Name: a protocol to characterize the morphological changes of clostridium difficile in response to antibiotic treatment
Uploaded: 2017-05-25T12:30:00.000+00:00
Duration: 12 min 58 s
Description: Watch this Scientific Journal Video about A Protocol to Characterize the Morphological Changes of Clostridium difficile in Response to Antibiotic Treatment at JoVE.com

<p class='jove_content'>These experiments have been supported by research grants from Merck and Co. and Summit, PLC. </p>

<p class="jove_title">1. Isolating <em>C. difficile</em> from Different Environmental or Clinical Sources</p>
<ol>
	<li>Environmental isolates: Using a pre-sterilized cotton gauge (lightly wetted with 0.85% NaCl), swab the surface of any area of interest (floor, door, handle, shelf, <em>etc.</em>).<sup class="xref">8</sup> Make sure to wear sterile gloves and place the swab in a sterilized tube after completed.</li>
	<li>Clinical isolates (stool): Plate 10 to 100 mg of clinical stool samples onto cefoxitin-cycloserine-fructose agar (CCFA) using an inoculation loop and incubate under strict anaerobic conditions for 48 - 72 h. Store isolated colonies of <em>C. difficile</em> stock in cryovials at -80 &#176;C for further analyses.<sup class="xref">7</sup><sup>,</sup><sup class="xref">9</sup><sup>,</sup><sup class="xref">10</sup></li>
	<li>Enrich the environmental swab samples in brain heart infusion (BHI) broth with 0.05% sodium taurocholate and place in an anaerobic chamber at 37 &#176;C for 5 days. Centrifuge 1 mL of the culture at 10,000 x g and resuspend the pellet in 100 &#181;L of ethanol.</li>
	<li>Plate the resuspended cells (50 &#181;L) onto cycloserinecefoxitin fructose agar (CCFA) plates and incubate in an anaerobic chamber at 37 &#176;C for 40 - 48 h. Store isolated colonies of <em>C. difficile</em> stock in cryovials at -80 &#176;C for further analyses.</li>
	<li>Test suspected <em>C. difficile</em> colonies using the latex agglutination reagent or PCR.</li>
</ol>
<p class="jove_title">2. Culturing <em>C. difficile</em> and Killing Kinetic Procedures</p>
<ol>
	<li>Grow purified environmental or clinical <em>C. difficile</em> strains on blood agar plates in an anaerobic chamber at 37 &#176;C for 48 h.</li>
	<li>Take one isolated colony with an inoculation loop, transfer it into 5 mL BHI medium in a 15 mL tube, and grow for 24 h in an anaerobic chamber at 37 &#176;C .</li>
	<li>Dilute the pre-cultures 1:100 to approximately 10<sup>6</sup> colony-forming units (CFU)/mL in fresh pre-reduced BHIS (BHI plus 5 g/L yeast extract and 1% L-cysteine) supplemented with 0.1% sodium taurocholate and the appropriate concentration of antibiotic (T0).</li>
	<li>Collect a 1 mL sample with a pipette at each time point (T0, T6, T24, T48) and plate/spread a small aliquot (100 &#181;L in serial dilutions) onto a blood agar plate. Let the cells grow on the blood agar plate for 48 h in an anaerobic chamber at 37 &#176;C and count the resulting number of colonies to determine CFU.</li>
</ol>
<p class="jove_title">3. Preparing Samples for Scanning Electron Microscopy</p>
<ol>
	<li>Collect 1 mL of cells from each time point in microcentrifuge tubes and centrifuge at 10,000 x g for 10 min. Discard the supernatant and wash cells in PBS.</li>
	<li>Centrifuge the samples at 10,000 x g for 10 min again and discard the supernatant. Re-dilute cells in 1 mL of 4% paraformaldehyde and incubate for 1 h at room temperature.</li>
	<li>Centrifuge samples again for 10 min at 10,000 x g and discard supernatant. Wash cells twice with distilled water and redilute in 100 &#181;L of distilled water. Adjust volume depending on the turbidity of the solution.<br />
	NOTE: Making multiple serial dilutions is a good idea.</li>
	<li>Label coverslips and add 40 &#181;L of the sample on it. Incubate for 15 min in a flow hood to evaporate the liquid and allow the cells to adhere to the coverslip. If liquid is still present, use a blower to remove the liquid.</li>
	<li>Place labeled coverslips with cells in a desk sputtering machine and tape down. Secure pure gold in the sputtering machine. Turn machine on and commence sputtering at low pressure (50 mTorr). Coat cells for 30 s at 80 mA, which translates to 20 nm of gold coating.</li>
	<li>Transfer coated cells to the scanning electron microscope.</li>
</ol>
<p class="jove_title">4. Imaging <em>C. difficile</em> Cells on a Scanning Electron Microscope</p>
<ol>
	<li>Vent the scanning electron microscope (SEM) properly by pressing the vent button on the computer software.</li>
	<li>Using carbon tape, secure the coated coverslips onto the metal stage. Once the SEM is vented, the door should easily open. Lock the metal stage into the SEM chamber by screwing it in.</li>
	<li>Click the PUMP button on the computer software. The SEM will be useable when the system reads &#34;Vac OK&#34;.</li>
	<li>Click on the SE detector under the detectors tab. Turn on the beam by clicking on the button that displays the voltage. Start imaging at a lower voltage (5 kV) before increasing voltage (up to 15 kV). An image will appear after the beam is turned on.</li>
	<li>Using the tracking function, find an area on the coated coverslips to image. Zoom into the region and find rod-shaped structures; these are <em>clostridium difficile</em>.</li>
	<li>Zoom in and focus the image to calibrate the system. This should be done at multiple working distances: 15, 9 and 5 mm. Image at a working distance of 5 mm.</li>
	<li>Switch on the ultra high resolution imaging mode and begin to focus and optimize astigmatism.</li>
	<li>Begin to focus at high magnification. Use the coarse and fine focus toggles for this. Adjust astigmatism as well to obtain a clearer image.
	<ol>
		<li>Adjust astigmatism at high magnification. To do this, adjust the astigmatism toggles (they look similar to the fine/coarse focus toggles) and check the image for clarity by digitally zooming in on the computer software.</li>
	</ol>
	</li>
	<li>se the slow scan function to collect a high quality image. Save the collected image as a .TIFF file, which will be used for analysis. Make sure the data bar is selected if measurements will be made during analysis.</li>
	<li>Collect images at different angles (up to 52&#176; by manually turning the angle directly on the SEM. Angled images tend to reveal more depth information.</li>
	<li>Change beam voltage depending on the cells that are being imaged. Keep this mostly between 5 - 15 kV for all experiments.</li>
	<li>After imaging is completed, turn the beam off and raise the working distance to 20 mm. The chamber can then be vented and the stage can be removed. Copy all of the images onto a drive for further analysis.</li>
</ol>
<p class="jove_title">5. Image Processing and Analysis</p>
<ol>
	<li>Download and install the freely available software, FIJI (http://fiji.sc), on to the computer.</li>
	<li>Open the image file in FIJI.</li>
	<li>Using the line function, precisely trace the scale bar.</li>
	<li>Click on the Analyze tab in the FIJI program and finally select the Select Scale function. A window will appear that will require the setting of the known distance based on the scale bar. Change the unit of length as well and click OK.</li>
	<li>Now that the program is calibrated for distance, use the line function to measure cell length. To obtain the length, use the line function to trace the cell in its entirety. Select the Analyze tab again and then click on Measure. The length should appear in the units denoted previously.</li>
</ol>

<ol>
	<li>Lessa, F. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Burden+of+Clostridium+difficile+infection+in+the+United+States.">Burden of Clostridium difficile infection in the United States.</a> <em style='font-style: italic'>N Engl J Med</em>. <b>372</b> (9), 825-834 (2015).</li><li>Vickers, R. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ridinilazole:+a+novel+therapy+for+Clostridium+difficile+infection.">Ridinilazole: a novel therapy for Clostridium difficile infection.</a> <em style='font-style: italic'>Int J Antimicrob Agents</em>. <b>48</b> (2), 137-143 (2016).</li><li>Shah, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clostridium+difficile+infection:+update+on+emerging+antibiotic+treatment+options+and+antibiotic+resistance.">Clostridium difficile infection: update on emerging antibiotic treatment options and antibiotic resistance.</a> <em style='font-style: italic'>Expert Rev Anti Infect Ther</em>. <b>8</b> (5), 555-564 (2010).</li><li>Ambrose, P. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+EMA+guideline+for+antimicrobial+development.">New EMA guideline for antimicrobial development.</a> <em style='font-style: italic'>Lancet Infect Dis</em>. <b>12</b> (4), 265-266 (2012).</li><li>Bass&#232;res, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impact+on+toxin+production+and+cell+morphology+in+Clostridium+difficile+by+ridinilazole+(SMT19969),+a+novel+treatment+for+C.+difficile+infection.">Impact on toxin production and cell morphology in Clostridium difficile by ridinilazole (SMT19969), a novel treatment for C. difficile infection.</a> <em style='font-style: italic'>J Antimicrob Chemother</em>. <b>71</b> (5), 1245-1251 (2016).</li><li>Endres, B. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+method+for+imaging+the+pharmacological+effects+of+antibiotic+treatment+on+Clostridium+difficile.">A novel method for imaging the pharmacological effects of antibiotic treatment on Clostridium difficile.</a> <em style='font-style: italic'>Anaerobe</em>. <b>40</b>, 10-14 (2016).</li><li>Endres, B. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluating+the+Effects+of+Surotomycin+Treatment+on+Clostridium+difficile+Toxin+A+and+B+Production,+Immune+Response,+and+Morphological+Changes.">Evaluating the Effects of Surotomycin Treatment on Clostridium difficile Toxin A and B Production, Immune Response, and Morphological Changes.</a> <em style='font-style: italic'>Antimicrob Agents Chemother</em>. <b>60</b> (6), 3519-3523 (2016).</li><li>Alam, M. J., Anu, A., Walk, S. T., Garey, K. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Investigation+of+potentially+pathogenic+Clostridium+difficile+contamination+in+household+environs.">Investigation of potentially pathogenic Clostridium difficile contamination in household environs.</a> <em style='font-style: italic'>Anaerobe</em>. <b>27</b>, 31-33 (2014).</li><li>Aitken, S. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+the+Endemic+Setting,+Clostridium+difficile+Ribotype+027+Is+Virulent+But+Not+Hypervirulent.">In the Endemic Setting, Clostridium difficile Ribotype 027 Is Virulent But Not Hypervirulent.</a> <em style='font-style: italic'>Infect Control Hosp Epidemiol</em>. , 1-6 (2015).</li><li>Basseres, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impact+on+toxin+production+and+cell+morphology+in+Clostridium+difficile+by+ridinilazole+(SMT19969),+a+novel+treatment+for+C.+difficile+infection.">Impact on toxin production and cell morphology in Clostridium difficile by ridinilazole (SMT19969), a novel treatment for C. difficile infection.</a> <em style='font-style: italic'>J Antimicrob Chemother</em>. <b>71</b> (5), 1245-1251 (2016).</li><li>Walters, B. A., Roberts, R., Stafford, R., Seneviratne, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Relapse+of+antibiotic+associated+colitis:+endogenous+persistence+of+Clostridium+difficile+during+vancomycin+therapy.">Relapse of antibiotic associated colitis: endogenous persistence of Clostridium difficile during vancomycin therapy.</a> <em style='font-style: italic'>Gut</em>. <b>24</b> (3), 206-212 (1983).</li><li>Chilton, C. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+the+effect+of+oritavancin+on+Clostridium+difficile+spore+germination,+outgrowth+and+recovery.">Evaluation of the effect of oritavancin on Clostridium difficile spore germination, outgrowth and recovery.</a> <em style='font-style: italic'>J Antimicrob Chemother</em>. <b>68</b> (9), 2078-2082 (2013).</li><li>Ofosu, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clostridium+difficile+infection:+a+review+of+current+and+emerging+therapies.">Clostridium difficile infection: a review of current and emerging therapies.</a> <em style='font-style: italic'>Ann Gastroenterol</em>. <b>29</b> (2), 147-154 (2016).</li><li>McDonald, L. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+epidemic,+toxin+gene-variant+strain+of+Clostridium+difficile.">An epidemic, toxin gene-variant strain of Clostridium difficile.</a> <em style='font-style: italic'>New Eng J Med</em>. <b>353</b> (23), 2433-2441 (2005).</li></ol>

