<p class="jove_content">Supported by R&#252;diger Foundation grants in 2008 &#38; 2010 (BVS), BONFOR/Gerok Scholarship O-137.0015 (BVS), BONFOR/Gerok Scholarship O-137.0019 (FT), Deutsche Forschungsgemeinschaft/ DFG (BVS) STA 1135/2-1, Chinese Scholarship Council No. 2008627116 (ZL) and an unrestricted grant by Geuder AG, Heidelberg (Fig. 2). Members of H. Skottman&#39;s laboratory, University of Tampere Finland are gratefully acknowledged for providing hES derived RPE shown in Fig 2.&#160;</p>

<p>Ethics of animal handling in ophthalmic research: We obtained approval from the Ethics committee of the Medical Faculty, University of Bonn, and adhere to the guidelines stated by The Association for Research in Vision and Ophthalmology (ARVO). Moreover, all procedures were approved by the state regulatory authorities of North Rhine-Westphalia. Animals were held indoors in a specialized facility in an air-conditioned room with temperatures between 18 - 20 &#176;C, exposure to regular daylight, in standardized individual cages with free access to food and water.</p>
<p></p>
<p>Note: To ensure the animals operative affinity, an animal health score sheet is followed which includes the following definitive animal exclusion criteria: 20% weight loss compared to weight on admission; apparent cyanosis of the animal; animal shivers, has cramps or cannot move in coordination; ataxia/ paresthesia, <em>e.g</em>., paralyses; apathy; extreme auto mutilation (skin wounds, severed limbs).</p>
<p></p>
<p class="jove_title">1. Instrument Sterilization</p>
<ol>
	<li>Place reusable instruments in ultrasonic bath.</li>
	<li>Add 500 ml distilled water and 2 ml of instrument disinfectant.</li>
	<li>Clean instruments using sweep function for 15 min.</li>
	<li>Remove instruments from ultrasonic bath and rinse thoroughly using distilled water for 5 min.</li>
	<li>Insert instruments in autoclave and use the standard program (sterilization of instruments under 121 &#186;C for 20 min).</li>
</ol>
<p class="jove_title">2. Instrument Preparation</p>
<ol>
	<li>Establish and maintain a sterile field, by working in a closed room, wearing surgical scrubs, mask and hair cover. Disinfect hands prior to wearing sterile surgical gloves. For detailed approach see <sup>28</sup>.</li>
	<li>Place sterilized instruments on a sterile drape.</li>
	<li>Place 1 ml syringe filled with 40 mg triamcinolone attached to a 27 G needle for injection, 10 ml syringe with Balance salt solution (BSS), and 5 ml syringe of lubricant on drape.</li>
	<li>Place 3-0 silk, 7-0 vicryl, ocular sticks (to stop conjunctival/ scleral bleeding), twister gauze sponges, wound closure strips (to fixate the vitrectomy tip tubing), and chandelier endoillumination fiber wire on a drape.</li>
	<li>Unwrap 25 G chandelier endoilluminator and connect to light machine using sterile techniques (see step 2.1). Connect vitrectomy set including high speed vitrector and Venturi cassette to vitrectomy machine using sterile techniques (see step 2.1).</li>
	<li>Open 500 ml BSS bottle and connect solution to Venturi cassette according manufacturer&#39;s instructions.</li>
</ol>
<p class="jove_title">3. Preparation of Anesthesia and Positioning of the Animal</p>
<ol>
	<li>Weigh animal to ensure accurate medication dosage.</li>
	<li>Prepare intramuscular (IM) anesthesia using 1 syringe with 27 G needle containing 0.35 mg/kg ketamine and 0.25 mg/kg medetomidine for start. Flip the syringe to mix.</li>
	<li>Prepare 2 syringes with 1/3 the doses to maintain the anesthesia during the operation.</li>
	<li>Prepare syringe containing 20 ml of 5% glucose solution and 18 G needle for subcutaneous injection as an intravenous infusion alternative.</li>
	<li>Give 3 x 1 drop of mydriatic eye drops prior to vitrectomy for pupil dilatation.</li>
	<li>Cover rabbit with blanket to calm before anesthesia injection, inject in the hind limb (gluteal muscle) and massage around injection site for 30 sec.<br />
	Note: The first shot of IM anesthesia lasts about 1 - 2 hr. depending on the rabbit&#39;s size, drug tolerance, fat layer, stress and body temperature. The first sign of the anesthesia fading away is a nystagmus (must be monitored by the surgeon), subsequent injections last about 30 - 45 min.</li>
	<li>Confirm proper anesthesia, by verifying hypnosis, hyporeflexia, analgesia and muscle relaxation of the animal.</li>
	<li>Give subcutaneous injection (Pt. 3.3) in neck skin fold, once the rabbit is unconscious.</li>
	<li>Add methylcellulose lubricant every 5 - 10 min in operated eye, add lubricant and tape lids in non-operated eye.</li>
	<li>Place the covered rabbit on the surgical table draped with a cover such as a cotton blanket in an optimal position (nose slightly elevated through a mold of the blanket, so it is level with ocular surface) under surgical microscope. Align eye perpendicular to microscope objective.</li>
	<li>Ensure proper body core temperature using rectal thermometer (normothermia 39 &#177;&#160;1 &#186;C)<sup>29</sup>.</li>
	<li>Cut eyelashes using scissors (some ointment on blade) to reduce postoperative infections.</li>
	<li>Disinfect the eye using 2 - 3 drops of 0.1 g/ml Povidone-Iodine topically for 1 min and rinse with sterile BSS.</li>
	<li>Cover eye with sterile drape with pre-cut opening in the middle for the eye and then cover with (sticky) surgical incision drape 12 x 17 cm.</li>
</ol>
<p class="jove_title">4. Vitrectomy</p>
<ol>
	<li>Proptose and secure eye with 3-0 silk using inverted caliper and, perform a conjunctival peritomy.
	<ol>
		<li>Incise the conjunctiva with a vannas scissor close to limbus but far enough from the blood vessels (~1 mm distance).</li>
		<li>Dissect the conjunctiva by creating a &#34;T-cut&#34;. First enlarge the peritomy with the scissor parallel to limbus and then incise the conjunctiva vertically in form of a &#34;T&#34; for about 6 - 7 mm. Carefully separate the conjunctiva bluntly.</li>
	</ol>
	</li>
	<li>Perform a sclerotomy using a 23 G microvitreoretinal (MVR) blade at 8 o&#39;clock on right eye/OD (4 o&#39;clock on left eye/OS) by carefully inserting the sharp tip of the blade in direction towards the optic nerve. Slowly retract the blade in the same direction and avoid enlarging the sclerotomy.</li>
	<li>Insert and suture custom side port-infusion cannula<sup>27</sup> using 7-0 silk suture and set intraocular pressure (IOP) at 24 mmHg.</li>
	<li>Perform a sclerotomy with 25 G flat head trocar at 2 o&#39;clock on OD (10 o&#39;clock on OS) similar to step 4.2.</li>
	<li>Insert 25 G chandelier light into flat head trocar, fixate with sticky tape and turn on light source at <em>ca.</em> 30%.</li>
	<li>If needed, remove edematous corneal epithelium using a #20 scalpel for better intraocular visualization.</li>
	<li>Perform a sclerotomy similar to step 4.2 at 10 o&#39;clock on OD (2 o&#39;clock on OS), (pre-) place u-shaped 7-0 sutures around sclerotomy without tying the knot, and insert vitrectomy cutter tip.</li>
	<li>Start vitrectomy<sup>30</sup> around the entry port, then continue over the optic disc and the fibrae medullares using high speed vitrector by cutting the vitreous humor into small pieces at max. 2,000 - 3,000 cuts/min, aspirating at max. 200 mmHg using the stated parameter setup of the vitrectomy machine (<strong>Table 1</strong>)</li>
	<li>Perform a posterior vitreous detachment (PVD) by separating the vitreous humor from the retina by holding the high speed vitrector over the posterior pole and (if feasible gently) superior of the disc<sup>31</sup> while aspirating only at max. 200 mmHg without cutting.</li>
