<p class="jove_content">The current project was funded by the University of Kansas Proof of Concept Fund. We would like to thank the nursing staff at KUMC (Kansas City, KS) for assisting us in obtaining human umbilical cords, and David Jorgensen for assisting with cochleae cultures.</p>

<p class="jove_content">All procedures, including animal euthanasia, were conducted according to the approved Institutional Animal Care and Use Committee (IACUC) protocol (ACUP #2014-2234) at the University of Kansas Medical Center (KUMC).</p>
<p class="jove_content">NOTE: HWJCs were isolated from human umbilical cords that were donated by patients that provided informed consent and specimens were used in accordance with the protocols approved by the University of Kansas Human Subjects Committee (KU-IRB #15402).</p>
<p class="jove_title">1. Temporal Bone Harvest and Cochlea Isolation</p>
<ol>
	<li>Decapitate an appropriately euthanized, 15-week-old mouse by cutting between the skull and first cervical vertebra with a sharp pair of surgical scissors, then spray down the head with 70% ethanol to disinfect (<strong class="xfig">Figure 1A</strong>).</li>
	<li>Bisect the skull in a mid-sagittal plane using the same sharp pair of surgical scissors (<strong class="xfig">Figure 1B</strong><strong>-C</strong>, dashed lines).</li>
	<li>Remove the brain tissue from the skull using a pair of forceps (<strong class="xfig">Figure 1D</strong>, arrowhead).</li>
	<li>Identify the temporal bone through the presence of the auditory and vestibular nerve roots (<strong class="xfig">Figure 1E</strong><strong>-F</strong>, arrowheads) on the interior of the skull and the ear canal from the exterior (<strong class="xfig">Figure 1F</strong>, arrow). Carefully cut through the skull to isolate the temporal bone from the remainder of the skull (<strong class="xfig">Figure 1G</strong>).<br />
	NOTE: In some instances, it may be possible to delicately pry the surrounding bones of the skull away from the temporal bone without requiring cutting.</li>
	<li>Insert the fine forceps into the opening of the ear canal approximately 5 mm (<strong class="xfig">Figure 1H</strong>), so the tips do not risk puncturing the cochlea. Gently break open the bone of the bulla (<strong class="xfig">Figure 1H</strong>, top red shaded area) so it fractures (<strong class="xfig">Figure 1I</strong>, red dotted line).<br />
	NOTE: If necessary, a dissection microscope may aid in this process to ensure accurate placement and manipulation of instruments.</li>
	<li>Using fine forceps, pry away the remainder of the bulla from the temporal bone, exposing the bony labyrinth of the cochlea (<strong class="xfig">Figure 1J</strong>, red brackets).</li>
	<li>Place a Petri dish large enough to hold the temporal bone (<em>e.g.</em>, 30 mm or greater) underneath the dissecting scope field of view and fill it with enough Phosphate Buffered Saline (PBS) to cover the temporal bone (<em>e.g.</em>, 0.5 to 1 mL).</li>
	<li>Submerge the temporal bone with the bulla removed in the PBS with the oval and round windows of the cochlea facing up.<br />
	NOTE: Removal of the vestibular system from the cochlea is not necessary, as the vestibular system serves well as a handle for manipulating and orienting features of the cochlea (<strong class="xfig">Figure 1I</strong> and <strong class="xfig">Figure 2B</strong>).</li>
	<li>Using an ultra-fine pair of forceps, remove the stapedial artery, which passes through the stapes.</li>
	<li>Insert one tip of the ultra-fine forceps through the arch of the stapes where the artery passed through, and delicately lift the stapes upward. The disarticulated stapes should lift off of the oval window.</li>
	<li>Using one tip of the ultra-fine forceps, puncture the oval and round windows.</li>
	<li>Using a 1x PBS filled 28.5-gauge syringe with tubing (inner diameter 0.28 mm, outer diameter 0.61 mm) attached, position the tubing over the oval window (<strong class="xfig">Figure 2A</strong><strong> - B</strong>).<br />
	NOTE: The opening of the tube can be manipulated using ultra-fine forceps helping to ensure accurate placement.
	<ol>
		<li>Use a fresh syringe and tubing for each cochlea.</li>
	</ol>
	</li>
	<li>Perfuse 2 mL of 10% antibiotic-antimycotic (anti-anti) and 10% penicillin-streptomycin (pen-strep) in PBS through the cochlea over 5 min to remove the perilymph. Perfusing too fast with too much pressure will damage the fine structures within the cochlea.<br />