<p class='jove_content'>KWG has received past and current research support from Merck &amp; Co. and Summit, PLC.</p>

<p class="jove_content">The goal of the current study was to create a high-throughput method for isolating <em>C. difficile </em>and testing antibiotic susceptibility using scanning electron microscopy (SEM) as a means for a more thorough characterization of the antibiotic&#39;s pharmacological action. Using the protocols outlined here, we have demonstrated that imaging the cell&#39;s phenotypic response to antibiotic treatment can reveal insight into the pharmacological action of the drug. In total, the imaging portion of this protocol takes r...

<p class="jove_content"><em>Clostridium difficile</em> is a gram positive, spore-forming bacterium, causing approximately 500,000 infections annually in the US and is considered a threat level urgent pathogen by the Centers for Disease Control and Prevention (CDC), the highest level of risk.<sup class="xref">1</sup> The past decade has seen considerable drug development in antimicrobials with activity against <em>C. difficile</em>.<sup class="xref">2</sup><sup>,</sup><sup class="xref">3</sup><em>In vitro </em>studies are a necessary component of the drug development process.<sup class="xref">4</sup> Traditionally, <em>in vitro</em> susceptibility and time kill studies are used to validate future animal and other <em>in vivo</em> studies.</p>
<p class="jove_content">While these methods serve an important role for evaluating killing action, they do not capture the cells&#39; phenotypic response to pharmacological treatment. By incorporating scanning electron microscopy (SEM) with standard killing kinetic studies, a more thorough characterization of the antibiotic direct effects is possible.<sup class="xref">5</sup><sup>,</sup><sup class="xref">6</sup><sup>,</sup><sup class="xref">7</sup> Here, we present a method where SEM is used as a means to profile the efficacy of antibiotic treatment.</p>

<table><tbody><tr><td>cotton gauze&amp;nbsp;</td><td>Caring</td><td>PRM21408C</td><td></td></tr><tr><td>NaCl</td><td>Macron</td><td>7532</td><td></td></tr><tr><td>50 mL tubes</td><td>Falcon</td><td>352098</td><td></td></tr><tr><td>Brain Heart Infusion (BHI)&amp;nbsp;</td><td>Criterion</td><td>C5141</td><td></td></tr><tr><td>L-cysteine</td><td>Alfa Aesar</td><td>A10389</td><td></td></tr><tr><td>yeast extract</td><td>Criterion</td><td>C741</td><td></td></tr><tr><td>sodium taurocholate</td><td>Alfa Aesar</td><td>A18346</td><td></td></tr><tr><td>anaerobic chamber</td><td>Coy</td><td></td><td>vinyl anaerobic chamber</td></tr><tr><td>cycloserinecefoxitin fructose agar (CCFA) plates</td><td>Anaerobe systems</td><td>AS-213</td><td></td></tr><tr><td>blood agar plates</td><td>Hardy diagnostics</td><td>A-10</td><td></td></tr><tr><td>latex agglutination reagent</td><td>Oxoid</td><td>DR1107A</td><td>C. diff test kit</td></tr><tr><td>microcentrifuge tubes</td><td>Eppendorf</td><td>222363204</td><td></td></tr><tr><td>PBS</td><td>Gibco</td><td>10010-031</td><td></td></tr><tr><td>4% paraformaldehyde</td><td>Fisher Scientific</td><td>50-259-98</td><td></td></tr><tr><td>microscope slides</td><td>J. Melvin freed brand</td><td>7525M</td><td>75 x 25mm</td></tr><tr><td>flow hood</td><td>Labconco</td><td>Class II type A2&amp;nbsp;</td><td>biosafety cabinet</td></tr><tr><td>desk sputtering machine</td><td>Denton Vacuum</td><td>Desk II</td><td></td></tr><tr><td>tape</td><td>Plastic Core</td><td>05072-AB</td><td>SPI Double Sided Adhesive Carbon Tape</td></tr><tr><td>gold</td><td>Denton Vacuum</td><td>TAR001-0158</td><td>2.375&amp;rdquo; Diameter x .002&amp;rdquo; Thick Gold foil</td></tr><tr><td>scanning electron microscope</td><td>FEI</td><td>XL-30</td><td></td></tr></tbody></table>

a protocol to characterize the morphological changes of <em>clostridium difficile</em> in response to antibiotic treatment

<p class="jove_content">Assessment of antibiotic action with new drug development directed towards anaerobic bacteria is difficult and technically demanding. To gain insight into possible MOA, morphologic changes associated with antibiotic exposure can be visualized using scanning electron microscopy (SEM). Integrating SEM imaging with traditional kill curves may improve our insight into drug action and advance the drug development process. To test this premise, kill curves and SEM studies were conducted using drugs with known but different MOA (vancomycin and metronidazole). <em>C. difficile </em>cells (R20291) were grown with or without the presence of antibiotic for up to 48 h. Throughout the 48 h interval, cells were collected at multiple time points to determine antibiotic efficacy and for imaging on the SEM. Consistent with previous reports, vancomycin and metronidazole had significant bactericidal activity following 24 h of treatment as measured by colony-forming unit (CFU) counting. Using SEM imaging we determined that metronidazole had significant effects on cell length (&#62; 50% reduction in cell length for each antibiotic; <em>P</em>&#60; 0.05) compared to controls and vancomycin. While the phenotypic response to drug treatment has not been documented previously in this manner, they are consistent with the drug&#39;s MOA demonstrating the versatility and reliability of the imaging and measurements and the application of this technique for other experimental compounds.</p>

<p class="jove_content" fo:keep-together.within-page="1"><em>Clostridium difficile</em> is a spore-forming bacterium and thus it is essential to determine the morphology differences between vegetative and spore cells prior to any functional analysis. <strong>Figure 1</strong> demonstrates representative images of vegetative cells that were captured during the exponential phase of the growth curve and spore cells. As depicted, vegetative cells are long, smooth, rod-shaped structures whereas spores are small, oval structures that have a rough ext...

Watch this Scientific Journal Video about A Protocol to Characterize the Morphological Changes of Clostridium difficile in Response to Antibiotic Treatment at JoVE.com

A Protocol to Characterize the Morphological Changes of Clostridium difficile in Response to Antibiotic Treatment

<p class='jove_content'>Antibiotic efficacy is most commonly determined by conducting killing kinetic studies and measuring colony forming units (CFUs). By integrating scanning electron microscopy (SEM) with these standard methods, we can distinguish the pharmacological effects of treatment between different antibiotics.</p>

A Protocol to Characterize the Morphological Changes of <em>Clostridium difficile</em> in Response to Antibiotic Treatment

Assessment of antibiotic action with new drug development directed towards anaerobic bacteria is difficult and technically demanding. To gain insight into ...

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Immunology and Infection

8.9K Views.  University of Houston College of Pharmacy. Antibiotic efficacy is most commonly determined by conducting killing kinetic studies and measuring colony forming units (CFUs). By integrating scanning electron microscopy (SEM) with these standard methods, we can distinguish the pharmacological effects of treatment between different antibiotics.

Video: A Protocol to Characterize the Morphological Changes of Clostridium difficile in Response to Antibiotic Treatment

