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">Label each plant with a letter from A to E and with both partners&#8217; names.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Record any natural differences you see between your plant samples. Note: All of the plants are the same age, so any observed variation is due to expressed phenotype, not development.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Use a hand lens to count the number of trichomes found on the petiole of the lowest leaf, not including the round, first-grown cotyledons for each plant. Note: It can be helpful to shine a light onto the plant and to hold it against a dark background, such as a lab bench, during the trichome analysis. 
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Both partners will conduct counts on each plant independently and compare answers. If the counts are different, enlist a third counter or take the average of the two results.
</div></li><li data-sub-note-item="1-6"><div class="lab-step">After this, determine the median number of trichomes for the entire class.  <a target="__blank" style="display:block;padding:20px;" href="https://www-jove-com.bdigitaluss.remotexs.co/CDNSource/lm/labs/2/Lab_2_Table_A.xlsx">Click Here to download Table A</a>
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Determine the top and bottom 10% of the plants from the class trichome count. These groups will make up the two artificially selected lines: high hairiness and low hairiness, respectively.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">If more than 10% of the plants have no trichomes, select 10% of those plants randomly for the low hairiness plant line.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">Separate the two groups of selected plants and label them clearly.
</div></li><li data-sub-note-item="1-10"><div class="lab-step">Move the plants to a well-lit area and keep them well watered until they develop flowers. Note: Flowers typically develop around 14 days.
</div></li><li data-sub-note-item="1-11"><div class="lab-step">When the flowers are fully developed, use a pollination wand or paintbrush to rub the stigma and anther of one plant&#8217;s flowers for a few seconds.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">Using the same wand or brush, rub the inside of each flower of every other plant in the same selection line.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">Using a different wand or brush, repeat steps 11 through 12 on the other line. Note: Take care not to cross the lines. Spread the pollen only among the appropriately selected plants.
</div></li><li data-sub-note-item="1-14"><div class="lab-step">Return the generation one plants to the growing areas. When the plants reach approximately 1 month old, they should produce mature seed pods.
</div></li><li data-sub-note-item="1-15"><div class="lab-step">Allow the plants to dry out once the seed pods turn yellow.
</div></li><li data-sub-note-item="1-16"><div class="lab-step">Break the seed pods open and collect the seeds.
</div></li><li data-sub-note-item="1-17"><div class="lab-step">Obtain new pots for the generation two seeds.
</div></li><li data-sub-note-item="1-18"><div class="lab-step">Fill the pots with moist potting soil about halfway and add a fertilizer pellet to each pot.
</div></li><li data-sub-note-item="1-19"><div class="lab-step">Fill the pots with more soil to fill just below the lip.
</div></li><li data-sub-note-item="1-20"><div class="lab-step">Make a shallow depression in the soil of each pot with a probe.
</div></li><li data-sub-note-item="1-21"><div class="lab-step">Touch a clean toothpick to a gluestick and use the toothpick to transplant the seeds into the soil one by one.
</div></li><li data-sub-note-item="1-22"><div class="lab-step">Cover the seeds with a thin layer of soil.
</div></li><li data-sub-note-item="1-23"><div class="lab-step">Label the pots with their hairiness line and place them in the designated growing area.
</div></li><li data-sub-note-item="1-24"><div class="lab-step">Use a plastic dropper to water the plants as needed to ensure they do not dry out.
</div></li><li data-sub-note-item="1-25"><div class="lab-step">When the plants are two weeks old, collect five plants from each line and label them A to E.
</div></li><li data-sub-note-item="1-26"><div class="lab-step">Then, count the trichomes on these plants as previously described (steps 4- 5).
<a target="__blank" style="display:block;padding:20px;" href="https://www-jove-com.bdigitaluss.remotexs.co/CDNSource/lm/labs/2/Lab_2_Table_B.xlsx">Click Here to download Table B</a>
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Data Analysis
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">Create a frequency graph based on the generation one class data with the trichome count on the x axis and the number of plants on the y axis. Label the mean. 
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Then, create frequency graphs for both of the selected hairiness lines.
</div></li><li data-sub-note-item="2-3"><div class="lab-step">Compare the frequency graphs for generation one and both hairiness lines.
</div></li><li data-sub-note-item="2-4"><div class="lab-step">Take note of phenotypic shifts observed in the selected hairiness lines.
</div></li></ul></li></ol>

Artificial Selection: Visualization in Plants and Data Analysis - Procedure

Sztuczna selekcja

Observing Artificial Selection in Plants 
ExpandTo begin the experiment, collect five two-week old grown plants from the generation one population that ...