	NOTE: Manually driving the syringe by hand to perfuse the fluid through the cochlea is effective, although a perfusion pump could be used as an alternative.
	<ol>
		<li>If manually perfusing, use a pair of fine, self-closing forceps to stabilize the cochlea (<strong class="xfig">Figure 2A</strong>) during perfusion by grasping the vestibular portion of the temporal bone.<br />
		NOTE: Although not required, this leaves both hands free to handle instruments; one hand can drive the syringe (<strong class="xfig">Figure 2A</strong>) while the other can position the tubing appropriately (<strong class="xfig">Figure 2B</strong>).</li>
		<li>From this step forward, take care to avoid exposing the cochlea unnecessarily to air. Introducing air bubbles into the fluid spaces will block fluid circulation, and further drying out will damage the fine structures within the cochlea.</li>
	</ol>
	</li>
	<li>Remove and discard any remaining muscle tissue and bone fragments using fine forceps.</li>
	<li>If necessary, store isolated cochleae at 4 &#176;C in 10% anti-anti and 10% pen-strep solution in PBS for up to seven days with regular changes of antibiotic solution every 48 h.</li>
</ol>
<p class="jove_title">2. Cochlear Processing</p>
<ol>
	<li><strong>Decellularization</strong>
	<ol>
		<li>For each cochlea, fill a 20-mL glass scintillation vial with a 1% sodium dodecyl sulfate (SDS) solution in de-ionized (DI) water, which is used to decellularize the cochlea.</li>
		<li>Cut the tip off of a plastic 7-mL transfer pipette using a razor blade so that the opening is large enough for a cochlea to pass through.</li>
		<li>Using the transfer pipette, gently draw the cochlea up into the pipette so it just passes the cut-off edge by a few millimeters.</li>
		<li>Expel the cochlea into the 1% SDS solution-filled scintillation vial.</li>
		<li>Circulate 2 mL of 1% SDS in de-ionized (DI) water through the cochlea in the scintillation vial&#160;using a transfer pipette.</li>
		<li>Place the scintillation vial into a rotator, and allow the scintillation vial to rotate at 10 rpm for 72 h at room temperature.
		<ol>
			<li>Change 1% SDS solution every 24 h over a 72 h period. Take care when changing SDS to never expose the cochlea to air.</li>
		</ol>
		</li>
		<li>Wash the cochlea three times for 30 min each with DI water. Perform the washes in the same scintillation vial used for the SDS charges; at this stage, the cochlea is decellularized.</li>
	</ol>
	</li>
	<li><strong>Decalcification</strong>
	<ol>
		<li>Remove the remaining DI water from the scintillation vial, leaving just enough to keep the cochlea submerged.</li>
		<li>To begin decalcification, add 5 mL of 10% ethylenediaminetetraacetic acid (EDTA) (0.02 M) in DI water to the scintillation vial containing the cochlea.</li>
		<li>Circulate 2 mL of 10% EDTA in DI water through the cochlea in the scintillation vial&#160;using a transfer pipette.</li>
		<li>Place the scintillation vial back in the rotator, and allow the scintillation vial to rotate at 10 rpm for 72 h at room temperature. Change the 10% EDTA solution every 24 h over a 72 h period. Take care when changing solutions so that the cochlea is not exposed to air.</li>
		<li>Rinse the cochlea 3 times for 2 h each with 1x PBS; at this stage, the cochlea is decalcified.</li>
	</ol>
	</li>
	<li><strong>Storage</strong>
	<ol>
		<li>Store the cochleae for up to 72 h in a 10% anti-anti, 10% pen-strep solution in PBS at 4 &#176;C.<br />
		NOTE: Longer storage periods may be possible with regular changes of antibiotic solution; however, extended storage of processed cochleae has not been validated in this protocol.</li>
	</ol>
	</li>
</ol>
<p class="jove_title">3. Procurement and Expansion of hWJCs</p>
<ol>
	<li>Isolate hWJCs according to previously published protocols<sup class="xref">22</sup>. Briefly, segment umbilical cords into 3-cm sections.</li>
	<li>Carefully make a shallow incision with a scalpel lengthwise on each umbilical cord segment to unroll and expose the Wharton&#39;s jelly of the umbilical cord. Use forceps to remove the blood vessels.</li>
	<li>Wash umbilical cord segments twice with enough PBS (<em>e.g.</em>, 40 mL in a 60 mm Petri dish) to submerge the tissue. Finely mince tissue segments with a scalpel. Digest tissue in a 100 mm Petri dish with 50 mL of digestion medium (0.2% type 2 collagenase, 1% penicillin-streptomycin, in low-glucose Dulbecco&#39;s Modified Eagle&#39;s Medium (DMEM)).</li>