Macromolecules

<h4>Biomolecules</h4>
<p>Organisms contain a wide variety of organic molecules with numerous functions which depend on the chemical structures and properties of these molecules. All organic molecules contain a carbon backbone and hydrogen atoms. The carbon atom is central in the formation of a vast variety of organic molecules ranging in size, shape and complexity; inorganic molecules on the other hand, generally have simpler structures. The outermost shell of a free carbon atom can accommodate eight electrons but is occupied by only four electrons, therefore it can form four covalent bonds and bond with up to four atoms. Alternatively, it can also bond with fewer atoms by forming double or triple bonds. This versatility of carbon atoms allows organic molecules to display intricate structures, such as chains, branches, and rings, among others.</p>
<p>Organic molecules that naturally occur in organisms are called biomolecules. Besides carbon and hydrogen, biomolecules also contain other elements such as oxygen, nitrogen, phosphorus, and sulfur. In general, smaller units of biomolecules come together, as repetitive sequences, to form larger biomolecules. These small modular units of biomolecules are called monomers. Two monomers typically join each other to form a dimer through a process known as <em>dehydration synthesis</em>, which is simply the removal of a hydrogen atom from one monomer and a hydroxyl (OH<sup>-</sup>) ion from the other monomer to create a water molecule to be expelled out while linking the two monomers with a covalent bond. The reverse of this process is called <em>hydrolysis</em>, in which the molecule splits back into its original monomers with a water molecule providing a hydrogen atom to one monomer and a hydroxyl ion to the other. Many monomers can be attached together by dehydration to form polymers. Sometimes different polymers can come together to form even larger and more complex molecules, which are known as biological macromolecules.</p>
<p>Biomolecules are classified based on the elements that compose them, and their structure and function inside living organisms. Almost all biomolecules can be classified into one of the four general categories: carbohydrates, lipids, proteins, and nucleic acids.</p>
<h4>Carbohydrates</h4>
<p>Carbohydrate simply means &#8220;carbon water&#8221; because these molecules are composed of carbon, hydrogen, and oxygen atoms roughly in the ratio of 1:2:1. Carbohydrate monomers are known as monosaccharides, which are also referred to as <em>simple sugars</em>. Glucose (C<sub>6</sub>H<sub>12</sub>O<sub>6</sub>) is the most common monosaccharide in living organisms and is a subunit of many polysaccharides. Numerous organisms also synthesize other six-carbon monosaccharides with the same chemical formula as glucose but slightly different structures, such as fructose and galactose. When two monosaccharides are linked together, they form disaccharides. For example, sucrose is composed of glucose and fructose, whereas lactose contains glucose and galactose. These monosaccharides and disaccharides are used for short-term energy storage in living organisms.  Maltose is another disaccharide that is made up of two glucose molecules and is usually formed when polysaccharide chains such as starch and glycogen are broken down during digestion. Starch is a polysaccharide that serves as an energy storage molecule in plants and is made up of two types of glucose polymers: amylose and amylopectin. Amylose constitutes 10-20% of starch and is a helical polymer of glucose. Amylopectin makes up the bulk of the starch and is a branched polymer of glucose. Glycogen is virtually the same as starch, however it is synthesized, stored and used in animal liver and muscle tissues.</p>
<p>Besides serving as energy stores, carbohydrates also have other functions in organisms. The five-carbon monosaccharides, ribose and deoxyribose, are integrated into the nucleic acid structure and are present in every living cell. Moreover, the polysaccharide cellulose, which is a long polymer made up of glucose, serves as a rigid structural material in plants. Humans do not have digestive enzymes to break down cellulose in food, which is also called dietary fiber. However, dietary fiber consumption helps to maintain a healthy gut flora, which in turn contributes to the health of digestive and immune systems<sup>1</sup>. Similar to plants, some animals and fungi use another polysaccharide, chitin, as a structural molecule. Arthropods use chitin to build and maintain their exoskeletons, whereas fungi incorporate it into their cell walls to maintain rigidity.</p>
<h4>Lipids</h4>
<p>The second class of biological macromolecules are lipids, which include fats, oils, and waxes. Lipids are hydrophobic molecules that are almost entirely made up of carbon and hydrogen atoms. Often, lipids are grouped in three major categories; triglycerides, phospholipids, and steroids.</p>
<p>The most common type of lipid is triglycerides, which include fats from animals and oils from plants. Triglycerides generally serve as long-term energy storage molecules, except indigestible waxes, which are instead used as a waterproofing substance in both plants and animals. Triglycerides contain three fatty acid chains, which can be either saturated or unsaturated, connected to a glycerol molecule. Saturated fatty acid chains are linear molecules with a maximum number of hydrogen atoms, where every carbon in the chain is connected via a single bond. On the other hand, unsaturated fatty acid chains have kinks due to the presence of at least one double bond. Additionally, unsaturated fats can be &#8220;trans&#8221; fats if the hydrogens around the double bond oppose each other. While trans fats occur naturally, they are generated during industrial production of saturated vegetable oils with hydrogen. Similar to saturated fatty acids, trans fats stack very well due to their relative linearity. However, trans fats cause problems for human heart health, such as the damaging the lining of arteries and causing inflammation when digested<sup>2</sup>.</p>
<p>Phospholipids are similar to triglycerides, however, one of the fatty acid chains is replaced with a phosphate-containing polar group. Therefore, phospholipids have a hydrophilic head and two hydrophobic fatty acid tails. These properties of phospholipids are crucial to the cell membrane structure and function.</p>
<p>Steroids are lipids that are composed of fused carbon rings with varying functional groups. Cholesterol is a steroid that is also a cell membrane component. Moreover, cholesterol is used to synthesize other steroids, including sex hormones such as estrogen and testosterone. Although cholesterol is essential for cell membrane structure and hormone synthesis, high levels of plasma cholesterol are implicated in plaque accumulation inside blood vessels and causing coronary disease<sup>3</sup>.</p>
<h4>Proteins</h4>
<p>The third class of biological macromolecules are proteins, which are made up of chains of amino acids. There are 20 different amino acids, all with a similar base structure but each has a unique side chain called an &#8216;R-group&#8217;. A single amino acid has an N-terminal end which is an amino group (NH<sub>3</sub><sup>+</sup>) and a C-terminal end which is a carboxyl group (-COOH). These groups link together, N-terminal to C-terminal, in a chain connected by peptide bonds. Proteins are important for maintaining body functions as enzymes, hormones, structural components and transport molecules, and play vital roles in muscle contractibility, immunity and blood clotting. However, issues can arise in protein structure and function, and these issues are often genetic. For instance, normal red blood cells are round, but in people affected by sickle cell anemia, cells have a curved shape with an exposed hydrophobic region, caused by a mutation in a protein called hemoglobin S. This shape reduces the capacity to carry oxygen and causes the cells to get stuck in blood vessels. This results in many detrimental symptoms to the person carrying the mutation, and people who inherit two copies of the sickle cell gene often suffer ill effects or even possibly die because of the reduced capacity of sickled cells to transport oxygen. In a twist, those carrying only one copy of the gene are resistant to infection from malaria, so the disease has been able to be passed on and persists in countries with elevated levels of malaria infections<sup>4</sup>.</p>
<h4>Nucleic Acids</h4>
<p>The fourth class of biological macromolecules are the nucleic acids, which are composed of monomers known as nucleotides. These monomers are composed of three parts: a phosphate group, a ribose sugar and a nitrogen base. Nucleotides differ from each other by their nitrogen bases and the type of ribose they contain.  Single nucleotides generally act as energy carriers inside cells, as well as function as messenger molecules. However, the nucleotide polymers or nucleic acids such as deoxyribonucleic acid (DNA) or ribonucleic acid (RNA) are hereditary molecules that contain the genetic information to construct cellular products.</p>
<h4>Macromolecule Detection</h4>
<p>Biological macromolecules in food or other substances can be detected by using their specific chemical properties. For example, monosaccharides have free aldehyde (-CHO) or ketone (-C=O) groups that can reduce other compounds, meaning they are substances that cause other molecules to lose electrons, thus monosaccharides are also known as <em>reducing sugars</em>. This property is used to detect the presence of monosaccharides with an indicator called Benedict&#8217;s reagent.  This indicator contains copper ions (Cu<sup>2+</sup>) which are reduced by monosaccharides as observed with a change of the solution color from blue to a reddish orange, though the intensity of the color varies based on the original concentration of the reducing sugar. As such, observation of a green color means that there is only a small amount of reducing sugars present in the solution. Amylose in starch exists in a coiled structure because of the bond angles in the polymer chain. The iodine indicator, iodine-potassium iodide (IKI), reacts with these coiled molecules and will turn the solution a dark bluish-black to indicate the presence of amylose. However, if starch amylose isn&#8217;t present, the reaction will not take place, and the solution will remain a yellowish-brown color.  A Sudan IV test is performed to test for the presence of lipids. This dye is lipophilic and solubilizes when lipids are present, thus a red color is retained in the presence of lipids. Biuret&#8217;s reagent, an indicator for the presence of proteins, contains copper ions (Cu<sup>2+</sup>) which react with the peptide bonds and turn the solution from blue to dark violet color. This reagent needs to react with a sufficient number of these peptide bonds to produce the expected violet color, so a pinkish color will result if the amino acid chains are not long enough.</p>
<h4>References</h4>
<ol class="references">
	<li><strong>Rasnik K. Singh, Hsin-Wen Chang, Di Yan, Kristina M. Lee, Derya Ucmak, Kirsten Wong, Michael Abrouk, Benjamin Farahnik, Mio Nakamura, Tian Hao Zhu, Tina Bhutani, Wilson Liao.</strong> Influence of diet on the gut microbiome and implications for human health. <em>J Transl Med. </em>2017, Vol. 15, 73 doi: 10.1186/s12967-017-1175-y.</li>
	<li><strong>Siri-Tarino PW, Sun Q, Hu FB, Krauss RM.</strong> Meta-analysis of prospective cohort studies evaluating the association of saturated fat with cardiovascular disease. <em>Am J Clin Nutr. </em>2010, Vol. 91, 3 (535-46).</li>
	<li><strong>Joseph L. Goldstein, Michael S. Brown.</strong> A Century of Cholesterol and Coronaries: From Plaques to Genes to Statins. <em>Cell. </em>2015, Vol. 161, 1 (161-72).</li>
	<li><strong>Michael Aidoo, Dianne J Terlouw, Margarette S Kolczak, Peter D McElroy, Feiko O ter Kuile, Simon Kariuki, Bernard L Nahlen, Altaf A Lal, Venkatachalam Udhayakumar.</strong> Protective effects of the sickle cell gene against malaria morbidity and mortality. <em>Lancet. </em>2002, Vol. 359, 9314 (1311-1312).</li>
</ol>

Macromolecules: Identification Using Indicator Solutions - Concept

Makrocząsteczek

Biomolecules
Organisms contain a wide variety of organic molecules with numerous functions which depend on the chemical structures and properties of these ...