	<li>Inject <em>ca.</em> 50 &#181;l (20 mg) triamcinolone or diluted fluorescein (<em>ca.</em> 0.1 mg/ml) intravitreally to visualize and facilitate (near total) removal of the floating vitreous over the posterior pole and midperiphery during vitrectomy. Avoid crossing over under the lens. Indentation to shave peripheral vitreous by a (skilled) assistant is recommended if gas tamponade is desired.</li>
	<li>Add 20 units/ml heparin and 0.5 mg epinephrine to final concentration of 0.001 mg/ml into the BSS infusion solution in parallel or after step 4.10.<br />
	Note: As heparin/epinephrine are not injected intraocular their effects are delayed depending on infusion flow rate.</li>
</ol>
<p class="jove_title">5. Loading Shooter</p>
<p>Note: The work described herein does not fall under the tenets of the Declaration of Helsinki; it did not involve human patients. Here, standard RPE cells were isolated from fetal human eyes, cultured and differentiated on uncoated 10-&#181;m-thick polyester (PET) inserts according to our previously published protocol<sup>14</sup>. A permission to work with the human fetal material was obtained from the ethics committee of the University of Bonn. Alternatively, hES-RPE were shipped from the Skottman lab (manuscript in prep.), where they were cultured according to the technique described by Vaajasaari <em>et al</em>.<sup>32</sup>; for these cells a permission has been obtained from the R. Koch Institute, Berlin, Germany.</p>
<ol>
	<li>Rinse cell culture prior to preparation of the implant 3x with ophthalmic grade BSS.</li>
	<li>Fill a standard cell culture dish (100 x 20 mm) with 10 ml ophthalmic grade BSS.</li>
	<li>Add the cell culture insert into the BSS and center the dish under a light microscope.</li>
	<li>Punch out a 2.4 x 1.1 mm implant with a blunt, oval, custom-made needle to obtain a flat, bean-shaped substrate with two long edges and two round edges.</li>
	<li>Gently flood the needle through the second port with BSS to flush out the implant into the BSS filled custom made loading station (<strong>Figure 1</strong>).</li>
	<li>Optionally cut one round end of implant (&#60;0.5 mm), just to obtain a third edge.</li>
	<li>Make sure that the implant is in the right orientation by ensuring that the monolayer is upside on the cell carrier. To change positioning carefully use two scalpels.</li>
	<li>Push the implant gently and completely into the shooter instrument using the needle holder until all of the implant is secured inside of the tip. The plunger should remain retracted.</li>
	<li>Keep the &#34;loaded&#34; shooter tip in loading station under BSS until the moment of implantation.</li>
</ol>
<p class="jove_title">6. Implantation</p>
<ol>
	<li>Approach neural retina with extendible 41 G subretinal injection needle connected to a gastight syringe (ensure that all air bubbles have been evacuated from tubing!).</li>
	<li>Inject BSS (with calcium and magnesium/ CM) subretinally and thereby create a bleb retinal detachment (bRD) of about 2 - 3 disc diameter (DD). Two bRD per eye can be raised safely.</li>
	<li>Enlarge retinotomy to 1.5 mm with vertical 23 G VR-scissors. The subretinal space is now accessible for implantation or further maneuvering.</li>
	<li>Extend sclerotomy (precisely) with a 1.4 mm incision knife to 20 G approach.</li>
	<li>Attempt passing through the sclerotomy using a 20 G shooter dummy, enlarge as needed to ensure smooth, yet snug transition of the loaded shooter.</li>
	<li>Pass with the loaded shooter<sup>27</sup> through sclerotomy ideally at 24 mmHg.</li>
	<li>Approach retinotomy edge and eject the implant subretinally from an epiretinal position.</li>
	<li>Adjust the implant with half-closed 23 G scissors, forceps or 41 G needle to make sure it is positioned well under the retina- reasonably away from the retinotomy.</li>
</ol>
<p class="jove_title">7. Ending Operation</p>
<ol>
	<li>Remove 25 G chandelier and infusion cannula.</li>
	<li>Suture all sclerotomies.</li>
	<li>Inject 25 &#181;l (10 mg) triamcinolone by the 8 o&#39;clock sclerotomy (prior to suturing last sclerotomy).</li>
	<li>Check/adjust IOP by palpation and inject BSS via 30 G needle/syringe, if needed.</li>
	<li>Suture conjunctiva with 7-0 vicryl.</li>
	<li>Remove proptosing 3-0 silk sling slowly (Avoid deep orbital venous plexus!).</li>
	<li>Add dexamethasone/antibiotic ointment under lid.</li>
	<li>Position for 1 hr covered with blanket with operated eye facing up (w/o gas), or down (with air/gas).</li>
	<li>Do not leave animal unattended until it regains sufficient consciousness to maintain sternal recumbence.</li>
	<li>Do not transport rabbit before the anesthesia fades completely, this can be speeded up by injecting medetomidine reversing agent equal to amount of medetomidine given.</li>
</ol>
<p class="jove_title">8. Post-operative Animal Care</p>
<ol>
	<li>Keep rabbits under appropriate conditions (temperature, light, food, water, space, <em>etc</em>.) and close monitoring in a specialized facility.</li>
	<li>Ensure that animal is well rested, <em>i.e</em>., no extended periods of food or water deprivation.</li>
	<li>Look for any wounds or injuries, especially on injection sites.</li>
	<li>Keep wounds dry to prevent infections. Give antibiotics when infection is suspected: dexamethasone 1 mg/g, neomycin sulfate 3,500 I.U./g, polymyxin B sulfate 6,000 I.U./g ointment was applied twice daily for 1 week postoperative onto the ocular surface.</li>
	<li>Add dexamethasone/antibiotic ointment for next 7 days postoperative twice daily for better ocular surface regeneration and reduced post-operative pain.</li>
	<li>Give systemic analgesics (Carprofene 4 mg/kg twice daily) for initial 48 hr.</li>
	<li>Do not leave an animal unattended until it has regained sufficient consciousness to maintain sternal recumbence.</li>
	<li>Do not return an animal that has undergone surgery to the company of other animals until fully recovered.</li>
	<li>Do not expose animals to unnecessary distress.</li>
</ol>
<p class="jove_title">9. SLO/SD-OCT Guidance</p>
<ol>
	<li>Prepare and inject intramuscular (IM) anesthesia using 1 syringe with 27 G needle containing 0.175 mg/kg ketamine and 0.125 mg/kg medetomidine for start. Flip the syringe to mix.</li>
	<li>Add a lubricant at least every 5 min to moisturize eye and maintain clear SD-OCT imaging, optionally a custom contact lens may be used.</li>
	<li>Attach a steel platform to the headrest to stabilize the animal in the required position.</li>
	<li>Place rabbit on steel platform, placing its eye perpendicular to probe.</li>
	<li>Hold the rabbit&#39;s head from the lower cheek bone (mandibula) avoiding the trachea.</li>
	<li>Tilt animal&#39;s head by roughly 45&#186; towards the SD-OCT probe to optimize the viewing angle on the implant.</li>
	<li>Use a 30-degree lens and the following parameters for optimal OCT imaging [HS]: 30 degrees settings for single line scans with ART mode set to 100 (averaging) and 20 x 20 degree settings for volume scans with ART mode set to 15; high resolution mode is not required.</li>
	<li>Use SLO infrared reflectance imaging to find the focal plane of the implant (<strong>Fig. 2A</strong>); optimal focus is reached when (all) the implant edges are sharp<sup>14</sup>.</li>
	<li>Add dexamethasone 1 mg/g, neomycin sulfate 3,500 I.U./g, polymyxin B sulfate 6,000 I.U./g ointment under lid when finished.</li>
</ol>