	<li>Place tissue in digesting medium on an orbital shaker at 50 rpm overnight in a 5% CO<sub>2</sub>, 37 &#176;C incubator. The following day, dilute digestion medium at a ratio of 1:16 in 2% antibiotic-antimycotic in PBS, and pellet cells by centrifugation at 500 x g at room temperature (~ 27 &#176;C) for 10 min. Discard supernatant.</li>
	<li>Resuspend pellets in Mesenchymal Stem Cell Growth Medium (MSCGM) and recombine cells. Plate cells at a density of 7 x 10<sup>3</sup> cells/cm<sup>2 </sup>in tissue culture treated T-75 flasks. Culture HWJCs in MSCGM, and expand to passage 5 for experiments.<br />
	NOTE: HWJCs may be sub-cultured into larger T-flasks (<em>e.g.</em>, T-150, T-300) when passaging to increase yield of hWJCs for experiments.</li>
</ol>
<p class="jove_title">4. Infusion of hWJCs into Decellularized Cochleae</p>
<ol>
	<li><strong>Prepare decellularized cochleae</strong>
	<ol>
		<li>Using aseptic technique, wash stored cochleae in 1x PBS three times for 30 min each.</li>
		<li>Transfer cochleae into separate wells of a 24-well plate, and add 1 mL of 37 &#176;C pre-warmed MSCGM.</li>
		<li>Incubate cochleae in a 5% CO<sub>2</sub>, 37 &#176;C cell culture incubator for a minimum of 1 h before infusion of cells.</li>
	</ol>
	</li>
	<li><strong>Dissociate hWJCs and resuspend</strong>
	<ol>
		<li>Wash hWJCs with 37 &#176;C pre-warmed 1x PBS twice.</li>
		<li>Add enough 37 &#176;C pre-warmed Trypsin with 0.5% EDTA to cover the cell vessel surface.</li>
		<li>Incubate hWJCs for up to 5 min in a 5% CO<sub>2</sub>, 37 &#176;C cell culture incubator.
		<ol>
			<li>Verify that 90% of cells have detached from the cell culture surface by viewing cells under an inverted microscope.<br />
			NOTE: Cells that move when the culture vessel is gently tapped, have detached. Cells that remain stationary, have not detached.</li>
			<li>Gently tap the sides of the flask to further dislodge attached cells.</li>
		</ol>
		</li>
		<li>Using aseptic technique, transfer hWJCs from the culture vessel to a 50-mL conical tube containing an equivalent volume of MSCGM to the volume of Trypsin used to dissociate the cells.</li>
		<li>Pellet the hWJCs by centrifuging at 500 x g at room temperature (~ 27 &#176;C) for 5 min.</li>
		<li>Aspirate supernatant, and resuspend hWJCs in MSCGM at a concentration of 500,000 cells/mL.</li>
	</ol>
	</li>
	<li><strong>Perfuse cochleae</strong>
	<ol>
		<li>Transfer cochleae to a new 24-well plate using a transfer pipette.</li>
		<li>Add a drop of MSCGM to each cochlea to prevent any cochlea from drying out.</li>
		<li>Delicately orient the cochlea using ultra-fine forceps so that the oval and round windows are facing up.<br />
		NOTE: Take extreme care when directly handling each cochlea as the cochlea is easily damaged.</li>
		<li>Using a sterile 28.5-gauge insulin syringe with connected tubing, draw up 0.2 mL resuspended hWJCs (100,000 cells).</li>
		<li>Using sterilized fine forceps, position tubing over the oval window.</li>
		<li>Delicately and slowly perfuse the cochlea with 0.2 mL of resuspended hWJCs using a 28.5-gauge insulin syringe connected to polyethylene tubing (Outer Diameter: 0.61 mm, Inner Diameter: 0.28 mm) over approximately 5 min.</li>
		<li>After perfusion, add 0.8 mL of 37 &#176;C pre-warmed MSCGM to the well containing the cochlea to bring the total volume in the well to 1 mL.</li>
		<li>Repeat steps 4.3.3-4.3.7 for each cochlea, using a fresh syringe and tubing each time.</li>
		<li>Place perfused cochleae in a 5% CO<sub>2</sub>, 37 &#176;C cell culture incubator, and change media three times per week.</li>
	</ol>
	</li>
</ol>
<p class="jove_title">5. Cochlea Harvest and Preservation</p>
<p class="jove_content">NOTE: Cochleae may be cultured and harvested at any time point up to 30 days post-perfusion.</p>
<ol>
	<li>To preserve cochleae, fix in 4% paraformaldehyde in 1x PBS overnight at 4 &#176;C on a rocking platform.</li>
	<li>After fixation, wash cochleae with 1x PBS three times for 5 min each.</li>
	<li>Gradually dehydrate cochlea with ethanol and clear with an aliphatic hydrocarbon solvent before embedding in paraffin.</li>
	<li>Section samples to a thickness of 10 &#181;m using a microtome and mount on glass microscope slides.<br />
	NOTE: Samples are now ready for histological or immunohistochemical processing using standard protocols.</li>
</ol>