Education

JoVE Lab Manual

Biology

Fundamentals

UV-Vis Spectroscopy of Dyes

<p>Source: Lara Al Hariri and Ahmed Basabrain at the University of Massachusetts Amherst, MA, USA</p>
<ol class="note-items">
	<li data-note-item="1" data-parent-collapse="">Absorbance Wavelength Comparison<button class="expand">Expand</button>
	<p>In the first part of the lab, you&#39;ll use ultraviolet and visible light absorption spectroscopy, or UV-Vis absorption spectroscopy, to analyze the absorption characteristics of fluorescein, &#946;-carotene, and indigo dye. To perform UV-Vis spectroscopy, you&#39;ll place a solution between a light source and a photodetector. The molecules will absorb light at wavelengths that correspond to the energies needed to excite their electrons and scatter or transmit the rest.</p>
	<p>An absorption spectrum represents the variation in the number of photons of each wavelength that reach the detector. A higher absorbance corresponds to fewer detected photons of that wavelength. You&#39;ll take a spectrum of the pure solvent first, which the instrument will subtract from the spectrum of the dye solution to show you just the absorbance data from the dye. This reference is called a solvent blank.</p>
	<ul class="lab-items">
		<li data-sub-note-item="1-3">
		<div class="lab-step">Before you start, put on a lab coat, safety glasses, and gloves. <strong>Note: </strong>The &#946;-carotene and indigo solutions use hexane and dimethylformamide, respectively, so you will work with the dyes in a fume hood. DMF and hexane will degrade thin nitrile gloves, so change your gloves if you get solvent on them.</div>
		</li>
		<li data-sub-note-item="1-4">
		<div class="lab-step">Now, obtain a clean quartz cuvette. Cuvettes typically have two transparent sides and two textured or opaque sides. Always hold the cuvette by the textured sides to keep the transparent sides clean.</div>
		</li>
		<li data-sub-note-item="1-5">
		<div class="lab-step">First, you&#39;ll analyze a solution of fluorescein in water. To make your solvent blank, fill your cuvette about 3/4 full with deionized water.</div>
		</li>
		<li data-sub-note-item="1-6">
		<div class="lab-step">Now, clean the transparent sides of the cuvette with a laboratory wipe. Check the cuvette for air bubbles and gently tap the cuvette to dislodge them.</div>
		</li>
		<li data-sub-note-item="1-7">
		<div class="lab-step">Then, bring the cuvette to the spectrometer. Open the sample chamber of the spectrometer and clean your cuvette one last time. Smudges and dust can absorb or reflect light, which will introduce errors into the data.</div>
		</li>
		<li data-sub-note-item="1-8">
		<div class="lab-step">Now, insert the cuvette into the sample holder so that the transparent sides are in line with the light source. Close the sample chamber door completely. Follow the instructions for your spectrometer software to set up a scan of absorbance from 200 &#8211; 800 nm and scan your cuvette as the solvent blank.</div>
		</li>
		<li data-sub-note-item="1-9">
		<div class="lab-step">When the scan finishes, empty the cuvette and use compressed air to remove large drops of water.</div>
		</li>
		<li data-sub-note-item="1-10">
		<div class="lab-step">Then, bring it to the shared hood and use a clean Pasteur pipette to fill the cuvette 3/4 full of the provided 3 &#181;M fluorescein solution.</div>
		</li>
		<li data-sub-note-item="1-11">
		<div class="lab-step">Check the cuvette for air bubbles and remove them as needed before bringing the cuvette to the spectrometer. Remember to clean the transparent sides one last time before placing the cuvette in the sample chamber. Then, scan the cuvette as your sample.</div>
		</li>
		<li data-sub-note-item="1-12">
		<div class="lab-step">When the scan finishes, identify the wavelengths that fluorescein absorbed. The wavelength with the highest absorbance is called &#955;<sub>max</sub>.</div>
		</li>
		<li data-sub-note-item="1-13">
		<div class="lab-step">Record the wavelengths of the absorbance peaks and the overall absorption range in your lab notebook and then save the data.
		<h4><strong>Table 1: Absorbance and &#955;<sub>max</sub> of dyes</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>Absorbance range (nm)</strong></td>
					<td style="text-align: center;"><strong>&#955;<sub>max</sub> (nm)</strong></td>
				</tr>
				<tr>
					<td>&#160;Fluorescein</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td>&#160;&#946;-carotene</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td>&#160;Indigo</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
			</tbody>
		</table>
		<a href="https://www.jove.com/CDNSource/lm/labs/70/Table_1._Absorbance_and_?max_of_dyes.xlsx" target="_blank">Click Here to download Table 1</a></div>
		</li>
		<li data-sub-note-item="1-14">
		<div class="lab-step">Now, empty your cuvette into the aqueous waste and rinse it with water to remove traces of fluorescein.</div>
		</li>
		<li data-sub-note-item="1-15">
		<div class="lab-step">You&#39;ll analyze &#946;-carotene next, so you need a hexane solvent blank. Rinse the cuvette with hexane a few times to remove leftover water before filling it 3/4 full of pure hexane.</div>
		</li>
		<li data-sub-note-item="1-16">
		<div class="lab-step">Remove air bubbles, clean the sides of the cuvette, and place it in the spectrometer. Make sure that the scan is set for absorbance from 200 &#8211; 800 nm, and then scan the cuvette as a solvent blank.</div>
		</li>
		<li data-sub-note-item="1-17">
		<div class="lab-step">When the scan finishes, empty the cuvette into the organic waste and dry it with compressed air.</div>
		</li>
		<li data-sub-note-item="1-18">
		<div class="lab-step">Use a clean pipette to fill the cuvette 3/4 full of the provided 3 &#181;M &#946;-carotene solution and return to the spectrometer.</div>
		</li>
		<li data-sub-note-item="1-19">
		<div class="lab-step">Clean the transparent sides of the cuvette, place it in the spectrometer, and scan it as your sample. Write the wavelengths of the peaks in your lab notebook and save the data.</div>
		</li>
		<li data-sub-note-item="1-20">
		<div class="lab-step">Now, analyze the indigo dye. The indigo solution is in DMF, so empty your cuvette into the organic waste and rinse out the leftover &#946;-carotene and hexane with pure DMF.</div>
		</li>
		<li data-sub-note-item="1-21">
		<div class="lab-step">Fill the cuvette 3/4 full of pure DMF, place it in the spectrometer, and make sure that the scan settings are the same as before. Run the solvent blank scan, and then empty the cuvette into the organic waste and dry it with compressed air.</div>
		</li>
		<li data-sub-note-item="1-22">
		<div class="lab-step">Use a clean pipette to fill the cuvette 3/4 full of the provided 3 &#181;M indigo dye solution and scan it as your sample. Write down the absorbance information and save your data.</div>
		</li>
		<li data-sub-note-item="1-23">
		<div class="lab-step">Lastly, empty your cuvette into the organic waste and rinse it with acetone.</div>
		</li>
	</ul>
	</li>
	<li data-note-item="2" data-parent-collapse="">&#946;-Carotene Calibration Curve<button class="expand">Expand</button>
	<p>In the second half of the lab, you&#39;ll use UV-Vis spectroscopy to relate absorbance intensity and concentration for &#946;-carotene by measuring the absorbances of 5 &#946;-carotene solutions with different concentrations. This will let you make a calibration curve of absorbance versus &#946;-carotene concentration and derive the equation describing that relationship.</p>
	<ul class="lab-items">
		<li data-sub-note-item="2-2">
		<div class="lab-step">To begin, make a table in your lab notebook with columns for &#946;-carotene concentration and absorbance at 450 nm.
		<h4><strong>Table 2: Absorbance of &#946;-carotene</strong></h4>
		<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
			<tbody>
				<tr>
					<td><strong>Test </strong></td>
					<td><strong>Concentration (&#181;M) </strong></td>
					<td><strong>Absorbance at &#955;<sub>max</sub> (450 nm) </strong></td>
				</tr>
				<tr>
					<td style="text-align: center;">1</td>
					<td style="text-align: center;">1.9</td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td style="text-align: center;">2</td>
					<td style="text-align: center;">3.7</td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td style="text-align: center;">3</td>
					<td style="text-align: center;">7.5</td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td style="text-align: center;">4</td>
					<td style="text-align: center;">11</td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td style="text-align: center;">5</td>
					<td style="text-align: center;">15</td>
					<td style="text-align: center;"></td>
				</tr>
			</tbody>
		</table>
		<a href="https://www.jove.com/CDNSource/lm/labs/70/Table_2._Absorbance_of_?-carotene .xlsx" target="_blank">Click Here to download Table 2</a></div>
		</li>
		<li data-sub-note-item="2-3">
		<div class="lab-step">Then, get a clean cuvette and fill it 3/4 full of hexane as your solvent blank.</div>
		</li>
		<li data-sub-note-item="2-4">
		<div class="lab-step">Remove any air bubbles, clean the transparent sides of the cuvette, and place it in the spectrometer. Set up an absorbance scan from 200 &#8211; 800 nm and scan the cuvette as the solvent blank.</div>
		</li>
		<li data-sub-note-item="2-5">
		<div class="lab-step">When the scan finishes, empty the cuvette into the organic waste and dry it with compressed air.</div>
		</li>
		<li data-sub-note-item="2-6">
		<div class="lab-step">Then, use a clean pipette to fill the cuvette 3/4 full of the 1.9 &#181;M &#946;-carotene solution, which is the lowest concentration solution.</div>
		</li>
		<li data-sub-note-item="2-7">
		<div class="lab-step">Clean the sides of the cuvette, insert it into the spectrometer, and scan it as a sample.</div>
		</li>
		<li data-sub-note-item="2-8">
		<div class="lab-step">When the scan finishes, display the absorbance intensity of the peak at 450 nm. Record this in your lab notebook as the absorbance for the 1.9 &#181;M solution.</div>
		</li>
		<li data-sub-note-item="2-9">
		<div class="lab-step">Now, empty the cuvette into the organic waste, rinse it with hexane, and dry it with compressed air.</div>
		</li>
		<li data-sub-note-item="2-10">
		<div class="lab-step">Use a clean pipette to fill the cuvette 3/4 full of the 3.7 &#181;M &#946;-carotene solution. Clean the sides of the cuvette, insert it into the spectrometer, and scan it as another sample.</div>
		</li>
		<li data-sub-note-item="2-11">
		<div class="lab-step">When the scan finishes, write down the absorbance of the peak at 450 nm. Repeat this for the remaining solutions, working from lowest to highest concentration. Save your data when you&#39;re done.</div>
		</li>
		<li data-sub-note-item="2-12">
		<div class="lab-step">Lastly, empty the cuvette into the organic waste and clean it with acetone.</div>
		</li>
	</ul>
	</li>
	<li data-note-item="3" data-parent-collapse="">Results<button class="expand">Expand</button>
	<p>First, look at the spectra of the three dyes. Fluorescein in water absorbs blue and purple light, with the maximum absorbance at 490 nm. It does not absorb red light, and it only absorbs some yellow and green light. Consistent with this, solid fluorescein is red, and fluorescein solutions are usually yellow to green.</p>
	<p>&#946;-carotene also absorbs blue and purple light. The maximum absorbance of &#946;-carotene in hexane is 450 nm, and you&#39;ll see another large peak at 478 nm. The strong absorption of purple light is part of why &#946;-carotene appears orange.</p>
	<p>Indigo in DMF absorbs UV light and red, orange, and yellow light, with a distinct peak at 611 nm. Thus, indigo dye reflects primarily blue and purple light, giving it its characteristic color.</p>
	<ul class="lab-items">
		<li data-sub-note-item="3-4">
		<div class="lab-step">Now, make the calibration curve for &#946;-carotene concentration. Plot the &#946;-carotene absorbance values at 450 nm from the second half of the lab against the concentrations of the solutions.</div>
		</li>
		<li data-sub-note-item="3-5">
		<div class="lab-step">Then, apply a trend line and find the linear equation that fits the data. According to the Beer-Lambert law, in this equation, y is absorbance, x is concentration, and the slope is the product of the relevant molar attenuation coefficient and the pathlength.</div>
		</li>
		<li data-sub-note-item="3-6">
		<div class="lab-step">Check your equation. Rearrange it to solve for concentration, and fill in the &#946;-carotene absorbance at 450 nm from the first part of the lab. The calculated concentration should be close to 3 &#181;M/L.</div>
		</li>
	</ul>
	</li>
</ol>

Ultraviolet-Visible UV-Vis Spectroscopy of Dyes - Procedure

Spektroskopia UV-VIS barwników

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

	Absorbance Wavelength ComparisonExpand
	In the first part ...