<p class="jove_content">RB, BVS, ZL, NE and Geuder AG have filed a European patent application on the shooter. NB is an employee of Geuder AG. Publication fees for this video-article were&#160;paid by Geuder AG.</p>

<p class="jove_content">Using a rabbit model, a safe and reproducible method is presented for transvitreal delivery of cultured RPE on cell carriers into the subretinal space with a custom-designed shooter instrument. The described method offers a short/optimized surgical technique for easy learning, as it involves standard techniques in vitrectomy with subretinal maneuvers. Outcome is greatly facilitated by a clean vitreoretinal interface, and intraocular infusion that avoids fluid turbulence over the implantation site, inducing bleb retinal detachment (bRD) at low IOP, preventing retina and sclera damage through dryness, and appropriate positioning of the rabbit.</p>
<p class="jove_content">We caution however, as several intra-operative complications may occur any time, hindering the success of the implantation, for instance intra ocular bleedings, anesthesia fading off during vital steps such as the implantation, collapsing of bRD due to instrument manipulation or ocular hypotony, rabbit death due to excessive doses of anesthesia, low blood pressure during long operation causing hypoxic brain damage, or hyperthermia. Yet these complications decrease with time as they are quickly tackled and resolved by increasing experience of the surgical team.</p>
<p class="jove_content">Some complications could be reduced by following a few simple, yet crucial steps. Lubricant should be added every 5 - 10 min to prevent corneal, scleral and conjunctival damage during the operation, and to maintain a clear intraocular media, as dried/blackened sclera may be a cause of wound dehiscence, which in turn leads to ocular hypotony and/ or intraoperative leakage from sclerotomies. Heparin should be added to prevent the formation of a fibrin film that makes particularly subretinal implantation challenging and simultaneously adding epinephrine to reduce bleeding under heparin<sup>16</sup>. Too long heparin/ epinephrine exposure times (&#62;1 hour) should be avoided to prevent corneal edema by endothelial decompensation<sup>33</sup>, hypertensive crisis or intraoperative fatality. Meticulous vitreous removal should be performed at instrument (entry) port to avoid retinal and/ or choroidal detachments. Intraocular instruments should be pointed towards posterior pole to avoid lens touch (causes iatrogenic cataract formation) or (entry site) retinal damage. An intraocular side-port infusion cannula should be used, as it attenuates the jet stream around the implantation area, thus preventing uncontrolled tearing of the retinotomy, and collapse of the bRD. bRD induction in the midline (vertical axis from optic nerve) or close to optic medullar fibers should be avoided to prevent extensive iatrogenic retinal detachments. Finally, last but not least bRD should be induced at low IOP, to avoid subretinal BSS injection using excessive flow rates which may lead to a retinal damage (<em>e.g</em>., by stretching).</p>
<p class="jove_content">Many study variables such as cell carrier variants, fetal, adult or stem cell derived RPE cell sources, choices for immunosuppressants, <em>etc</em>., can be explored <sup>14,26,27,34</sup>. Further improvement such as serum-free RPE culture methods, characterization of xenoRPE in subretinal space, removal of the host RPE layer<sup>14</sup> or strategies for implant anchorage are current work in progress.</p>
<p class="jove_content">To date the described techniques have been used on 5 different rabbit strains, including chinchilla bastard, Chinchilla bastard/KBL hybrids, New Zealand White/Red Cross, New Zealand White (albino) and Dutch belted. Both male and female rabbits were operated on, with rabbits at least 1.5 kg or 2 months of age (depending on species). Most surgeries were on pigmented rabbits (chinchilla bastard or chinchilla bastard hybrids) with weights between 2.5 - 3 kg.</p>
<p class="jove_content">All of the rabbit strains we have had the opportunity to work with seem to have some peculiarities. Given the exclusive availability of pigmented rabbits of the chinchilla bastard strain in Germany in 2009-13, we have collected the most experience with these animals. Unfortunately it is no longer available, since breeding has been discontinued, but compares very well to New Zealand White/Red Cross except for the more advantageous thicker sclera and larger eye volumes in the latter. Chinchilla bastard hybrids have significant intraoperative fibrin formation and require heparin/epinephrine use as outlined above to ensure successful subretinal maneuvers. This protocol has also been performed in non-pigmented albino rabbits (New Zealand White), however particularly bRD creation and subretinal implantation is more challenging given reduced contrast appreciation. The feasibility of inducing a posterior vitreous detachment did not seem rabbit strain dependent in our hands.</p>
<p class="jove_content">Transvitreal subretinal delivery is likely the future surgical strategy of choice given it is the most common route nowadays clinically to access the retina. As a result many other groups have presented such techniques for cultured RPE on carrier supports in animal work<sup>11,15,23,35</sup>. Aramant <em>et al</em>.<sup>36</sup> have an instrument, which places rather than pushes their hydrogel-encapsulated soft implant to its subretinal target site. The design of Thumann <em>et al.</em> utilizes a hollow spatula, which releases the carrier-supported graft by floating it off through fluid injection <sup>19</sup>. Both former strategies require subretinal insertion of the instrument, which in our view is more prone to complications, when compared to an epiretinally appositioned instrument. Montezuma <em>et al</em>.<sup>22</sup> described a subretinal inserter instrument for the delivery of subretinal chip implants in pigs but no further work has been published since to the best of our knowledge. We have been able to extend the described technique with some modification to pig.</p>
<p class="jove_content">Our preferred cell carriers are 10 micron thick polyester terephthalate (PET) membranes. From a surgical perspective, this material has favorable stiffness and elasticity parameters, in addition to its broad versatility during cell culture experiments. We found similar experiences with expanded tetrafluoroethylene (ePTFE) <sup>37</sup> or nanofiber membranes electrospun from PET, poly-lactic/capronolactic acid (PLCL) or poly--lactic-co-glycolic acid (PLGA), as well as composite nanofiber (PLGA or PET) and ultrathin PET <sup>26</sup>. When PET membranes are used with our metallic shooter instrument, they do have an occasional tendency to exhibit electrostatic charge, which challenges their ejection from the shooter <sup>27</sup>. Ultrathin polyimide membranes could in our hands not be implanted in the subretinal space with the protocol outlined above (manuscript in preparation).</p>
<p class="jove_content">Marmor <em>et al</em>. have systematically studied spontaneous resorption of subretinal fluid in iatrogenic localized retinal detachments<sup>38-41</sup>. Even following manipulation in the subretinal space these were found to be reabsorbed by postoperative day 4 in uneventful surgeries. Laser retinopexy is not performed to secure the edges of the retinotomy. Although counterintuitive when compared to human surgery, air/ gas tamponade is not required. Unless meticulous removal of peripheral vitreous can be achieved, particularly in the superior quadrant, this may in fact result in giant retinal tears originating from the retinotomy site. It is only recommended to perform fluid air exchange with subsequent 20% SF6 gas tamponade to salvage intraoperative iatrogenic retinal detachments or in case a particular implant position needs to be secured.</p>
<p class="jove_content">Although the mechanically induced ablation of the neural retina can cause RPE and photoreceptor damage in rabbits<sup>42,43</sup>, its extent varies greatly (even with regular BSS) depending on factors such as IOP, syringe type used, injection volume with thereby induced retinal stretching, <em>etc</em>. We have also tested the often recommended Ca/Mg-free BSS facilitated detachment <sup>42-44</sup>, but found that it causes intraoperative lens opacification (particularly with elevated temperature), and significantly delays or even impairs retinal re-attachment <sup>27</sup>. Slow subretinal injection of 20 - 30 &#181;l volume of regular BSS with a 100 &#181;l syringe is therefore recommended; injection needle movements should be minimal so the retinotomy seals around it and prevent Bruch&#39;s membrane damage. Some of the iatrogenic damage may be resolved by RPE wound healing, and the observed relative preservation of ONL thickness after reattachment, suggests that the RPE/photoreceptor complex can tolerate this impairment, as also described by others<sup>45</sup>.</p>
<p class="jove_content">Cell-based therapeutics or retinal prosthetics require preclinical animal testing prior to regulatory approval and commencing human safety studies. The former vary from country to country. The rabbit model described here may serve as a cost-efficient and less challenging platform for establishing or even carrying out all requirements by regulatory authorities. Moreover, it may subsequently serve for training of surgeons in eventual multicenter clinical trials or further improvements of the technique along the way.</p>

<p class="jove_content">Age-related macular degeneration (AMD) is the most common cause of visual impairment in adults aged 50 or older in industrially developed countries, as it causes loss of central vision. About 15% of these patients suffer from the &#34;wet&#34; form of the disease, in which neovascularization originates from the choroid and disrupts retinal function<sup>1</sup>. This variant can be treated by a highly effective therapy with repeated intra-vitreal injections of antiangiogenic drugs<sup>2</sup>. However, the vast majority of patients (~85%) suffer from the dry form, which is characterized by extracellular deposits (<em>e.g.</em>, drusen) under the retinal pigment epithelium (RPE). These deposits cause RPE dysfunction leading to retinal atrophy in the macula. Given the lack of any curative therapeutic options, AMD evolved into an intensively developing research field, where many different curative therapeutic approaches are being tested. Surgical RPE replacement is one attractive future possibility to defeat this debilitating disease<sup>3</sup>.</p>
<p class="jove_content">Autologous subretinal RPE transplantation replaces dysfunctional or lost RPE in macula, and has the potential to restore its physiological function<sup>4-9</sup>. This surgical technique had a breakthrough with the development of RPE differentiation protocols from human embryonic stem cells (hESC) and induced pluripotent stem cells (iPSC), giving the scientist an unlimited cell source of RPE for transplantation<sup>10</sup>. RPE transplantation is now recognized as an attractive first-in-human application for stem cell derived therapeutics. The eye offers excellent surgical access and sophisticated <em>in vivo</em> monitoring tools<sup>11-13</sup>.</p>
<p class="jove_content">To transplant the RPE, one way is with a minimally invasive delivery using a cell suspension, alternatively, to better preserve RPE characteristics and transplant function, arti&#64257;cial carrier substrates (scaffolds) for RPE replacement are being considered<sup>4,14,15</sup>. Large animal models are required for preclinical validation, yet detailed technical information on animal handling and surgical technique is missing to date<sup>16-23</sup>.</p>
<p class="jove_content">We and others<sup>11,24</sup> despite some evidence to the contrary<sup>25</sup>, suggest the use of a rigid yet elastic carrier substrate as it provides safer handling, preserves monolayer integrity and functionality. Over time we have tested several custom-designed instruments and ancillary techniques for the implantation of cell-carrier supported RPE transplants into the subretinal space (SRS). We utilized intraoperative video recordings, <em>in vivo</em> scanning laser ophthalmoscopy combined with spectral domain optical coherence tomography (SLO/ SD-OCT), and histology to evaluate the implantation success<sup>14,26,27</sup>. Here we provide our current recommendation for subretinal RPE implants in rabbits, which were tested in 5 different rabbit strains, 7 cell carrier materials and 4 RPE cell sources in over 150 procedures.</p>