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J., Converse, G. L., Hopkins, R. A., Detamore, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+bioactivity+of+cartilage+extracellular+matrix+in+articular+cartilage+regeneration.">The bioactivity of cartilage extracellular matrix in articular cartilage regeneration.</a> <em style='font-style: italic'>Adv Healthc Mater</em>. <b>4</b> (1), 29-39 (2015).</li></ol>

<p class="jove_content">The authors have nothing to disclose.</p>

<p class="jove_content">We have successfully demonstrated that native cochlear cells can be removed from the cochlea via a decellularization process, which allows for the use of the cochlea as an intricate, three-dimensional tissue scaffold. Santi <em>et al</em>.<sup class="xref">15</sup> developed the initial method for decellularizing cochleae, and have accurately estimated the volumes of many cochlear structures through with the aid of light sheet microscopy<sup class="xref">23</sup>. Such early work served as a strong basis for the tissue engineering and cell culture techniques presented here. Decellularized cochlea can successfully be infused with cells, and then cultured for a prolonged period of time. Cochleae from a wide array of animals could theoretically be utilized in combination with a similarly wide array of cells types. These combinations allow the techniques presented here to be utilized in numerous possible experimental designs, such as tissue engineering applications that examine sensory epithelium development<sup class="xref">14</sup>, drug testing of new pharmaceutical agents<sup class="xref">24</sup>, and development of cochlear repair models<sup class="xref">25</sup>. However, despite the broad applicability of these techniques they are not without limitations.</p>
<p class="jove_content">One of the limitations of the current technique is that the perfusion is variable based on individual. Developing a holder, for the cochlea, and using a syringe pump to conduct perfusions may increase the accuracy and success of the process. Additionally, definitively proving successful decellularization could potentially be accomplished through several methods. We employed DAPI staining that was used to quantify remaining cell nuclei, and we observed no residual nuclei post-decellularization. Additionally, standard DNA quantification assays utilize the reagent, green dye detecting picogram quantities of DNA, to identify nucleic acid remnants, which may visually be more sensitive than DAPI for identifying the presence of small fragments of nucleic acids in decellularized tissues<sup class="xref">26</sup>. Regardless of which techniques are used to validate successful decellularization, it may not be possible to show complete decellularization of an individual cochlea prior to utilizing it as a scaffold. Nevertheless, we have found the decellularization process to be successful and robust.</p>
<p class="jove_content">The two most critical steps in the current protocol are the isolation and handling of the cochleae, and the perfusion of the cochleae. Great care must be taken when the cochlea is initially isolated, as the cochlea is delicate and brittle. If too much force is applied while handling the cochlea with forceps, the cochlea could fragment. Thus, it is imperative to handle each cochlea gently before decellularization and decalcification. The vestibular system, if left intact, serves as a great handle for grabbing the inner ear complex with forceps, and orienting the cochlea. Similarly, when perfusing the cochlea with cells, the pressure behind the perfusion must not be too great, otherwise cells may not survive the perfusion. As noted in the current protocol, attaching tubing to the end of a 1-mL insulin syringe allows for easier handling of the cochlea with one hand, and greater precision in perfusing cells through the cochlea with the other hand.</p>
<p class="jove_content">As a next step, it would be natural to assess developmental and functional markers to determine if the cochlear ECM is inducing differentiation of stem cells, and if so, which specific ECM components in the cochlea are inducing changes in cells. Perfusing different types of cells (<em>e.g.</em>, stem cells, neuroprogenitors, fibroblasts, <em>etc.</em>) through the cochlea to monitor cell differentiation and behavior would be another fascinating follow-up investigation. In addition, cells that have been genetically modified or are exposed to a growth factor differentiation protocol may provide new insights into how the cochlear ECM supports neurosensory development, and possibly even function in hearing. Thus, the current protocol is a significant first step in using the cochlear ECM to study inner ear neurosensory development and maintenance, which may one day lead to the development of new therapies to restore hearing loss.</p>