Evolution

Genetics of Organisms

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">REF and WEF Fly Genetics
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">To prepare for the experiment, be sure to obtain cultures of <em>Drosophila melanogaster</em> at least two weeks ahead of the planned experiment day. NOTE: Because adult females store sperm after mating, you will need this time to generate virgin females by breeding the flies.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">You should get two stock cultures: wildtype, that is homozygous dominant red eyed flies, and white chromosome one mutant, which are homozygous recessive white eyed flies.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Purchase or create <em>Drosophila </em>habitats consisting of a clean vial, 3 cm of prepared food media, and a stopper to close the open end of the vial.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">When not being moved or counted, keep the flies in an incubator or room kept at 23 - 25 &#176;C, where the lifecycle will take around 12 days. NOTE: From the egg, <em>Drosophila </em>progress through four main stages of development: embryo, larva, pupa, and adult. The complete lifecycle takes about 10 days at 25 &#176;C, but it can be affected by temperature. For example, at 18 &#176;C, the lifecycle is about 19 days and at 29 &#176;C, the lifecycle is only seven days.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">As an overview, to set up for the student experiment, you will generate vials for each student group containing white eyed virgin female flies only, white eyed males only, red eyed virgin females only, and red eyed males only. These will be referred to as the experimental vials.
</div></li><li data-sub-note-item="1-6"><div class="lab-step">When your vials from the supplier arrive, referred to as the stock vials, these will contain a mix of male and female flies, as well as larvae, pupae, and eggs.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Beginning with your stock vials, you will take around 3 - 5 males and the same number of females, keeping white eyed and red eyed flies separate at all times, and place these together into a new nursery vial containing food. NOTE: You can sex the flies using a magnifying glass. Look at the posterior of the fly. Males are slightly smaller and have darker solid pigment on the end of their abdomen. Females are larger and have stripes rather than a solid spot.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">You will leave the flies in these nursery vials for around 3 - 5 days to mate and lay eggs.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">After this, you will move them back to their stock vials.
</div></li><li data-sub-note-item="1-10"><div class="lab-step">Next, you will observe as the eggs and larvae in the nursery vials mature, checking the vials every 6 - 8 hours. As it takes females around 8 - 10 hours to become sexually mature, this should ensure that any females found in the nursery vials are virgin.
</div></li><li data-sub-note-item="1-11"><div class="lab-step">As the new adult flies appear, you should transfer them from the nursery to the appropriate experimental tube. For example, if it is a male, transfer it to the experimental male tube for the appropriate eye color. If it is a female, it should be placed in an experimental vial labeled with eye color and virgin female.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">Keep adding flies to the appropriate vials until each experimental vial has at least five flies. NOTE: You will need to make enough experimental vials so that each student group has one of each eye color type and sex, with a few class spares. You should create enough nursery vials to account for this. A good rule of thumb is that it generally takes about two weeks for 25 healthy adults to produce about 50 new adults.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">To begin the process, first label empty habitats to make the nursery vials to raise larvae and pupae for each strain of flies: red eyed and white eyed.
</div></li><li data-sub-note-item="1-14"><div class="lab-step">To move the adult flies into the nursery vial to lay eggs, use an anesthesia wand according to the manufacturer&#39;s instructions, placing it into the stock vial of mature flies. Wait until the animals become unconscious. Now they can be gently tapped onto a piece of white paper and sorted into piles of the same sex using a brush.
</div></li><li data-sub-note-item="1-15"><div class="lab-step">Transfer a few male and female flies to the appropriate vial labeled with their eye color phenotype and sex.
</div></li><li data-sub-note-item="1-16"><div class="lab-step">Place the nursery vials to incubate at 23 to 25 &#176;C and the flies will lay eggs.
</div></li><li data-sub-note-item="1-17"><div class="lab-step">Remove the adults after 3 - 4 days using anesthesia and put them back into the stock vial.
</div></li><li data-sub-note-item="1-18"><div class="lab-step">Remove any adult flies that have eclosed in the 6 - 8 hour period using an anesthesia wand.
</div></li><li data-sub-note-item="1-19"><div class="lab-step">Then, tip out the adults onto paper and sort them into groups based on their sex before placing each fly into an appropriate experimental vial marked male or virgin female and red or white for the phenotype. NOTE: Each experimental vial should contain five flies. Once this number is reached, then a new vial should be started. If you miss the 6 - 8 hour window, females may not be virgins, so these should be discarded into a morgue tube containing 70% ethanol or returned to the stock vial. Males do not need to be virgins, but do need to be kept in male only containers.
</div></li><li data-sub-note-item="1-20"><div class="lab-step">Each lab group will need one tube of five virgin red eyed females, one of five virgin white eyed females, one tube of red eyed males, and one of five white eyed males. In addition, they will need a morgue vial containing ethanol, two anesthesia wands, fly anesthetic, a paintbrush, a magnifying glass, some plain paper, and two empty fly vials for crosses.
</div></li></ul></li></ol>

Genetics of Organisms: REF and WEF Fly Genetics - Prep

Genetyka organizmów

REF and WEF Fly Genetics
ExpandTo prepare for the experiment, be sure to obtain cultures of Drosophila melanogaster at least two weeks ahead of the ...

Genetics

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.bdigitaluss.remotexs.co/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.bdigitaluss.remotexs.co/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.bdigitaluss.remotexs.co/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.bdigitaluss.remotexs.co/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>

Pomiar różnorodności biologicznej

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