<table><tbody><tr><td>s30 ultrasonic cleaning unit</td><td>Elmasonic</td><td>100 4631</td><td>2.75 L</td></tr><tr><td>DE-23&amp;nbsp; autoclave</td><td>Systec</td><td>C 2209</td><td>23 L</td></tr><tr><td>Syringe&amp;nbsp;</td><td>BD&amp;nbsp;</td><td>300013&lt;br/&gt; 300995&amp;nbsp;&lt;br/&gt; 301285&lt;br/&gt; 300294&lt;br/&gt; 300330&amp;nbsp;</td><td>1 ml x3&lt;br/&gt; 2 ml x3&lt;br/&gt; 5 ml x1&lt;br/&gt; 10 ml x1&lt;br/&gt; 20 ml x1</td></tr><tr><td>Needle&amp;nbsp;</td><td>BD</td><td>305196&lt;br/&gt; 305136</td><td>18 G x1&amp;nbsp;&lt;br/&gt; 27 G x5</td></tr><tr><td>Scalpel</td><td>Feather</td><td>2975#20</td><td>blade#20 x3</td></tr><tr><td>Surgical drape</td><td>HARTMANN&lt;br/&gt; LOHMANN &amp;amp; RAUSCHER</td><td>277 502&lt;br/&gt; 25 440</td><td>60 x 40 cm x2&lt;br/&gt; 12 x 17 cm</td></tr><tr><td>Ocular sticks</td><td>LOHMANN &amp;amp; RAUSCHER</td><td>16 516</td><td>66 x 5 mm</td></tr><tr><td>Twister gauze sponges</td><td>HARTMANN</td><td>481 274</td><td>x2</td></tr><tr><td>Closure strips</td><td>HARTMANN</td><td>540 686</td><td>x4</td></tr><tr><td>Opmi Visu CS Microscope</td><td>Zeiss</td><td>N/a</td><td>incl. fundus imaging system BIOM II</td></tr><tr><td>Chandelier endoillumination</td><td>Geuder</td><td>G-S03503 +&lt;br/&gt; G-S03504</td><td>25 G incl. trocar</td></tr><tr><td>Light machine</td><td>Geuder</td><td>G-26033</td><td>Xenotron III</td></tr><tr><td>Vitrectomy machine</td><td>Geuder</td><td>G-60000</td><td>MegaTRON S4 S4/ HPS</td></tr><tr><td>Vitrector</td><td>Geuder</td><td>G-46301</td><td>MACH2 vitreous cutter 23 G</td></tr><tr><td>Venturi cassette&amp;nbsp;</td><td>Geuder</td><td>G-60700</td><td></td></tr><tr><td>Sideport-infusion cannula</td><td>Geuder</td><td>custom&amp;nbsp;</td><td>23 G x1</td></tr><tr><td>3-0 silk suture</td><td>ETHICON</td><td>V546G</td><td>x1</td></tr><tr><td>Caliper</td><td>Geuder</td><td>G-19135</td><td>x2</td></tr><tr><td>Vannas scissors</td><td>Geuder</td><td>G-19777</td><td>x1</td></tr><tr><td>Sclerotomie blade</td><td>Ziemer</td><td>21-2301</td><td>1x 23 G&lt;br/&gt; 1x 20 G</td></tr><tr><td>7-0 silk suture</td><td>ETHICON</td><td>EH6162H</td><td>x1</td></tr><tr><td>Needle holder</td><td>Geuder</td><td>G-32320</td><td>x2</td></tr><tr><td>Iris forceps</td><td>Geuder</td><td>G-18910</td><td>x1</td></tr><tr><td>Colibri forceps</td><td>Geuder</td><td>G-18950</td><td>x1</td></tr><tr><td>Extendible subretinal injection needle</td><td>DORC</td><td>1270.EXT</td><td>41 G</td></tr><tr><td>VR scissor</td><td>Geuder</td><td>G-36542</td><td>25 G</td></tr><tr><td>Grieshaber forceps holder</td><td>Alcon</td><td>712.00.41</td><td>23 G</td></tr><tr><td>Curved scissor forceps tips</td><td>Alcon</td><td>723.52</td><td>23 G</td></tr><tr><td>Implant loading station</td><td>Dow Corning</td><td>3097358-1004</td><td>SYLGARD 184 Silicone Elastomer Kit</td></tr><tr><td>Blunt oval implant trephine&amp;nbsp;</td><td>Geuder</td><td>custom-made&amp;nbsp;</td><td>2.4 x 1.1 mm&amp;nbsp;</td></tr><tr><td>Shooter dummy</td><td>Geuder</td><td>G-32227</td><td>x1</td></tr><tr><td>Shooter</td><td>Geuder</td><td>G-S03443</td><td>x1</td></tr><tr><td>Flute needle</td><td>DORC</td><td>1281.SD</td><td>20 G (Vacuum)</td></tr><tr><td>Manual microliter syringe</td><td>Hamilton</td><td>24535</td><td>100 &amp;micro;l</td></tr><tr><td>Tissue culture plates</td><td>Greiner bio-one</td><td>&amp;nbsp;664160</td><td>100 x 20 mm</td></tr><tr><td>Spectralis Multi-Modality&lt;br/&gt; Imaging System</td><td>Heidelberg&lt;br/&gt; Engineering</td><td>N/a</td><td>Spectralis HRA&lt;br/&gt; &amp;nbsp;+ OCT</td></tr><tr><td>&lt;strong&gt;Drugs and solutions&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Active agent&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>Mucadont-IS</td><td>Merz Hygiene</td><td>virucidal instrument disinfectant&amp;nbsp;</td><td>2 L</td></tr><tr><td>Mucocit&amp;nbsp;T</td><td>Merz Hygiene</td><td>Aldehyde-free instrument disinfectant</td><td>2 L</td></tr><tr><td>Ketamin 10%</td><td>WDT</td><td>Ketamine&amp;nbsp;</td><td>10 ml (100 mg/ml)</td></tr><tr><td>Domitor</td><td>Orion Pharma</td><td>Medetomidine hydrochloride</td><td>10 ml (1 mg/ml)</td></tr><tr><td>Antisedan</td><td>Orion Pharma</td><td>Atipamezole hydrochloride</td><td>10 ml (5 mg/ml)</td></tr><tr><td>Neosynephrin POS 10%</td><td>URSAPHARM</td><td>Phenylephrine HCl</td><td>10 ml</td></tr><tr><td>Mydriacyl</td><td>Alcon</td><td>Tropicamid</td><td>10 ml (5 mg/ml)</td></tr><tr><td>Methocel 2%</td><td>Omni Vision</td><td>hydroxypropyl methylcellulose</td><td>10 g</td></tr><tr><td>PURI CLEAR</td><td>ZEISS</td><td>Balance salt solution (BSS)&amp;nbsp;</td><td>500 ml</td></tr><tr><td>Glucose 5%&amp;nbsp;&amp;nbsp;</td><td>B.Braun</td><td>Glucose 5%&amp;nbsp; solution</td><td>100 ml</td></tr><tr><td>Heparin-Natrium-25,000</td><td>Ratiopharm</td><td>Heparin</td><td>5 ml (2,500 unit/ml)</td></tr><tr><td>Suprarenin</td><td>SANOFI</td><td>Epinephrine</td><td>1 ml (1 mg/ml)</td></tr><tr><td>Triamcinolone</td><td>University of Bonn pharmacy</td><td>preservative-free Triamcinolone&amp;nbsp;</td><td>1 ml (40 mg/ml)&amp;nbsp;</td></tr><tr><td>Isoptomax eye ointment</td><td>Alcon</td><td>dexamethasone 1 mg/g&lt;br/&gt; neomycin sulfate 3,500 IU/g&lt;br/&gt; polymyxin B sulfate 6,000 IU/g</td><td>10 ml</td></tr><tr><td>Betaisodona</td><td>Mundipharma</td><td>Povidon-Iod</td><td>30 ml (1 g/10 ml)</td></tr><tr><td>Optive</td><td>ALLERGAN</td><td>sodium carboxymethylcellulose glycerol</td><td>10 ml</td></tr></tbody></table>

a step by step protocol for subretinal surgery in rabbits

<p class="jove_content">Age related macular degeneration (AMD), retinitis pigmentosa, and other RPE related diseases are the most common causes for irreversible loss of vision in adults in industrially developed countries. RPE transplantation appears to be a promising therapy, as it may replace dysfunctional RPE, restore its function, and thereby vision.</p>
<p class="jove_content">Here we describe a method for transplanting a cultured RPE monolayer on a scaffold into the subretinal space (SRS) of rabbits. After vitrectomy xenotransplants were delivered into the SRS using a custom made shooter consisting of a 20-gauge metallic nozzle with a polytetrafluoroethylene (PTFE) coated plunger. The current technique evolved in over 150 rabbit surgeries over 6 years. Post-operative follow-up can be obtained using non-invasive and repetitive <em>in vivo</em> imaging such as spectral domain optical coherence tomography (SD-OCT) followed by perfusion-fixed histology.</p>
<p class="jove_content">The method has well-defined steps for easy learning and high success rate. Rabbits are considered a large eye animal model useful in preclinical studies for clinical translation. In this context rabbits are a cost-efficient and perhaps convenient alternative to other large eye animal models.</p>