<p class="jove_content">The cochlea is an intricate spiral structure found in the temporal bone. It is composed of an outer bony labyrinth and a concentric, inner membranous labyrinth<sup class="xref">1</sup>. The membranous labyrinth consists of three fluid spaces: Scala vestibuli, Scala media, and Scala tympani<sup class="xref">1</sup>. The scala media houses the sensory epithelium, which is composed of a multitude of cell types, but the sensory hair cells (HC), which transduce mechanical energy in sound waves to nerve impulses<sup class="xref">2</sup>, are of particular interest. Exposure to acoustic trauma<sup class="xref">3</sup><sup>,</sup><sup class="xref">4</sup><sup>,</sup><sup class="xref">5</sup>, medication<sup class="xref">6</sup>, disease<sup class="xref">7</sup><sup>,</sup><sup class="xref">8</sup>, and aging<sup class="xref">9</sup> can all result in impaired auditory function via HC death. Hair cell loss in mammals is permanent, unlike avian HCs, which can regenerate after injury<sup class="xref">10</sup>.</p>
<p class="jove_content">A variety of contemporary research efforts have sought to restore lost HCs, although the specific experimental approaches vary. Manipulation of gene expression in the sensory epithelium and implantation of stem cells differentiated outside the body are dominant approaches in the field, although induction methods that seek to differentiate stem cells into cochlear organoids have been attempted<sup class="xref">11</sup><sup>,</sup><sup class="xref">12</sup><sup>,</sup><sup class="xref">13</sup>. Each of these approaches is either reliant directly on stem cells, or the developmental cues used by stem cells; however, a second shared, and potentially critical, element is the ECM of the cochlea itself<sup class="xref">14</sup><sup>,</sup><sup class="xref">15</sup>.</p>
<p class="jove_content">The ECM not only provides physical support for cells and tissue, which includes a surface for cell adhesion, proliferation, survival, and migration, but also plays critical roles in the development of HCs and the spiral ganglion<sup class="xref">15</sup><sup>,</sup><sup class="xref">16</sup><sup>,</sup><sup class="xref">17</sup>. Naturally occurring ECM provides inductive signals that may guide cell phenotype determination and/or cell adhesion, proliferation, and survival<sup class="xref">18</sup>. Consequently, the use of decellularized cochlea in combination with cultured hWJCs offer a unique opportunity to explore the role of the ECM and HC regeneration. HWJCs are a readily available, non-controversial cell type isolated from human umbilical cords that behave like mesenchymal stem cells<sup class="xref">19</sup>. HWJCs have shown the ability to differentiate down neurosensory cell lineages<sup class="xref">20</sup><sup>,</sup><sup class="xref">21</sup>. Thus, the current protocol details the isolation, decellularization, and perfusion of cochleae from C57BL mouse carcasses with hWJCs for inner ear tissue engineering.</p>

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

a protocol for decellularizing mouse cochleae for inner ear tissue engineering

<p class="jove_content">In mammals, mechanosensory hair cells that facilitate hearing lack the ability to regenerate, which has limited treatments for hearing loss. Current regenerative medicine strategies have focused on transplanting stem cells or genetic manipulation of surrounding support cells in the inner ear to encourage replacement of damaged stem cells to correct hearing loss. Yet, the extracellular matrix (ECM) may play a vital role in inducing and maintaining function of hair cells, and has not been well investigated. Using the cochlear ECM as a scaffold to grow adult stem cells may provide unique insights into how the composition and architecture of the extracellular environment aids cells in sustaining hearing function. Here we present a method for isolating and decellularizing cochleae from mice to use as scaffolds accepting perfused adult stem cells. In the current protocol, cochleae are isolated from euthanized mice, decellularized, and decalcified. Afterward, human Wharton&#39;s jelly cells (hWJCs) that were isolated from the umbilical cord were carefully perfused into each cochlea. The cochleae were used as bioreactors, and cells were cultured for 30 days before undergoing processing for analysis. Decellularized cochleae retained identifiable extracellular structures, but did not reveal the presence of cells or noticeable fragments of DNA. Cells perfused into the cochlea invaded most of the interior and exterior of the cochlea and grew without incident over a duration of 30 days. Thus, the current method can be used to study how cochlear ECM affects cell development and behavior.</p>