Chemistry

Lab Manual Sub Articles

Concentration Dependence

<p>Source: Smaa Koraym at Johns Hopkins University, MD, USA</p>
<ol class="note-items">
	<li data-note-item="1" data-parent-collapse="">Concentration Dependence<button class="expand">Expand</button>
	<p>In this experiment, you will combine varying concentrations of aqueous HCl and sodium thiosulfate to make solid sulfur, which rapidly gathers into opaque yellow particles at a certain sulfur concentration. Since the reaction solution starts clear and colorless, you can easily tell when it reaches that concentration.</p>
	<p>You&#39;ll use the same test tube and total volume each time, so it will take the same amount of sulfur to make each solution completely opaque. Thus, you&#39;ll measure the reaction progress by timing how long that takes. You&#39;ll then use that data to estimate the reaction orders of the individual reactants and of the overall reaction.</p>
	<p>Before starting the lab, make a table listing the reactant concentrations, the time to solution opacity, and the solution temperature for the trials.</p>
	<h4><strong>Table 1: Reaction progress</strong></h4>
	<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
		<tbody>
			<tr>
				<td><strong>Trial</strong></td>
				<td><strong>[Na<sub>2</sub>S<sub>2</sub>O<sub>3</sub>] (M)</strong></td>
				<td><strong>[HCl] (M)</strong></td>
				<td><strong>Time (s)</strong></td>
				<td><strong>Temp (&#176;C)</strong></td>
			</tr>
			<tr>
				<td style="text-align: center;">1</td>
				<td style="text-align: center;">0.1 M</td>
				<td style="text-align: center;">3.0 M</td>
				<td></td>
				<td></td>
			</tr>
			<tr>
				<td style="text-align: center;">2</td>
				<td style="text-align: center;">0.1 M</td>
				<td style="text-align: center;">3.0 M</td>
				<td></td>
				<td></td>
			</tr>
			<tr>
				<td style="text-align: center;">3</td>
				<td style="text-align: center;">0.2 M</td>
				<td style="text-align: center;">3.0 M</td>
				<td></td>
				<td></td>
			</tr>
			<tr>
				<td style="text-align: center;">4</td>
				<td style="text-align: center;">0.15 M</td>
				<td style="text-align: center;">3.0 M</td>
				<td></td>
				<td></td>
			</tr>
			<tr>
				<td style="text-align: center;">5</td>
				<td style="text-align: center;">0.05 M</td>
				<td style="text-align: center;">3.0 M</td>
				<td></td>
				<td></td>
			</tr>
			<tr>
				<td style="text-align: center;">6</td>
				<td style="text-align: center;">0.1 M</td>
				<td style="text-align: center;">6.0 M</td>
				<td></td>
				<td></td>
			</tr>
			<tr>
				<td style="text-align: center;">7</td>
				<td style="text-align: center;">0.1 M</td>
				<td style="text-align: center;">4.5 M</td>
				<td></td>
				<td></td>
			</tr>
			<tr>
				<td style="text-align: center;">8</td>
				<td style="text-align: center;">0.1 M</td>
				<td style="text-align: center;">1.5 M</td>
				<td></td>
				<td></td>
			</tr>
		</tbody>
	</table>
	<p><a href="https://www.jove.com/CDNSource/lm/labs/50/Table_1._Reaction_progress.xlsx" target="_blank">Click Here to download Table 1</a></p>
	<p>We want to make sure that the reactions are all taking place at room temperature. You&#39;ll perform two benchmark trials, three trials with different sodium thiosulfate concentrations, and three trials with different HCl concentrations. You will vary the concentrations by diluting stock solutions of sodium thiosulfate and HCl, as shown in the following tables.</p>
	<h4><strong>Table 2: Sodium thiosulfate dilutions</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>Target Concentration</strong></td>
				<td style="text-align: center;"><strong>Volume of 0.2 M sodium thiosulfate</strong></td>
				<td style="text-align: center;"><strong>Volume of DI water</strong></td>
			</tr>
			<tr>
				<td style="text-align: center;">0.05 M</td>
				<td style="text-align: center;">5 mL</td>
				<td style="text-align: center;">15 mL</td>
			</tr>
			<tr>
				<td style="text-align: center;">0.10 M</td>
				<td style="text-align: center;">10 mL</td>
				<td style="text-align: center;">10 mL</td>
			</tr>
			<tr>
				<td style="text-align: center;">0.15 M</td>
				<td style="text-align: center;">15 mL</td>
				<td style="text-align: center;">5 mL</td>
			</tr>
			<tr>
				<td style="text-align: center;">0.20 M</td>
				<td style="text-align: center;">20 mL</td>
				<td style="text-align: center;">0 mL</td>
			</tr>
		</tbody>
	</table>
	<p><a href="https://www.jove.com/CDNSource/lm/labs/50/Table_2._Sodium_thiosulfate_dilutions.xlsx" target="_blank">Click Here to download Table 2</a></p>
	<h4><strong>Table 3: HCl Dilutions</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>Target Concentration</strong></td>
				<td style="text-align: center;"><strong>Volume of 6.0 M HCl</strong></td>
				<td style="text-align: center;"><strong>Volume of DI water</strong></td>
			</tr>
			<tr>
				<td style="text-align: center;">1.5 M</td>
				<td style="text-align: center;">2.5 mL</td>
				<td style="text-align: center;">7.5 mL</td>
			</tr>
			<tr>
				<td style="text-align: center;">3.0 M</td>
				<td style="text-align: center;">5.0 mL</td>
				<td style="text-align: center;">5.0 mL</td>
			</tr>
			<tr>
				<td style="text-align: center;">4.5 M</td>
				<td style="text-align: center;">7.5 mL</td>
				<td style="text-align: center;">2.5 mL</td>
			</tr>
			<tr>
				<td style="text-align: center;">6.0 M</td>
				<td style="text-align: center;">10 mL</td>
				<td style="text-align: center;">0 mL</td>
			</tr>
		</tbody>
	</table>
	<p><a href="https://www.jove.com/CDNSource/lm/labs/50/Table_3._HCl_Dilutions.xlsx" target="_blank">Click Here to download Table 3</a></p>
	<p>Remember to use caution when handling HCl, which is toxic and highly corrosive. Sulfur dioxide, which is a gaseous product of this reaction, is also toxic. You will leave the reaction waste in the fume hood overnight to let the sulfur dioxide escape harmlessly.</p>
	<ul class="lab-items">
		<li data-sub-note-item="1-6">
		<div class="lab-step">First, put on a lab coat, splash-proof safety glasses, and nitrile gloves.</div>
		</li>
		<li data-sub-note-item="1-7">
		<div class="lab-step">Fill a plastic wash bottle with deionized water and get a test tube marked with an X.</div>
		</li>
		<li data-sub-note-item="1-8">
		<div class="lab-step">Clamp the test tube over the center of a small stir plate. The label must be on the opposite side from you. Make sure that you can clearly see the X. Place a small magnetic stir bar in the test tube.</div>
		</li>
		<li data-sub-note-item="1-9">
		<div class="lab-step">Use a thermometer clamp to secure a glass thermometer in the lower third of the tube, keeping it clear of the X and the stir bar.</div>
		</li>
		<li data-sub-note-item="1-10">
		<div class="lab-step">Cut a sheet of about 20 squares of plastic paraffin film from a roll and bring the sheet back to your hood.</div>
		</li>
		<li data-sub-note-item="1-11">
		<div class="lab-step">Lay paper towels in the fume hood as a clean surface for labware to be reused.</div>
		</li>
		<li data-sub-note-item="1-12">
		<div class="lab-step">Label a 600-mL beaker as &#8216;waste&#8217; for the reaction, a 100-mL beaker as &#8216;0.2 M sodium thiosulfate&#8217;, and a 50-mL beaker as &#8216;6 M HCl&#8217;.</div>
		</li>
		<li data-sub-note-item="1-13">
		<div class="lab-step">Fill the 100-mL beaker with 0.2 M sodium thiosulfate from the stock bottle in the dispensing hood.</div>
		</li>
		<li data-sub-note-item="1-14">
		<div class="lab-step">After that, bring the 50-mL beaker and a watch glass to the acid-dispensing hood to obtain 50 mL of 6 M HCl. Cover the filled beaker with the watch glass and bring your portion of HCl back to your hood.</div>
		</li>
		<li data-sub-note-item="1-15">
		<div class="lab-step">Prepare 20 mL of 0.1 M sodium thiosulfate for the first benchmark trial. Use a 10-mL volumetric pipette to measure 10 mL of 0.2 M sodium thiosulfate and dispense it into a 20-mL volumetric flask. Fill the 20-mL volumetric flask with deionized water to the mark.</div>
		</li>
		<li data-sub-note-item="1-16">
		<div class="lab-step">Then, cut a square of plastic paraffin film, seal the flask, and invert it several times to thoroughly mix the solution.</div>
		</li>
		<li data-sub-note-item="1-17">
		<div class="lab-step">Remove the film and pour the solution into the test tube.</div>
		</li>
		<li data-sub-note-item="1-18">