<p class="jove_content" fo:keep-together.within-page="1">The results from the described method for subretinal implantation are shown in <strong>Table 2.</strong> Engraftment under the retina had a success rate of <em>ca.</em> 61% when a core vitrectomy was performed and rose up to 76%, when posterior vitreous detachment was induced. These numbers include <em>ca.</em> 21% of animals who died either intraoperatively or in the first 3 postoperative days. This technique can be used to implant two scaffolds on different retinal areas in one eye simultaneously.</p>
<p class="jove_content" fo:keep-together.within-page="1">The rabbits underwent postoperative <em>in vivo</em> follow up using SD-OCT and histologic processing as described by Stanzel <em>et al</em>.<sup>27</sup> (<strong>Figure 2</strong>). <strong>Fig. 2A</strong>&#160;shows scanning laser ophthalmoscope infrared reflectance image of implanted cultured RPE on a polyester membrane (PET) after an uncomplicated transplantation. The halo around implant corresponds to photoreceptor atrophy. <strong>Fig. 2B</strong>&#160;shows corresponding SD-OCT, notice retinal, mainly outer nuclear layer (ONL) thinning, hyper reflective band on SD-OCT above implant, while the neural retina adjacent to the implant shows near-normal reflection bands. These results suggest an atraumatic delivery. <strong>Fig. 2C</strong>&#160;shows Hematoxiline/Eosin (H/E) stain of the implant which shows subretinal scarring and ONL atrophy around the retinotomy site likely as a result of iatrogenic manipulation, a contiguous yet irregular pigmented layer over PET. Bruch&#39;s membrane underneath the implant also appeared to be contiguous, and the choriocapillaris contains some scattered erythrocytes. These morphologic result are comparable to the SD-OCT and strengthen the thesis of an atraumatic delivery.</p>
<p class="jove_content" fo:keep-together.within-page="1"><img alt="Figure 1" src="/files/ftp_upload/53927/53927fig1.jpg" /><br />
<strong>Figure 1: </strong> <strong>Loading Shooter with Human iPSC-RPE Cultured on PET Cell Carrier. A</strong>) Shows punching out an implant using custom made needle. <strong>B</strong>) Bean-shaped implant cut from cell culture. <strong>C</strong>) Positioning of implant before loading. <strong>D</strong>) Loading of shooter with implant. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53927/53927fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a></p>
<p class="jove_content" fo:keep-together.within-page="1"><img alt="Figure 2" src="/files/ftp_upload/53927/53927fig2.jpg" /><br />
<strong>Figure 2: </strong> <strong>Cultured Human hES-RPE on PET Cell Carrier 4 Weeks in Rabbit Subretinal Space. A</strong>) Shows SLO infrared reflectance image, the green line demarcates the cross-section shown in <strong>Fig. 2B</strong>. <strong>B</strong>) Corresponding SD-OCT. <strong>C</strong>) H/E stain, see text or<sup>26,27</sup> for details. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/53927/53927fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a></p>
<table fo:keep-together.within-page="1">
	<colgroup>
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:97px;"><strong>Parameter</strong></td>
			<td style="width:131px;"><strong>Used settings</strong></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Vitrectomy</td>
			<td>6,000 cuts/min</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Vacuum</td>
			<td>200 mmHg</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Rise time</td>
			<td>1 sec</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">&#160;Air</td>
			<td>24 mmHg</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Irrigation&#160;</td>
			<td>24 cmH<sub>2</sub>O</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Diathermy</td>
			<td>30%</td>
		</tr>
	</tbody>
</table>
<p class="jove_content" fo:keep-together.within-page="1"><strong>Table 1:</strong><strong> Parameter Setup of the Vitrectomy Machine.</strong></p>
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td><strong>Vitrectomy</strong></td>
			<td><strong>Implant</strong></td>
			<td><strong>Rabbits operated</strong></td>
			<td><strong>Successful implant</strong></td>
			<td><strong>Failed implant</strong></td>
			<td><strong>Death</strong></td>
			<td><strong>Success rate%</strong></td>
		</tr>
		<tr>
			<td>Core vitrectomy</td>
			<td>PET</td>
			<td>30</td>
			<td>19</td>
			<td>4</td>
			<td>7</td>
			<td>63.33</td>
		</tr>
		<tr>
			<td></td>
			<td>PET + RPE</td>
			<td>70</td>
			<td>42</td>
			<td>12</td>
			<td>16</td>
			<td>60</td>
		</tr>
		<tr>
			<td>PVD, &#177;&#160;plasmin w/PPV</td>
			<td>PET</td>
			<td>28</td>
			<td>21</td>
			<td>2</td>
			<td>5</td>
			<td>75</td>
		</tr>
		<tr>
			<td></td>
			<td>PET + RPE</td>
			<td>22</td>
			<td>17</td>
			<td>2</td>
			<td>3</td>
			<td>77.27</td>
		</tr>
	</tbody>
</table>
<p class="jove_content" fo:keep-together.within-page="1"><strong>Table 2:</strong><strong> Summary of the Last 150 Operations Including Method and Implant Type. </strong></p>

Watch this Scientific Journal Video about A Step by Step Protocol for Subretinal Surgery in Rabbits at JoVE.com

A Step by Step Protocol for Subretinal Surgery in Rabbits

<p class="jove_content">Retinal pigment epithelium (RPE) replacement strategies and gene-based therapy are considered for several retinal degenerative conditions. For clinical translation, large eye animal models are required to study surgical techniques applicable in patients. Here we present a rabbit model for subretinal surgery geared towards RPE transplantation, which is versatile and cost-efficient.</p>

Age related macular degeneration (AMD), retinitis pigmentosa, and other RPE related diseases are the most common causes for irreversible loss of vision in ...

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15.3K Views.  University of Bonn. Retinal pigment epithelium (RPE) replacement strategies and gene-based therapy are considered for several retinal degenerative conditions. For clinical translation, large eye animal models are required to study surgical techniques applicable in patients. Here we present a rabbit model for subretinal surgery geared towards RPE transplantation, which is versatile and cost-efficient.