<p class="jove_content" fo:keep-together.within-page="1">Using the methods presented here, successful decellularization of cochleae was assessed by examining the presence or absence of DNA through 4&#39;,6-diamidino-2-phenylindole (DAPI) staining. Cochleae were considered fully decellularized if DNA was not identified within the decellularized cochlea. A native cochlea from a previous experiment that did not undergo decellularization or decalcification was used as a positive control to illustrate the structures and cells traditionally found in a C57BL mouse cochlea. The decellularized-decalcified cochlea, which did not have any hWJCs infused, was used as a negative control. No quantifiable DAPI stained cell nuclei were observed in any region of the tissue section (<strong class="xfig">Figure 3</strong>). Two cochleae were decellularized and decalcified, and then infused with hWJCs for the current project. The decellularized-decalcified cochleae that were perfused with hWJCs were observed to have numerous DAPI stained nuclei within the cochlea and growing on the bony shell outside of the cochlea (<strong>Table 1</strong>). Total estimated cell densities for the cell infused first cochlea were 113,759 cells/cochlea after 30 days of culture (<strong class="xfig">Figure 4</strong> and <strong class="xfig">Figure 5</strong>) and for the second cochlea were 118,732 cells/cochlea after 30 days of culture (<strong class="xfig">Figure 6</strong> and <strong class="xfig">Figure 7</strong>). These cell density estimates include the Scala tympani, Scala media, and Scala vestibule, and volumes of the structures found within the three fluid spaces, but these estimates excluded any of the remaining bony labyrinth. In addition, samples from each cochlea were stained with Hematoxylin and Eosin (H&#38;E) to show the presence of cells and anatomical features within each cochlea (<strong class="xfig">Figure 8</strong>). The gross anatomy of the cochlear microstructures remained predominantly unchanged throughout all stained sections; however, no cells were identifiable in the decellularized-decalcified section (<strong class="xfig">Figure 8B</strong>). The identification of cell nuclei was dramatically different between the native cochlea (<strong class="xfig">Figure 8A</strong>) and the decellularized-decalcified cochleae infused with cells (<strong class="xfig">Figure 8C</strong><strong>-</strong><strong>D</strong>).</p>
<p class="jove_content" fo:keep-together.within-page="1"><img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56523/56523fig1.jpg" /><br />
<strong>Figure 1: Mouse temporal bone isolation.</strong> Once the mouse has been appropriately euthanized, decapitate the animal by cutting between the base of the skull and the first cervical vertebra, and spray the head with ethanol to disinfect (<strong>A</strong>). Bisect the skull in a sagittal plane of section (<strong>B</strong><strong>-</strong><strong>C</strong>) using a sharp pair of surgical scissors, cutting through both the brain tissue and bone. The bisected mouse head (<strong>D</strong>, lower mandible removed for unrelated experimental work) must then have the brain tissue removed to reveal the temporal bone. The two foramina through which the auditory and vestibular nerves pass (<strong>E</strong>, red arrowheads) are useful cues for successfully identifying the temporal bone. Remove the flesh from the skull (<strong>F</strong>). The external ear canal (<strong>F</strong>, arrow, top panel) provides a useful external cue as well. Carefully trim away the excess bone, leaving just the isolated temporal bone (<strong>G</strong>). The bulla (<strong>H</strong>, top panel, red highlighted area) must then be delicately removed using fine forceps (<strong>H</strong>, lower panel). The bulla is thin bone and can be readily chipped away (<strong>I</strong>) until the bony labyrinth of the cochlea is revealed (<strong>J</strong>, red brackets). Scale bars on all images are 1 mm. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56523/56523fig1large.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" class="xfigimg" src="/files/ftp_upload/56523/56523fig2.jpg" /><br />
<strong>Figure 2: Cochlear Perfusion.</strong> Once the temporal bone has been isolated and the bulla has been removed, the cochlea can then be set up for perfusion at a microscope. When perfusing using a manually driven syringe with tubing attached (<strong>A</strong>, dotted line) it may be helpful to stabilize the cochlea using a pair of self-closing forceps. Gently attach the forceps to the vestibular part of the temporal bone (<strong>B</strong>) and rest the forceps against the lip of the Petri dish (<strong>A</strong>). This leaves both hands free to manipulate instruments during perfusion. One hand can then manipulate the syringe, controlling the flow rate of the fluid, while the other hand can position the tubing over the oval window using a second pair of forceps (<strong>B</strong>). Cutting the free end of the tubing at an angle allows for easier positioning, allowing the tubing to be placed correctly without blocking the field of view. Scale bar is 1 mm. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56523/56523fig2large.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 3" class="xfigimg" src="/files/ftp_upload/56523/56523fig3.jpg" /><br />
<strong>Figure 3: Tissue section of a decellularized mouse cochlea.</strong> An upright epi-fluorescent microscope was used to image samples at a magnification of 200X (10X eyepiece and 20X objective). Images were stitched together to create approximately a 4 x 5 montage. DAPI nuclear staining showing absence of cells is shown in panel <strong>A </strong>using a DAPI common filter set (350 nm excitation, 470 nm emission), and an overexposed, grayscale image of the autofluorescence using a fluorescein isothiocyanate (FITC) common filter set (490 nm excitation, 525 nm emission), which provides anatomical landmarks, is shown in panel <strong>B</strong>. Automated cell counts were performed in three regions of interest, which are delineated on both panels <strong>A</strong> and <strong>B</strong>. Total cell counts were calculated for the entire tissue section (<strong>1</strong>, solid white line) and the cochlea (<strong>2</strong>, dashed line). These two numbers were used to calculate the number of cells in the bony structures outside of the cochlea but within the exterior, bony edges of the tissue section (<strong>3</strong>, space between the dotted line and dashed line). Scale bars on all images are 500 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56523/56523fig3large.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 4" class="xfigimg" src="/files/ftp_upload/56523/56523fig4.jpg" /><br />