		<div class="lab-step">Prepare 10 mL of 3 M HCl. Pipette 5 mL of 6 M HCl into a 10-mL graduated cylinder. Add deionized water to fill the graduated cylinder to the 10-mL line.</div>
		</li>
		<li data-sub-note-item="1-19">
		<div class="lab-step">Seal the top of the graduated cylinder with plastic paraffin film and thoroughly mix the HCl solution by inverting it several times.</div>
		</li>
		<li data-sub-note-item="1-20">
		<div class="lab-step">Now, start the stir motor and get a reset stopwatch. Confirm that you can see the X through the solution in the test tube.</div>
		</li>
		<li data-sub-note-item="1-21">
		<div class="lab-step">Simultaneously start the stopwatch and pour the 3 M HCl into the test tube.</div>
		</li>
		<li data-sub-note-item="1-22">
		<div class="lab-step">Watch the X through the solution, which will start becoming cloudy when enough sulfur has been produced to saturate it. Stop the stopwatch exactly when the solution becomes so opaque that the X disappears.</div>
		</li>
		<li data-sub-note-item="1-23">
		<div class="lab-step">Record the time in your lab notebook for benchmark trial 1 and then reset the stopwatch. Also, record the temperature of the solution.</div>
		</li>
		<li data-sub-note-item="1-24">
		<div class="lab-step">Now, turn off the stir motor, remove the thermometer, and rinse it with deionized water into the waste beaker.</div>
		</li>
		<li data-sub-note-item="1-25">
		<div class="lab-step">Then, empty the test tube into the waste beaker, retrieve the stir bar with long forceps, and rinse the test tube and stir bar with deionized water.</div>
		</li>
		<li data-sub-note-item="1-26">
		<div class="lab-step">Thoroughly clean the test tube with a test tube brush. Briefly rinse the volumetric flask and graduated cylinder with deionized water as well. Dry everything with paper towels.</div>
		</li>
		<li data-sub-note-item="1-27">
		<div class="lab-step">Repeat the trial in the exact same way for the second benchmark time. If the time is not within 3 - 5 s of the first trial, repeat the benchmark trial until you have two consistent trials.</div>
		</li>
		<li data-sub-note-item="1-28">
		<div class="lab-step">Follow the same overall procedure for each subsequent trial, changing the dilutions as indicated in your table and cleaning your glassware between each trial. Use a 5-mL volumetric pipette as needed when measuring the sodium thiosulfate stock solution in trials four and five.</div>
		</li>
		<li data-sub-note-item="1-29">
		<div class="lab-step">Once you have performed all eight trials, rinse your glassware and tools one last time and neutralize the reaction waste with baking soda. Use caution because sulfur dioxide is one of the reaction products.</div>
		</li>
		<li data-sub-note-item="1-30">
		<div class="lab-step">Transfer the neutralized reaction waste to the designated waste container. Then, pour any remaining sodium thiosulfate solution into its labeled waste container and rinse the beaker with deionized water.</div>
		</li>
		<li data-sub-note-item="1-31">
		<div class="lab-step">Neutralize any remaining HCl with baking soda before flushing it down the drain with tap water.</div>
		</li>
		<li data-sub-note-item="1-32">
		<div class="lab-step">Finally, clean and put away your labware according to your standard practices and throw out any trash remaining in your hood. Your instructor will dispose of the reaction waste later.</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">Now, estimate the reaction order of sodium thiosulfate and HCl. Calculate the starting concentration of sodium thiosulfate in the reaction mixture for trials 1 through 5. Looking at the concentration of sodium thiosulfate and the reaction times, we see that as the concentration doubles, the time to opacity is halved.
		<h4><strong>Table 4: Estimating reaction order sodium thiosulfate</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>[Na<sub>2</sub>S<sub>2</sub>O<sub>3</sub>] added</strong></td>
					<td style="text-align: center;"><strong>Na<sub>2</sub>S<sub>2</sub>O<sub>3</sub> volume (mL)</strong></td>
					<td style="text-align: center;"><strong>Total volume (mL)</strong></td>
					<td style="text-align: center;"><strong>[Na<sub>2</sub>S<sub>2</sub>O<sub>3</sub>] in mixture </strong></td>
					<td style="text-align: center;"><strong>Time (s)</strong></td>
					<td style="text-align: center;"><strong>Inverse time (s<sup>-1</sup>)</strong></td>
				</tr>
				<tr>
					<td style="text-align: center;">1, 2</td>
					<td style="text-align: center;">0.1</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td style="text-align: center;">3</td>
					<td style="text-align: center;">0.2</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td style="text-align: center;">4</td>
					<td style="text-align: center;">0.15</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td style="text-align: center;">5</td>
					<td style="text-align: center;">0.05</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
			</tbody>
		</table>
		<a href="https://www.jove.com/CDNSource/lm/labs/50/Table_4._Estimating_reaction_order_sodium_thiosulfate.xlsx" target="_blank">Click Here to download Table 4</a></div>
		</li>
		<li data-sub-note-item="2-2">
		<div class="lab-step">Remember that each trial used the same volume, so it took the same amount of sulfur to make each solution opaque. Thus, the reaction is twice as fast when the sodium thiosulfate concentration is doubled.</div>
		</li>
		<li data-sub-note-item="2-3">
		<div class="lab-step">When the concentration increases by a factor of 4, the reaction rate also increases by a factor of 4. This one-to-one relationship suggests that the reaction was first order for sodium thiosulfate. If so, we would expect a plot of rate versus sodium thiosulfate concentration to be linear.</div>
		</li>
		<li data-sub-note-item="2-4">
		<div class="lab-step">We didn&#39;t measure how much sulfur it took to turn the solution opaque, but we assume that it was the same for each trial. This lets us use the inverse of time to opacity as a stand-in for the rate. The slope won&#39;t be equal to k, but that&#39;s OK for this lab.</div>
		</li>
		<li data-sub-note-item="2-5">
		<div class="lab-step">Plot inverse reaction time as a function of sodium thiosulfate concentration. This confirms the linear relationship between sulfur production and thiosulfate concentration. Thus, the reaction seems to be first order with respect to thiosulfate.</div>
		</li>
		<li data-sub-note-item="2-6">
		<div class="lab-step">Use the same method to estimate the reaction order of HCl. First, calculate the starting concentration of HCl in the benchmark trials and in trials 6 through 8.
		<h4><strong>Table 5: Estimating reaction order HCl</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>[HCl] added</strong></td>
					<td style="text-align: center;"><strong>HCl volume (mL) </strong></td>
					<td style="text-align: center;"><strong>Total volume (mL) </strong></td>
					<td style="text-align: center;"><strong>[HCl] in mixture </strong></td>
					<td style="text-align: center;"><strong>Time (s)</strong></td>
				</tr>
				<tr>
					<td style="text-align: center;">1, 2</td>
					<td style="text-align: center;">0.1</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td style="text-align: center;">3</td>
					<td style="text-align: center;">0.2</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td style="text-align: center;">4</td>
					<td style="text-align: center;">0.15</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
				<tr>
					<td style="text-align: center;">5</td>
					<td style="text-align: center;">0.05</td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
					<td style="text-align: center;"></td>
				</tr>
			</tbody>
		</table>
		<a href="https://www.jove.com/CDNSource/lm/labs/50/Table_5._Estimating_reaction_order_HCl.xlsx" target="_blank">Click Here to download Table 5</a></div>
		</li>
		<li data-sub-note-item="2-7">
		<div class="lab-step">Then, look at the concentration of HCl and the reaction times for these trials. The reaction times are nearly identical, so the HCl concentration has no effect on the reaction rate. This means that HCl seems to be zeroth order for this reaction.</div>
		</li>
		<li data-sub-note-item="2-8">
		<div class="lab-step">Lastly, add up the reaction orders of the reactants to get the overall reaction order.</div>
		</li>
	</ul>
	</li>
</ol>