Video: A Step by Step Protocol for Subretinal Surgery in Rabbits

Extraction

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

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

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<p>Source: Smaa Koraym at Johns Hopkins University, MD, USA</p>
<ol class="note-items">
	<li data-note-item="1" data-parent-collapse=""><strong>Determining the Density of an Egg</strong><button class="expand">Expand</button>
	<p>Density is a measure of how compact a substance or object is, and it is calculated as mass divided by volume. When a dense object is placed in a liquid with a lower density, the object sinks; when the object is less dense than the liquid, it floats. When the densities are the same, the object will be suspended in the liquid. In this experiment, we&#39;ll determine the density of an egg by creating a solution with the same density. We&#39;ll measure the density of the salt solution using different volume measurement glassware and determine which is the most accurate.</p>
	<ul class="lab-items">
		<li data-sub-note-item="1-1">
		<div class="lab-step">To begin, wear the appropriate personal protective equipment, including close-toed shoes, long pants, long-sleeved shirt, lab coat, chemical splash goggles, and gloves.</div>
		</li>
		<li data-sub-note-item="1-2">
		<div class="lab-step">Measure the mass of the egg. Set a weigh boat on the balance and tare it. Carefully set the egg in the weigh boat while you record the mass in your notebook.</div>
		</li>
		<li data-sub-note-item="1-3">
		<div class="lab-step">Prepare the salt solution that will be used to suspend your egg. Use the 600-mL beaker and fill it 3/4 of the way full with deionized water.</div>
		</li>
		<li data-sub-note-item="1-4">
		<div class="lab-step">Gently place your egg in the beaker, and then add about one teaspoon of NaCl into the water. Stir the solution gently until the NaCl dissolves. Continue to add NaCl until the egg floats just below the surface.</div>
		</li>
		<li data-sub-note-item="1-5">
		<div class="lab-step">Use different glassware to measure the volume of the salt solution. Weigh the 50-mL graduated cylinder on the top loading balance and record the mass. Then, measure 20 mL of your salt solution and record the volume in your notebook. <strong>Note: </strong>Place a blank sheet of paper behind the graduated cylinder to read the volume more clearly. Always view the measurement at eye level and measure the volume from the bottom of the meniscus. The lines on the graduated cylinder are 0.2 mL apart, meaning that you can read the volume to the hundredths place.
		<h4><b>Table 1: Measuring Mass</b></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>Container</strong></td>
					<td style="text-align: center;"><strong>Mass of empty container (g)</strong></td>
					<td style="text-align: center;"><strong>Mass of full containter (g)</strong></td>
					<td style="text-align: center;"><strong>Mass of solution (g)</strong></td>
					<td style="text-align: center;"><strong>Percent error</strong></td>
				</tr>
				<tr>
					<td>50-mL graduated cylinder</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td>10-mL volumetric pipette</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td>50-mL beaker</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td>20-mL volumetric flask</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
			</tbody>
		</table>
		<a href="https://jove.unicartagenaproxy.elogim.com/CDNSource/lm/labs/41/Table_1._Measuring_Mass.xlsx" target="_blank">Click Here to download Table 1</a></div>
		</li>
		<li data-sub-note-item="1-6">
		<div class="lab-step">Weigh the filled graduated cylinder and record the mass in your notebook. Then, pour the salt solution back into the 600-mL beaker.</div>
		</li>
		<li data-sub-note-item="1-7">
		<div class="lab-step">Use the volumetric pipette to measure volume. First, weigh the empty 50-mL beaker and record the mass in your notebook. Then, carefully measure 10 mL of the salt solution using the volumetric pipette and dispense the salt solution into the beaker.</div>
		</li>
		<li data-sub-note-item="1-8">
		<div class="lab-step">Measure another 10 mL and add it to the beaker. Weigh the filled beaker and record the mass in your notebook. Then, empty, rinse, and dry the beaker.</div>
		</li>
		<li data-sub-note-item="1-9">
		<div class="lab-step">Use the volumetric flask to measure volume. Weigh the empty 20-mL volumetric flask and record the mass. Then, carefully fill the flask with the salt solution to just below the 20-mL fill mark. Do not fill it all the way. Use an eyedropper to finish filling the flask so that the bottom of the meniscus just touches the fill line on the neck of the flask. Then, weigh the full flask and record the weight.</div>
		</li>
		<li data-sub-note-item="1-10">
		<div class="lab-step">Lastly, use the 50-mL beaker to measure volume. Weigh the empty beaker, then fill the beaker with salt solution to the 20-mL line and weigh the full beaker. Record the masses in your notebook.</div>
		</li>
		<li data-sub-note-item="1-11">
		<div class="lab-step">Measure the displacement of the egg when it is submerged in the salt solution. First, remove the egg and look at the level of the salt solution in the 600-mL beaker. <strong>Note:</strong> If it is not at a marked line, add deionized water to the solution so that the meniscus is at a readable volume and record the volume.&#160;
		<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
			<tbody>
				<tr>
					<td><strong>Measurements</strong></td>
					<td><strong>Salt solution using 20-mL volumetric flask&#160;</strong></td>
					<td><strong>Egg</strong></td>
				</tr>
				<tr>
					<td>Mass (g)</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td>Displacement of water (cm)</td>
					<td style="text-align: center;">-</td>
					<td></td>
				</tr>
				<tr>
					<td>Inner diameter of 600-mL beaker (cm)</td>
					<td style="text-align: center;">-</td>
					<td></td>
				</tr>
				<tr>
					<td>Volume (cm&#179;)</td>
					<td style="text-align: center;">-</td>
					<td></td>
				</tr>
				<tr>
					<td>Volume (mL)</td>
					<td style="text-align: center;">20</td>
					<td></td>
				</tr>
				<tr>
					<td>Density (g/mL)</td>
					<td></td>
					<td></td>
				</tr>
			</tbody>
		</table>
		<strong><a href="https://jove.unicartagenaproxy.elogim.com/CDNSource/lm/labs/41/Table_2._Measuring_Density.xlsx" target="_blank">Click Here to download Table 2</a> </strong></div>
		</li>
		<li data-sub-note-item="1-12">
		<div class="lab-step">Gently place the egg in the beaker.</div>
		</li>
		<li data-sub-note-item="1-13">
		<div class="lab-step">Use a ruler to measure the new height of the solution level and record the height. Then, place the ruler across the top of the beaker to measure the inner diameter.</div>
		</li>
		<li data-sub-note-item="1-14">
		<div class="lab-step">Calculate the new volume from these measurements. Subtract the initial volume to obtain the displacement of the egg (volume).</div>
		</li>
		<li data-sub-note-item="1-15">
		<div class="lab-step">To clean up from the experiment, return the egg to your instructor and pour the salt solution down the sink. Rinse all used glassware with tap water and set them out to dry. Finally, return the sodium chloride to your instructor.</div>
		</li>
	</ul>
	</li>
	<li data-note-item="2" data-parent-collapse=""><strong>Results</strong><button class="expand">Expand</button>
	<ul class="lab-items">
		<li data-sub-note-item="2-1">
		<div class="lab-step">Calculate the mass of the solution in each of the different volumetric glassware by subtracting the empty glassware mass from the full glassware mass.</div>
		</li>
		<li data-sub-note-item="2-2">
		<div class="lab-step">Keep track of uncertainty in your measurement by using significant figures. For example, when we look at the graduated cylinder measurement, the weight of the empty container is 90.92, and the weight of the full container is 111.59. The weight measurements have significant figures to the hundredths place, so it is important that significant figures are maintained throughout our calculations. Thus, the mass of the 20-mL salt solution is 20.67 g.</div>
		</li>
		<li data-sub-note-item="2-3">
		<div class="lab-step">The 50-mL beaker weighed 28.65 g when it was empty and 47.96 g when it contained 20 mL of salt solution. Therefore, the mass of 20 mL of the salt solution is 19.31 g.</div>
		</li>
		<li data-sub-note-item="2-4">
		<div class="lab-step">Repeat the calculation to obtain the mass of 20 mL of salt solution that was measured using the 10-mL pipette and the 200-mL volumetric flask.<b> Note:</b> Generally speaking, volumetric measurements are more accurate when using a volumetric flask.</div>
		</li>
		<li data-sub-note-item="2-5">
		<div class="lab-step">Look at the accuracy of the other methods compared to the volumetric flask in terms of percent error. Percent error is calculated by subtracting the theoretical value from the experimental value, then dividing by the theoretical value times 100.
		<ul>
			<li><img src="https://jove.unicartagenaproxy.elogim.com/CDNSource/lm/labs/41/41_Procedure_1b.jpg" /></li>
		</ul>
		</div>
		</li>
		<li data-sub-note-item="2-6">
		<div class="lab-step">Use the volumetric flask measurement as the theoretical value to determine the percent error for the other volume measurement methods. As expected, the volume measurement that uses the markings on the beaker is the least accurate.<strong> Note: </strong>We also know this because the known density of pure water is 1 g/mL. Thus, 20 mL of pure water weighs 20 g. The density of the salt solution must be higher than 20 g since salt was added to the water.</div>
		</li>
		<li data-sub-note-item="2-7">
		<div class="lab-step">Calculate the density of the salt solution using the volumetric flask measurement. Divide the mass of the solution by the volume of the solution. Thus, the density of the salt solution is 1.053 g/mL.</div>
		</li>
		<li data-sub-note-item="2-8">
		<div class="lab-step">Compare the calculated density of the salt solution to the density of the egg. The volume calculated using the displacement of water in a beaker was found to be 61.14 cm<sup>3</sup> or 61.14 mL. The weight of the egg was measured to be 60.15 g, so its density is 0.9838 g/mL. The disparity between the density of the egg and the density of the salt solution arises from error introduced in both calculations, primarily due to volume measurement readings.</div>
		</li>
	</ul>
	</li>
</ol>

Accuracy, Precision, Uncertainty, and Significant Figures - Procedure

Techniki laboratoryjne

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

	Determining the Density of an EggExpand
	Density is a measure of how compact a substance or ...