<strong>Figure 4: A near-modiolar section of decellularized mouse cochlea 1 infused with hWJCs.</strong> DAPI nuclear staining is shown in panel <strong>A</strong> overlaid onto an overexposed, grayscale autofluorescence from the green channel, which provides anatomical landmarks. Automated cell counts were performed in three regions of interest, which are delineated in panel <strong>A</strong>. Total cell counts were calculated for the entire tissue section (<strong>1</strong>, solid white line) and the cochlea (<strong>2</strong>, dashed line). These two numbers were used to calculate the number of cells in the bony structures outside of the cochlea but within the exterior, bony edges of the tissue section (<strong>3</strong>, space between the dotted line and dashed line). Isolated DAPI staining is shown in panel <strong>B</strong>, and the threshold of the DAPI staining used during automated quantification is shown in panel <strong>C</strong>. Scale bars on all images are 500 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56523/56523fig4large.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 5" class="xfigimg" src="/files/ftp_upload/56523/56523fig5.jpg" /><br />
<strong>Figure 5: High magnification view of a near-modiolar section of decellularized mouse cochlea 1 infused with hWJCs.</strong> DAPI nuclear staining is shown in panel <strong>A</strong> overlaid onto an overexposed, grayscale image of the autofluorescence from the green channel, which provides anatomical landmarks. Isolated DAPI staining is shown in panel <strong>B</strong>, and the threshold image used during automated cell counting is shown in panel <strong>C</strong>. The fluid spaces and finer microstructures housed within the cochlea are nicely preserved during the decellularization and cell culture phases of the experiment. Scala Vestibuli (SV), Scala Tympani (SM), Basilar Membrane (BM), Tectorial Membrane (TM), Spiral Limbus (L), Rosenthal&#39;s Canal (RC), Modiolus (M), Spiral Ligament (SL), and Stria Vascularis (<strong>*</strong>). Reissner&#39;s membrane was not clearly defined in tissue sections, and as a result the Scala Media was not clearly defined. Scale bars on all images are 250 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56523/56523fig5large.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 6" class="xfigimg" src="/files/ftp_upload/56523/56523fig6.jpg" /><br />
<strong>Figure 6: A near-modiolar section of decellularized mouse cochlea 2 that was infused with hWJCs.</strong> DAPI nuclear staining is shown in panel <strong>A</strong>, overlaid onto grayscale autofluorescence from the green channel, which provides anatomical landmarks. Automated cell counts were performed in three regions of interest, which are delineated in panel <strong>A</strong>. Total cell counts were calculated for the entire tissue section (<strong>1</strong>, solid white line) and the cochlea (<strong>2</strong>, dashed line). These two numbers were used to calculate the number of cells in the bony structures outside of the cochlea but within the exterior, bony edges of the tissue section (<strong>3</strong>, space between the dotted line and dashed lines). Isolated DAPI staining is shown in panel <strong>B</strong>, and the threshold of the DAPI staining used during automated quantification is shown in panel <strong>C</strong>. Scale bars on all images are 500 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56523/56523fig6large.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 7" class="xfigimg" src="/files/ftp_upload/56523/56523fig7.jpg" /><br />
<strong>Figure 7: High magnification view of a near-modiolar section of decellularized mouse cochlea 2 that was infused with hWJCs.</strong> DAPI nuclear staining is shown in panel <strong>A</strong>, overlaid onto grayscale autofluorescence from the green channel, which provides anatomical landmarks. Isolated DAPI staining is shown in panel <strong>B</strong>, and the threshold image used during automated cell counting is shown in panel <strong>C</strong>. The fluid spaces and finer microstructures housed within the cochlear are preserved during the decellularization and cell culture phases of the experiment. Scala Vestibuli (SV), Scala Tympani (SM), Basilar Membrane (BM), Tectorial Membrane (TM), Spiral Limbus (L), Rosenthal&#39;s Canal (RC), Modiolus (M), Spiral Ligament (SL), and Stria Vascularis (<strong>*</strong>). Reissner&#39;s Membrane was not clearly defined in tissue sections, and as a result the Scala Media was not clearly defined. Scale bars on all images are 250 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56523/56523fig7large.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 8" class="xfigimg" src="/files/ftp_upload/56523/56523fig8.jpg" /><br />
<strong>Figure 8: Hematoxylin and Eosin staining of control and treated cochleae.</strong> Hematoxylin and Eosin stained sections displaying a typical cochlea with native cells present is shown in panel <strong>A</strong>. Panel <strong>B</strong> contains the same structures as panel <strong>A</strong>, but is from a fully decellularized cochlea, which leaves behind the extracellular matrix. The two previously shown cochleae infused with hWJCs are seen in panels <strong>C</strong> and <strong>D</strong>. The gross cochlear anatomy remains predominantly unchanged through the decellularization and cell culture processes, although the specific cell populations and locations are dramatically altered. Scala Vestibuli (SV), Scala Tympani (SM), Basilar Membrane (BM), Tectorial Membrane (TM), Spiral Limbus (L), Rosenthal&#39;s Canal (RC), Modiolus (M), Spiral Ligament (SL), and Stria Vascularis (*). Scale bars on all images are 250 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56523/56523fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a></p>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td><strong>A1 Cochlea + Cells</strong></td>
			<td><strong>A2 Cochlea + Cells</strong></td>
			<td><strong>B2 Neg. Control Cochlea</strong></td>
		</tr>
		<tr>
			<td><strong>Outside Bony Labyrinth</strong></td>
			<td>3,758</td>
			<td>549</td>
			<td>0</td>
		</tr>
		<tr>
			<td><strong>Bone Outside Cochlea Spaces</strong></td>
			<td>248</td>
			<td>586</td>
			<td>0</td>
		</tr>
		<tr>
			<td><strong>Within Cochlea</strong></td>
			<td>518</td>
			<td>701</td>
			<td>0</td>
		</tr>
		<tr>
			<td><strong>Total Cells</strong></td>
			<td>4,524</td>
			<td>1,836</td>
			<td>0</td>
		</tr>
		<tr>
			<td><strong>Section Volume (&#956;l)</strong></td>
			<td>0.0077</td>
			<td>0.0100</td>
			<td>0.0147</td>
		</tr>
		<tr>
			<td><strong>Percent of Cochlea Volume</strong></td>
			<td>0.46%</td>
			<td>0.59%</td>
			<td>0.87%</td>
		</tr>
		<tr>
			<td><strong>Estimated Cell Density within Cochlea</strong></td>
			<td>113,759</td>
			<td>118,732</td>
			<td>0</td>
		</tr>
	</tbody>
</table>
<p class="jove_content" fo:keep-together.within-page="1"><strong>Table 1: Cell Counts. </strong>DAPI stained cochlear sections were thresholded and quantified using automatic counting tools in ImageJ in 3 non-overlapping regions of interest (outside the bony labyrinth, temporal bone outside of the cochlear fluid spaces, and within the cochlea). Cross-sectional cochlear area was also measured in ImageJ, and this value was used to calculate section volume in combination with section thickness (10 &#181;m). Using an estimate of total cochlear volume published by Santi <em>et al.</em><sup class="xref">23</sup>, the relative cochlear volume of a given section was calculated. The cell density of the entire cochlea was estimated using the relative cochlear volume of a given section in combination with the paired cell count.</p>