Concentration Dependence and Reaction Order - Procedure

Zależność od koncentracji

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

	Concentration DependenceExpand
	In this experiment, you will combine varying concentrations of ...

Recrystallization

<p>Source: Lara Al Hariri and Ahmed Basabrain at the University of Massachusetts Amherst, MA, USA</p>
<ol class="note-items">
	<li data-note-item="1" data-parent-collapse="">Recrystallization of Acetanilide<button class="expand">Expand</button>
	<p>In this part of the lab, you&#39;ll purify acetanilide by recrystallization from an aqueous solution. Acetanilide is moderately soluble in boiling water, but it&#39;s much less soluble in room temperature and cold water. Aniline, a potential impurity, is much more soluble in room temperature water. Thus, you&#39;ll first make a saturated solution of acetanilide in boiling water. Then, you&#39;ll slowly cool the solution to room temperature.</p>
	<p>As the acetanilide&#39;s solubility decreases, it will slowly form flat colorless or white crystals. Water-soluble impurities will remain in solution. You will then cool the solution to further decrease the solubility of acetanilide. Finally, you&#39;ll recover the crystals with vacuum filtration and measure the melting point of your crystals.</p>
	<ul class="lab-items">
		<li data-sub-note-item="1-3">
		<div class="lab-step">Before you start the lab, put on a lab coat, safety glasses, and nitrile gloves.</div>
		</li>
		<li data-sub-note-item="1-4">
		<div class="lab-step">To begin, bring a 125-mL Erlenmeyer flask and your lab notebook to the analytical balance to obtain your acetanilide.</div>
		</li>
		<li data-sub-note-item="1-5">
		<div class="lab-step">Measure 2 g of acetanilide in a tared weighing boat and record the precise mass in your lab notebook. Add the acetanilide to the flask and bring it back to your fume hood.
		<h4><strong>Table 1: Recrystallization of acetanilide</strong></h4>
		<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
			<tbody>
				<tr>
					<td><strong>Compound</strong></td>
					<td><strong>Mass (g) </strong></td>
					<td><strong>Melting point (&#176;C) </strong></td>
				</tr>
				<tr>
					<td>Acetanilide</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td>Seed crystal (if used)</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td>Recrystallized acetanilide<br />
					(<em>subtract seed crystal if used</em>)</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td></td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td><strong>Percent recovery</strong></td>
					<td></td>
					<td></td>
				</tr>
			</tbody>
		</table>
		<a href="https://www.jove.com/CDNSource/lm/labs/58/Table_1._Recrystallization_of_acetanilide.xlsx" target="_blank">Click Here to download Table 1</a></div>
		</li>
		<li data-sub-note-item="1-6">
		<div class="lab-step">Then, measure 30 mL of deionized water and carefully pour it into the flask of acetanilide.</div>
		</li>
		<li data-sub-note-item="1-7">
		<div class="lab-step">Place the flask on a stirring hotplate, add a stir bar, and set the hotplate to medium heat and stirring. Let the mixture boil for 4 &#8211; 5 min to dissolve the acetanilide.</div>
		</li>
		<li data-sub-note-item="1-8">
		<div class="lab-step">If you see solid acetanilide after the solution has boiled for 5 min, measure another 10 mL of deionized water. Use a Pasteur pipette to add water dropwise until either the solid dissolves or you&#39;ve added all 10 mL.</div>
		</li>
		<li data-sub-note-item="1-9">
		<div class="lab-step">Let the solution boil for at least a minute after each milliliter before adding more water.<strong> Note:</strong> It&#39;s important to add water slowly so that you don&#39;t cool the solution too much or add more water than you need.</div>
		</li>
		<li data-sub-note-item="1-10">
		<div class="lab-step">If you still see some solid after that, those are impurities that you must remove by hot filtration. If you do not see solid, continue the lab at step 14.</div>
		</li>
		<li data-sub-note-item="1-11">
		<div class="lab-step">To perform hot filtration, place about 15 mL of deionized water and one or two boiling chips in a 50-mL beaker and start heating it on a hotplate.</div>
		</li>
		<li data-sub-note-item="1-12">
		<div class="lab-step">Clamp another 125-mL Erlenmeyer flask on a hotplate and place a wide stem or stemless glass funnel in the flask. Fold a large piece of circular filter paper in quarters, place it point down in the funnel, and open it into a cone.</div>
		</li>
		<li data-sub-note-item="1-13">
		<div class="lab-step">Once the water boils, use a pipette to wet the filter paper with hot water. Then, use tongs to pick up the flask of acetanilide solution and pour the hot liquid into the funnel, being careful not to let it overflow. <strong>Note:</strong> If the stir bar falls into the funnel, retrieve it with tweezers and rinse it with hot water.</div>
		</li>
		<li data-sub-note-item="1-14">
		<div class="lab-step">Then, rinse the inside of the first flask with a few milliliters of hot water, pour the contents into the funnel, and dissolve any crystals that form with hot water.</div>
		</li>
		<li data-sub-note-item="1-15">
		<div class="lab-step">Once the funnel is empty, use tongs to move it to the first flask. Carefully unclamp the second flask. You can now continue with the lab as usual.</div>
		</li>
		<li data-sub-note-item="1-16">
		<div class="lab-step">Once there is no visible solid in your hot acetanilide solution, turn off the heat and stir motor and use tongs to place the flask on the floor of the hood.</div>
		</li>
		<li data-sub-note-item="1-17">
		<div class="lab-step">Leave the solution undisturbed as it cools to room temperature, which usually takes at least 15 min. Moving or shaking the flask will make it harder for crystals to form.</div>
		</li>
		<li data-sub-note-item="1-18">
		<div class="lab-step">While you wait, place a small amount of the starting acetanilide material in a weighing boat and bring it to the melting point apparatus.</div>
		</li>
		<li data-sub-note-item="1-19">
		<div class="lab-step">Pack a capillary tube with the acetanilide and measure and record its melting point.</div>
		</li>
		<li data-sub-note-item="1-20">
		<div class="lab-step">Once the solution cools, look for crystals without moving the flask too much. If you don&#39;t see any, use a glass rod to gently scratch the inside of the flask under the solution. It&#39;s easier for crystals to start to form on scratched or rough surfaces.</div>
		</li>
		<li data-sub-note-item="1-21">
		<div class="lab-step">If crystals still haven&#39;t formed after 10 more min, ask your instructor for a seed crystal of high purity acetanilide. Measure and record the mass of the seed crystal, and then drop it into the room temperature acetanilide solution. Crystal growth will then occur on the seed crystal.</div>
		</li>
		<li data-sub-note-item="1-22">
		<div class="lab-step">Once you see several crystals in the flask, prepare two ice baths in 600-mL beakers. One ice bath should nearly fill the beaker, and the other should occupy about half of the beaker.</div>
		</li>
		<li data-sub-note-item="1-23">
		<div class="lab-step">Measure 5 mL of deionized water and place it in the full ice bath. Confirm that the flask is cool to the touch and that several crystals have formed before gently placing it in the half-full ice bath. Leave the flask undisturbed to improve crystal growth.</div>
		</li>
		<li data-sub-note-item="1-24">
		<div class="lab-step">Check the crystal growth progress every 15 min. Once it looks like most of your acetanilide has crystallized, or you don&#39;t see much crystal growth over a 15-min period, set up for a vacuum filtration using a 125- or 250-mL filter flask.</div>
		</li>
		<li data-sub-note-item="1-25">
		<div class="lab-step">Wet the filter paper with 1 mL of ice-cold water. Then, start the vacuum and pour the contents of the flask into the B&#252;chner funnel of the vacuum filtration setup.</div>
		</li>
		<li data-sub-note-item="1-26">
		<div class="lab-step">Scrape crystals from the flask into the funnel using a glass rod or flexible spatula and rinse the remaining crystals into the funnel with about 1 mL of cold deionized water.</div>
		</li>
		<li data-sub-note-item="1-27">
		<div class="lab-step">Then, stop the vacuum, break the seal, and pour the rest of the cold deionized water into the funnel. Gently stir the crystals with the glass rod for a few seconds to wash them, being careful not to tear the filter paper.</div>
		</li>
		<li data-sub-note-item="1-28">
		<div class="lab-step">Turn on the vacuum to drain the funnel. Then, stop the vacuum, break the seal, and empty the filtrate into a waste beaker. Turn the vacuum back on, and leave the washed crystals in the funnel while the vacuum pump pulls air through them until they are dry, which usually takes about half an hour. You&#39;ll start working on your next recrystallization while you wait.</div>
		</li>
	</ul>
	</li>
	<li data-note-item="2" data-parent-collapse="">Recrystallization of <em>trans</em>-Cinnamic Acid<button class="expand">Expand</button>
	<p>In the second half of the lab, you&#39;ll recrystallize <em>trans</em>-cinnamic acid, which we&#39;ll just call cinnamic acid. Cinnamic acid is soluble in ethanol and nearly insoluble in water. So, you&#39;ll use a mixture of 95% ethanol and 5% water so that cinnamic acid is still soluble at high temperatures but less soluble at low temperatures.</p>
	<ul class="lab-items">
		<li data-sub-note-item="2-2">
		<div class="lab-step">To get started, fill a 5- or 10-mL graduated cylinder with deionized water and bring it to your hood.</div>
		</li>
		<li data-sub-note-item="2-3">
		<div class="lab-step">Then, weigh 0.5 g of cinnamic acid, record its mass, and place it in a 25-mL Erlenmeyer flask.
		<h4><strong>Table 2: Recrystallization of<em> trans</em>-Cinnamic acid</strong></h4>
		<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
			<tbody>
				<tr>
					<td><strong>Compound</strong></td>
					<td><strong>Mass (g) </strong></td>
					<td><strong>Melting point (&#176;C) </strong></td>
				</tr>
				<tr>
					<td><em>trans</em>-Cinnamic acid</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td>Seed crystal (if used)</td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td>Recrystallized<em> trans</em>-cinnamic acid<br />
					<em>(subtract seed crystal if used)</em></td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td></td>
					<td></td>
					<td></td>
				</tr>
				<tr>
					<td><strong>Percent recovery</strong></td>
					<td></td>
					<td></td>
				</tr>
			</tbody>
		</table>
		<a href="https://www.jove.com/CDNSource/lm/labs/58/Table_2._Recrystallization_of_trans-Cinnamic_acid.xlsx" target="_blank">Click Here to download Table 2</a></div>
		</li>
		<li data-sub-note-item="2-4">
		<div class="lab-step">Next, obtain 4 mL of 95% ethanol, and pour it into the flask containing cinnamic acid. Place the flask on a hotplate set to medium heat and stir the solution with a glass rod until the cinnamic acid dissolves, which usually takes 7 &#8211; 10 min.</div>
		</li>
		<li data-sub-note-item="2-5">
		<div class="lab-step">Then, turn off the heat and use a Pasteur pipette to add 5 drops of deionized water to the solution. This reduces cinnamic acid solubility. <strong>Note:</strong> If you don&#39;t see precipitation, add up to 5 more drops of water one by one. Stop adding water if you see solids starting to form.</div>
		</li>
		<li data-sub-note-item="2-6">
		<div class="lab-step">Then, remove the flask from the hotplate and let it cool to room temperature undisturbed.</div>
		</li>
		<li data-sub-note-item="2-7">
		<div class="lab-step">While you wait, get a small amount of the starting cinnamic acid and measure its melting point.</div>
		</li>
		<li data-sub-note-item="2-8">
		<div class="lab-step">Once the flask has cooled to room temperature, inspect it for crystals. If you don&#39;t see any, gently scratch the inside of the flask under the solution with the glass rod and wait for 10 min. <strong>Note:</strong> If that still doesn&#39;t work, get a seed crystal of cinnamic acid from your instructor. Remember to weigh the seed crystal and record its mass before you add it to your solution.</div>
		</li>
		<li data-sub-note-item="2-9">
		<div class="lab-step">Once the flask is cool to the touch and contains several crystals, remake the ice baths with one ice bath shallow enough that the 25-mL flask will not be completely submerged.</div>
		</li>
		<li data-sub-note-item="2-10">
		<div class="lab-step">Refill the graduated cylinder with deionized water. Place the flask of cinnamic acid crystals in the shallow ice bath and the water-filled graduated cylinder in the other ice bath. Let the crystals grow for at least 15 min.</div>
		</li>
		<li data-sub-note-item="2-11">
		<div class="lab-step">Your acetanilide crystals should be dry by now, so measure their mass and melting point while you wait for your cinnamic acid to finish crystallizing. Turn off the vacuum and break the vacuum seal.</div>
		</li>
		<li data-sub-note-item="2-12">
		<div class="lab-step">Gently place the B&#252;chner funnel in a clean beaker and bring it to the balance. Measure and record the mass of your pure acetanilide crystals. Remember to subtract the mass of the seed crystal if you used one.</div>
		</li>
		<li data-sub-note-item="2-13">
		<div class="lab-step">Then, bring your crystals to the melting point apparatus and measure and record the melting point.</div>
		</li>
		<li data-sub-note-item="2-14">
		<div class="lab-step">Now, place a clean, dry B&#252;chner funnel with a new piece of filter paper in the filter flask and check on your cinnamic acid. Once it looks like most of the cinnamic acid has crystallized, wet the filter paper with cold deionized water.</div>
		</li>
		<li data-sub-note-item="2-15">
		<div class="lab-step">Start the vacuum and pour the contents of the cinnamic acid flask into the funnel. Scrape the remaining crystals out of the flask with the glass rod and rinse the flask with about 1 mL of cold water.</div>
		</li>
		<li data-sub-note-item="2-16">
		<div class="lab-step">Now, turn off the vacuum, break the seal, and pour the rest of the cold water into the funnel. Gently stir the crystals with the glass rod to wash them, being careful not to damage the filter paper.</div>
		</li>
		<li data-sub-note-item="2-17">
		<div class="lab-step">Turn on the vacuum to drain the funnel. Leave the vacuum running until the cinnamic crystals are dry.</div>
		</li>
		<li data-sub-note-item="2-18">
		<div class="lab-step">Place the B&#252;chner funnel in a beaker and bring it to the balance. Measure and record the mass of your pure cinnamic acid crystals. Remember to subtract the mass of the seed crystal if you used one. Then, measure the melting point of your recrystallized cinnamic acid.</div>
		</li>
		<li data-sub-note-item="2-19">
		<div class="lab-step">Now, dispose of your acetanilide and cinnamic acid in the designated solid waste container. Flush the filtrate down the drain with running water and pour leftover ice and water into the sink.</div>
		</li>
		<li data-sub-note-item="2-20">
		<div class="lab-step">Clean your glassware using your usual methods and put away your other lab equipment.</div>
		</li>
		<li data-sub-note-item="2-21">
		<div class="lab-step">Lastly, discard your used pipettes in the glass waste and throw out used filter paper and other trash in the lab trash container.</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">First, calculate the percent recovery for each compound. We expect that some of each compound stayed in solution, so these percentages will be below the reported purity of your starting material.</div>
		</li>
		<li data-sub-note-item="3-2">
		<div class="lab-step">Now, look at the differences in melting points between the starting material and the crystals. For each compound, the less pure starting material has a lower melting point and a broader range than the recrystallized compound.</div>
		</li>
		<li data-sub-note-item="3-3">
		<div class="lab-step">Lastly, compare your results to the literature values. The melting point of acetanilide is 114 &#176;C, and the melting point of <em>trans</em>-cinnamic acid is 133 &#176;C. The melting points of your crystals should span 1 &#8211; 2 &#176;C and should be at or just below those values.</div>
		</li>
	</ul>
	</li>
</ol>

Recrystallization of Acetanilide and trans-Cinnamic Acid - Procedure

Rekrystalizacja

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

	Recrystallization of AcetanilideExpand
	In this part of ...