Buffers

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

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

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

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

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

Extinction: Diversity and Extinction during the Paleogene Epoch - Procedure

Wymieranie

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

Biology

Ecology

Optimal Foraging

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

Optimal Foraging: Foraging with Artificial Habitats and Prey - Procedure

Optymalne żerowanie

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

Group Behavior

<ol class="note-items"><li data-note-item="1" data-parent-collapse="">The Hawk-Dove Game
<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 will play the hawk and dove game with a single partner and then with multiple partners to observe hypothetical costs and benefits of the antagonist or cooperative behavior respectively. 
</div></li><li data-sub-note-item="1-2"><div class="lab-step">Before the activity starts, review the cost-benefit table so that you understand how to score the game. <a href="https://jove.unicartagenaproxy.elogim.com/CDNSource/lm/labs/33/Table_1_Group_behavior.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">HYPOTHESES: The experimental hypothesis for this exercise could be that the dove strategy will become more frequent between repeated partners, and the hawk strategy will become more frequent when interactions are with a variety of partners. The null hypothesis could be that there will be no change in strategy over time with repeat or different partner combinations. NOTE: Benefit can be determined by looking at the left column and choosing the row that corresponds to the card you played. The box that corresponds to the choice of your partner can be found in the top row. For the purposes of this game, B equals four and C equals three. So depending on the cards played, player one here, which is you, would gain a score of two, four, zero or one half. 
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Collect a set of two game cards. 
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Take out the cards labeled hawk and dove and place them face down on the table or conceal them in your hands. 
</div></li><li data-sub-note-item="1-6"><div class="lab-step">When the instructor gives the signal show either of the two cards to the student sitting across from you. Try to strategize how to receive the highest benefit from every interaction on your own but do not communicate with your partner. 
</div></li><li data-sub-note-item="1-7"><div class="lab-step">In the Single Partner Recording Table (Table 2) provided by your instructor, note the cards played as well as the benefit you received for that round.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">Repeat showing cards to the same partner and determining benefit on your instructor&#39;s signal for a total of 15 rounds.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">At the end of those rounds, the student with the highest total benefit value was determined to be the &#8220;winner&#8221;, or the most fit.
</div></li><li data-sub-note-item="1-10"><div class="lab-step">As a class, determine the percentages of hawks and doves played as well as the average benefit received for each round, using this formula: Class average benefit=(sum total benefit)/(number of players)
</div></li><li data-sub-note-item="1-11"><div class="lab-step">For the second game, and then for each round of play, one row of students should rotate to their right, leaving all players with a new partner.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">Continue playing for 15 rounds or until students end up back in their starting positions, recording the data in the Multiple Partner Recording Table (Table 3) given to you by your instructor.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">For each round, determine the percentages of hawks and doves played that round, as well as the average benefit received.
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Cheating and Cooperation Exercise
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">NOTE: In this activity, you will exchange benefits, represented by colored or similar items, with your classmates. In this simulation, a cheater is defined as a person who gives fewer benefits than they receive. During this activity, you will observe the frequency of cheating behavior over time. 
</div></li><li data-sub-note-item="2-2"><div class="lab-step">HYPOTHESES: The experimental hypothesis is that cheaters will receive fewer benefits over time and cooperators will continue to provide equal benefits to one another. The null hypothesis is that the frequency of cheaters and cooperators does not change over time. 
</div></li><li data-sub-note-item="2-3"><div class="lab-step">Each person should collect beads of one color to represent a tangible benefit, as well as one paper bag and a three-ring binder.
</div></li><li data-sub-note-item="2-4"><div class="lab-step">Get into a group of three so that all members have objects of the same color.
</div></li><li data-sub-note-item="2-5"><div class="lab-step">Now find another group with a different color to pair with. In each round, partners will exchange benefits without knowing what they are receiving from their partner and without discussion within their group.  NOTE: Benefits of a different color from your starting color are worth twice as much.
</div></li><li data-sub-note-item="2-6"><div class="lab-step">Turn face to face with a member from the other group while still being able to conceal the objects in your possession.  TIP:<strong> </strong>Setting up a screen using a notebook or sitting behind a desk can be helpful here.
</div></li><li data-sub-note-item="2-7"><div class="lab-step">When given a signal from the instructor, place zero to two benefits into the paper bag and pass it to your partner.
</div></li><li data-sub-note-item="2-8"><div class="lab-step">Simultaneously, receive the bag from your partner and then remove the benefits from the bag.
</div></li><li data-sub-note-item="2-9"><div class="lab-step">Make a note of how many benefits you gave away and how many you received in the Cheating and Cooperation Table (Table 4) provided by your instructor.
</div></li><li data-sub-note-item="2-10"><div class="lab-step">Turn to a new member from the other group and repeat the exchange.
</div></li><li data-sub-note-item="2-11"><div class="lab-step">Rotate through all of the members of the other group for as many rounds as the instructor determines - which will not be revealed until the game ends so as not to alter player behavior.
</div></li><li data-sub-note-item="2-12"><div class="lab-step">Once the game has ended, note for each round in the cheating and cooperation table whether you gave more than your partner, received more from them or if you gave the same amount.
</div></li><li data-sub-note-item="2-13"><div class="lab-step">Now total up all of the benefits in your possession, with benefits you&#39;ve received from the other group being worth twice as much.
</div></li><li data-sub-note-item="2-14"><div class="lab-step">For each round, the whole class should total up the number of cheaters - with cheaters defined as those who gave less than they received.
</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">For both the single partner hawk-dove game and the multiple partners hawk-dove game, use the whole class data to plot the percentages as two separate lines - one for the percentage of hawks, and one for the percentage of doves, for each of the 15 rounds.
</div></li><li data-sub-note-item="3-2"><div class="lab-step">Take note of whether the cards played changed as more rounds were played.
</div></li><li data-sub-note-item="3-3"><div class="lab-step">Manipulate the values for B and C to see if you can find the point at which values of B and C would render the hawk strategy no longer effective (this would be the point when there is no benefit to choosing the hawk strategy over always choosing the dove strategy).
</div></li><li data-sub-note-item="3-4"><div class="lab-step">As a class, use the data from the cheating and cooperation activity to make a graph showing the percentage of rounds each individual player cheated versus the amount of benefits they ended with.
</div></li><li data-sub-note-item="3-5"><div class="lab-step">Consider whether it is better to be a cheater or a cooperator in this scenario and discuss as a class what makes this scenario different from the hawk-dove game.
</div></li><li data-sub-note-item="3-6"><div class="lab-step">Now, use the class data to graph the percentage of cheaters against the number of rounds, and look for any trends over time.
</div></li><li data-sub-note-item="3-7"><div class="lab-step">Consider which type of strategy would most likely evolve in a setting where partners interact repeatedly.
</div></li><li data-sub-note-item="3-8"><div class="lab-step">Then, note which tactic might be beneficial for a species that relies on cooperation. 
</div></li><li data-sub-note-item="3-9"><div class="lab-step">Discuss as a class what would likely happen if you increased or decreased the relative value of the benefits you receive.
</div></li></ul></li></ol>

Group Behavior: Hawk-Dove Game and Trading Exercise - Procedure

Zachowanie grupy

The Hawk-Dove Game
ExpandNOTE: In this activity, you will play the hawk and dove game with a single partner and then with multiple partners to observe ...