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

A Protocol for Decellularizing Mouse Cochleae for Inner Ear Tissue Engineering

<p class="jove_content">The goal of this protocol is to demonstrate an effective method to decellularize and decalcify mouse cochleae for utilization as scaffolds for tissue engineering applications.</p>

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

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10.7K Views.  University of Kansas Medical Center. The goal of this protocol is to demonstrate an effective method to decellularize and decalcify mouse cochleae for utilization as scaffolds for tissue engineering applications.

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

Evolutionary Relationships

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

Evolutionary Relationships: Using BLAST to Test Evolutionary Hypotheses - Concept

Związki ewolucyjne

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

Education

JoVE Lab Manual

Biology

Evolution

Extraction

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

Wydobywanie

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

	Cellulose RecoveryExpand
	Extraction and filtration can ...

Chemistry

Lab Manual Sub Articles

Electrolytic Cells

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

Electrolytic Cells: Construction and Electroplating - Procedure

Ogniwa elektrolityczne

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

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

Galvanic Cells

Galvanic Cells: Construction and Reduction Potential Measurement - Procedure

Ogniwa galwaniczne

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

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

Cell Division

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

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

Podział komórek

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

Cellular Processes

Cell Structure

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

Cell Structure: Visualizing Onion and Human Cells - Procedure

Struktura komórki

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

Fundamentals

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

Microbial and Fungal Diversity

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

Microbial Colony and Fungal Diversity - Procedure

Różnorodność mikrobiologiczna i grzybicza

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

Community Diversity

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

Community Diversity: Measuring Biodiversity in Different Areas - Procedure

Różnorodność społeczności

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

Ecology

Cellular Respiration

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

Cellular Respiration: Quantification using Microrespirometers - Procedure

Oddychanie komórkowe

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