Genetics of Organisms

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

Genetics of Organisms: REF and WEF Fly Genetics - Procedure

Genetyka organizmów

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

Genetics

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://jove.unicartagenaproxy.elogim.com/CDNSource/lm/labs/15/Table_1_Animal_behavior.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 1</a>
</div></li><li data-sub-note-item="2-8"><div class="lab-step">Hold the chamber over the fly vial and carefully open the cap to return all of the <em>Drosophila</em> back to their empty vial. 
</div></li></ul></li><li data-note-item="3" data-parent-collapse="">Phototaxis
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="3-1"><div class="lab-step">To perform a phototaxis analysis, first cover side A of the choice chamber with tin foil, taking care that the foil reaches to the exact center between the two sides. 
</div></li><li data-sub-note-item="3-2"><div class="lab-step">Ensure the caps are secured on both ends of the chamber.  HYPOTHESES: In this exercise, the alternative hypothesis is that more flies will be found in the light side of the choice chamber, if the flues are attracted to light, or on the dark side if they are repelled by light. The null hypothesis is that <em>Drosophila</em> will have no preference with regard to either stimulus. Therefore, equal numbers of flies will be found on either side of the choice chamber.
</div></li><li data-sub-note-item="3-3"><div class="lab-step">Open the tape flap in the center of the choice chamber, insert the funnel into the hole, and tap all 40 flies into the chamber as previously described.
</div></li><li data-sub-note-item="3-4"><div class="lab-step">Set the timer to sound an alarm at 10 minutes.
</div></li><li data-sub-note-item="3-5"><div class="lab-step">When the alarm sounds, stop the timer, and count the number of flies in the open side B of the chamber.
</div></li><li data-sub-note-item="3-6"><div class="lab-step">Subtract that number from the total number of flies added to the choice chamber to calculate the number of flies in the covered side A.
</div></li><li data-sub-note-item="3-7"><div class="lab-step">Record these data in the table for time point 10.  <a href="https://jove.unicartagenaproxy.elogim.com/CDNSource/lm/labs/15/Table_2_Animal_behavior.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 2</a>
</div></li><li data-sub-note-item="3-8"><div class="lab-step">Carefully, return all of the flies back to their empty vial.
</div></li></ul></li><li data-note-item="4" data-parent-collapse="">Chemotaxis
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="4-1"><div class="lab-step">HYPOTHESES: In this exercise, the alternative hypothesis is that more flies will be found on the side of the chamber with the apple cider vinegar if the flies are attracted to the odor. Or, if the flies are repelled by the vinegar, then more of them will be found on the side of the chamber presenting the odorless water. The null hypothesis is that <em>Drosophila</em> will have no preference with regard to either stimulus and will move randomly in the chamber.  Use an individual disposable plastic pipette to dispense two drops of the apple cider vinegar onto one cotton ball for side A of the chamber. 
</div></li><li data-sub-note-item="4-2"><div class="lab-step">Use an individual disposable plastic pipette to dispense two drops of the distilled water onto a cotton ball for side B of the chamber. 
</div></li><li data-sub-note-item="4-3"><div class="lab-step">Fold two pieces of tape over themselves to create individual pieces of double-sided tape and use the tape to attach each cotton ball to the inside of the single bottle cap.
</div></li><li data-sub-note-item="4-4"><div class="lab-step">Screw the bottle caps with the cotton balls onto their corresponding sides of the chamber.
</div></li><li data-sub-note-item="4-5"><div class="lab-step">Transfer all of the flies into the chamber and set the timer to sound an alarm after 10 minutes. 
</div></li><li data-sub-note-item="4-6"><div class="lab-step">When the alarm sounds, stop the timer and record the number of flies in each side of the chamber. <a href="https://jove.unicartagenaproxy.elogim.com/CDNSource/lm/labs/15/Table_3_Animal_behavior.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 3</a>
</div></li><li data-sub-note-item="4-7"><div class="lab-step">At the end of the final experiment, funnel the flies into a morgue tube containing 70% ethanol. 
</div></li><li data-sub-note-item="4-8"><div class="lab-step">Dismantle the choice chambers for recycling.  NOTE: Since all of the chemicals used in these experiments are lab safe, the liquids can be disposed of down the sink, the chambers rinsed, and the solids can be thrown away in the trash. 
</div></li></ul></li><li data-note-item="5" data-parent-collapse="">Results
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="5-1"><div class="lab-step">Gather all of the data collected for the geotaxis, phototaxis, and chemotaxis experiments.
</div></li><li data-sub-note-item="5-2"><div class="lab-step">Plot these values as bar graphs for different conditions and different time points.
</div></li><li data-sub-note-item="5-3"><div class="lab-step">Use these data to draw inferences about the behavior of <em>drosophila melanogaster</em>. 
</div></li></ul></li></ol>

Animal Behavior: Studying Taxis Behavior of Drosophila - Procedure

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 ...

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://jove.unicartagenaproxy.elogim.com/CDNSource/lm/labs/25/Table_1_Measuring_biodiversity.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 1</a>
</div></li><li data-sub-note-item="1-6"><div class="lab-step">Once all of the species within a quadrat have been identified, pick up your hoop and move on to the next knot along the same transect.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">Continue surveying and moving to the next knot until you have completed the entire rope transect.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">Once you have completed the rope, move to a transect in a different habitat, and begin identifying the different species within each quadrat along the new transect, until each group has identified plants along three lengths of rope, one in each habitat, for a total of 15 quadrats surveyed.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">When all of the species data have been collected, carry the transects and quadrats back to the classroom.
</div></li></ul></li><li data-note-item="2" data-parent-collapse="">Species-Accumulation Curves and Richness/Evenness 
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">In the next activity, you will be calculating and comparing the species richness and evenness in two distinct populations, represented here by two opaque bags containing beads of various colors. For this exercise, start by taking one paper bag of beads per group.
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Have one person reach into the bag and randomly pull out three beads without looking.
</div></li><li data-sub-note-item="2-3"><div class="lab-step">Write down the bead colors, as well as the total number of colors pulled from the bag in the table. <a href="https://jove.unicartagenaproxy.elogim.com/CDNSource/lm/labs/25/Table_2_Measuring_biodiversity.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 2</a>HYPOTHESES: In this activity, the alternative hypothesis states that species evenness in the two communities will be different, while the richness will be the same. The null hypothesis might be that species richness and evenness will be the same in both communities.
</div></li><li data-sub-note-item="2-4"><div class="lab-step">Place the beads back in the bag after their colors are recorded.
</div></li><li data-sub-note-item="2-5"><div class="lab-step">Then, shake the bag and have the next person in the group draw another set of three beads.
</div></li><li data-sub-note-item="2-6"><div class="lab-step">Record the colors and numbers of beads that they pulled in the table.
</div></li><li data-sub-note-item="2-7"><div class="lab-step">Record the colors in the second row and the cumulative number of colors drawn in the last column.
</div></li><li data-sub-note-item="2-8"><div class="lab-step">Continue to take turns drawing three beads from the bag until 20 samples have been drawn.
</div></li><li data-sub-note-item="2-9"><div class="lab-step">Next, switch bags with a group with a bag from the other community, and repeat the exercise of drawing 20 samples, recording the color data in the appropriate community column of the table.
</div></li><li data-sub-note-item="2-10"><div class="lab-step">When all of the data have been recorded, return the beads to the bag and return the bags to your instructor.
</div></li></ul></li><li data-note-item="3" data-parent-collapse="">Results
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="3-1"><div class="lab-step">To calculate the alpha diversity of the transect and quadrat data collected by your group, use the data your data recorder entered for each species and enter it into your diversity table. <a href="https://jove.unicartagenaproxy.elogim.com/CDNSource/lm/labs/25/Table_3_Measuring_biodiversity.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 3</a>
</div></li><li data-sub-note-item="3-2"><div class="lab-step">Then, fill out the alpha diversity for each habitat into the appropriate column in the table.
</div></li><li data-sub-note-item="3-3"><div class="lab-step">Use the species data from your quadrat and transect table and enter the beta diversity data into the appropriate data cells within the diversity table. NOTE: The beta diversity is the number of species that are unique between two habitats. In this video, there are three species in common between the two sites, but a total of six unique species, giving a beta diversity of six between site A and B.
</div></li><li data-sub-note-item="3-4"><div class="lab-step">To calculate the gamma diversity for your activity, use the data from your quadrat and transect table and enter the gamma diversity data into the appropriate data cell. NOTE: The gamma diversity is the number of species in all of the habitats within a single study. It is not the sum of all of the alpha diversities, because species that are common between habitats should be counted only once.
</div></li><li data-sub-note-item="3-5"><div class="lab-step">To construct species accumulation curves for both of the bead sampling communities, use your bead color selection data to plot the species by sampling effort. That is the number of total colors by sample in the graph for each bead bag community.
</div></li><li data-sub-note-item="3-6"><div class="lab-step">Then, estimate the asymptote of the plotted data by appearance. Predict the number of species you would expect to find in each community. Note any differences in appearance between the two curves. If it was easier to estimate the number of species in one of the communities, make a hypothesis to explain this observation.
</div></li><li data-sub-note-item="3-7"><div class="lab-step">To construct rank abundance curves for both of the bead sampling communities, use the color bead data to fill in the table for both community A and community B, listing the colors that were identified during the sampling in the color column, and adding the total number of beads of each color that was sampled in the abundance column. <a href="https://jove.unicartagenaproxy.elogim.com/CDNSource/lm/labs/25/Table_4_Measuring_biodiversity.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 4</a>
</div></li><li data-sub-note-item="3-8"><div class="lab-step">Now, assign a rank to each color in the rank column, designating the color of the highest abundance the highest rank, rank one, and the color with the lowest abundance the lowest rank, rank 10. If there are multiple colors with the same number, just assign them consecutive ranks.
</div></li><li data-sub-note-item="3-9"><div class="lab-step">Plot individual rank abundance curves for the number of individuals by species ranked for the most to least common in both community A and community B. Compare the steepness of the two rank curves. Compare the richness and evenness between the two communities. Identify the community that you consider more biodiverse. 
</div></li></ul></li></ol>

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 ...

Scientific Method

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