Thin-Layer Chromatography

<p>Source: Lara Al Hariri and&#160;Ahmed Basabrain at the University of Massachusetts Amherst, MA, USA</p>
<ol class="note-items">
	<li data-note-item="1" data-parent-collapse="">Effects of Mobile Phase Composition on Compound Separation<button class="expand">Expand</button>
	<p>In this section of the lab, you will use TLC to compare how well pure hexane and a mixture of 60% hexane and 40% ethyl acetate can separate phenol, benzoic acid, and butyl phenyl ether by measuring the distance the compound traveled in each solvent.</p>
	<ul class="lab-items">
		<li data-sub-note-item="1-2">
		<div class="lab-step">Before starting the lab, put on a lab coat, safety glasses, and nitrile gloves. Note: Hexane and ethyl acetate are both volatile, toxic, and flammable, so always work with them in a fume hood. Change your gloves if you get solvent on them, as hexane and ethyl acetate will permeate nitrile gloves within minutes.</div>
		</li>
		<li data-sub-note-item="1-3">
		<div class="lab-step">Now, obtain a paper towel and lay it on the benchtop. Carefully pick up a TLC plate using tweezers at the very end of the plate.<strong> Note: </strong>Always hold TLC plates by the edges or at the very top with tweezers to avoid damaging or contaminating the silica gel.</div>
		</li>
		<li data-sub-note-item="1-4">
		<div class="lab-step">Look for chips or scratches in the silica gel and get a new plate if necessary. It&#39;s fine if there is a little chipping at only one end, since that can be the top.</div>
		</li>
		<li data-sub-note-item="1-5">
		<div class="lab-step">Place the plate on the paper towel in your fume hood, and lay a metric ruler next to it. Use a graphite pencil to gently make a small mark on the silica gel ~1 cm from the top of the plate. Write &#8216;100&#8217; on the plate above the mark.</div>
		</li>
		<li data-sub-note-item="1-6">
		<div class="lab-step">Next, gently mark the plate 1 cm above the bottom on both sides. Carefully trace a straight line between the bottom marks to make the starting line.</div>
		</li>
		<li data-sub-note-item="1-7">
		<div class="lab-step">Then, add 3 evenly spaced small marks to the starting line with the outer marks at least 0.5 cm from the sides of the plate. <strong>Note:</strong> It&#39;s important for the silica gel to be undamaged here, so if you scratch it with the pencil, prepare a new plate.</div>
		</li>
		<li data-sub-note-item="1-8">
		<div class="lab-step">Now, trace the outline of the TLC plate in your lab notebook and precisely copy the marks on the plate. Label the three marks as &#8216;phenol&#8217;, &#8216;benzoic acid&#8217;, and &#8216;butyl phenyl ether&#8217;. You may also want to label the marks on the plate itself.</div>
		</li>
		<li data-sub-note-item="1-9">
		<div class="lab-step">Prepare a second TLC plate labeled &#8216;60/40&#8217; in the same way. Now, bring the labeled plates to the common bench and locate the 1 mg/mL solutions of phenol, benzoic acid, and butyl phenyl ether in ethanol.</div>
		</li>
		<li data-sub-note-item="1-10">
		<div class="lab-step">Dip a clean capillary into one of the three solutions. Briefly and lightly dab the capillary squarely against the intersection between the starting line and the mark for that compound. You should spot twice for each compound.</div>
		</li>
		<li data-sub-note-item="1-11">
		<div class="lab-step">Use the same capillary to spot the same solution on your second TLC plate. The spots should be no more than 2 mm in diameter. Then, discard the capillary and close the vial.</div>
		</li>
		<li data-sub-note-item="1-12">
		<div class="lab-step">Spot the other two solutions on the plates, in the same way, using clean capillaries. Be careful not to cross-contaminate the solutions. <strong>Note</strong>: If a spot is very light or very small, wait for it to dry, and then spot the same solution on top of the dry spot. If the spot is still wet, capillary action could make it too wide and dilute.</div>
		</li>
		<li data-sub-note-item="1-13">
		<div class="lab-step">Once you have made three spots on each plate, discard the capillaries and bring the plates back to your hood. Keep the plates lying flat while they dry.</div>
		</li>
		<li data-sub-note-item="1-14">
		<div class="lab-step">Now, label a 50-mL jar &#8216;100% hexane&#8217; and bring the jar, its cap, and a 10-mL graduated cylinder to the solvent hood.</div>
		</li>
		<li data-sub-note-item="1-15">
		<div class="lab-step">Measure 2 &#8211; 3 mL of pure hexane with your graduated cylinder and pour it into the jar to fill it to a depth of 0.62 &#8211; 0.7 cm. Cap the bottle and jar and return to your fume hood.</div>
		</li>
		<li data-sub-note-item="1-16">
		<div class="lab-step">Label a second 50-mL jar &#8216;60% hexane, 40% ethyl acetate&#8217; and bring it and a clean graduated cylinder to the solvent hood. Measure 2 &#8211; 3 mL of the 60/40 mixture into the jar and return to your fume hood.</div>
		</li>
		<li data-sub-note-item="1-17">
		<div class="lab-step">Stand the plates next to the jars to compare the relative position of the solvent and the spots. The solvent level should be below the spots for each plate.</div>
		</li>
		<li data-sub-note-item="1-18">
		<div class="lab-step">Then, obtain three pieces of filter paper, put a filter paper upright against the wall in each jar, and then cap the jars tightly.</div>
		</li>
		<li data-sub-note-item="1-19">
		<div class="lab-step">Initial each plate above the mark so you can identify them later. Confirm that the filter papers in the jars have become saturated with solvent and that the spots on the plates are completely dry.</div>
		</li>
		<li data-sub-note-item="1-20">
		<div class="lab-step">Now, open the 100% hexane jar and use tweezers to gently place the 100 plate in the jar, keeping the plate level with the solvent surface as it enters the solvent. Close the jar tightly, being careful not to jostle the plate.</div>
		</li>
		<li data-sub-note-item="1-21">
		<div class="lab-step">Leave the jar still as the solvent travels up the plate. Development in pure hexane usually takes about 7 &#8211; 10 min. Keep an eye on the plate to make sure that the solvent doesn&#39;t reach the top.</div>
		</li>
		<li data-sub-note-item="1-22">
		<div class="lab-step">When the pure hexane solvent front is about 1 cm from the top of the plate, retrieve it using tweezers. Lay it on the paper towel with the silica gel facing up and quickly mark the solvent front with a line and pencil across the plate. Leave the plate lying flat as the solvent evaporates.</div>
		</li>
		<li data-sub-note-item="1-23">
		<div class="lab-step">Place the other plate in the 60/40 eluent in the same way. Wait until the solvent front of the 60/40 plate is about 1 cm from the top, and then retrieve it and quickly mark the solvent front in the same way.</div>
		</li>
		<li data-sub-note-item="1-24">
		<div class="lab-step">Once the 100% hexane plate is completely dry, pick it up with tweezers and bring it and a pencil to the UV lamp on the common bench.</div>
		</li>
		<li data-sub-note-item="1-25">
		<div class="lab-step">Place the plate under the lamp and illuminate it with UV light. The plate will glow green because of a fluorescent marker in the silica gel layer. The dark spots are the compounds.</div>
		</li>
		<li data-sub-note-item="1-26">
		<div class="lab-step">Carefully outline every spot in pencil, even if they haven&#39;t moved very far.</div>
		</li>
		<li data-sub-note-item="1-27">
		<div class="lab-step">Then, turn off the lamp and bring your plate to the iodine development hood. Open the iodine chamber. Use tweezers to gently place the plate upright in the iodine chamber, and then cover the chamber again.</div>
		</li>
		<li data-sub-note-item="1-28">
		<div class="lab-step">Wait 3 &#8211; 4 min for the iodine vapor to stay in the spots. Then, retrieve the plate with tweezers, close the chamber, and outline the brown and yellow spots on the plate. These spots will fade, so mark them quickly.</div>
		</li>
		<li data-sub-note-item="1-29">
		<div class="lab-step">Visualize the dry 60/40 plate under UV light and then with iodine vapor in the same way.<strong> Note: </strong>To avoid side reactions, always check plates under UV light before exposing them to iodine.</div>
		</li>
		<li data-sub-note-item="1-30">
		<div class="lab-step">Now, precisely copy the spots and the solvent fronts onto the corresponding drawings of the plates in your lab notebook. For each plate, measure and record the height of the solvent front relative to the starting line.</div>
		</li>
		<li data-sub-note-item="1-31">
		<div class="lab-step">Lastly, measure the distance between the center of each spot and its starting line, and record it in your lab notebook. You can now move on to the next part of the lab.</div>
		</li>
	</ul>
	</li>
	<li data-note-item="2" data-parent-collapse="">Unknown Compound Identification<button class="expand">Expand</button>
	<p>In the second half of the lab, you&#39;ll use TLC to find out what&#39;s in an unknown analgesic. It could contain one or more of these compounds: aspirin, caffeine, and acetaminophen. These compounds are highly polar, so you&#39;ll use a premade mixture of 90% ethyl acetate and 10% hexane as the eluent.</p>
	<p>You&#39;ll run the three reference compounds and the unknown all on the same plate. Later, you&#39;ll use the Rf values of the reference compounds to identify the unknown. First, double-check which unknown material you were assigned to analyze before starting.</p>
	<ul class="lab-items">
		<li data-sub-note-item="2-3">
		<div class="lab-step">Now, lay a clean paper towel in the hood and obtain a new TLC plate. Prepare the top mark and the starting line the same way as before. Make four evenly spaced ticks on the plate, making sure that the outer marks are at least 0.5 cm away from the edges.</div>
		</li>
		<li data-sub-note-item="2-4">
		<div class="lab-step">Draw an exact copy of the plate in your lab notebook and label the tick marks as &#8216;aspirin&#8217;, &#8216;acetaminophen&#8217;, &#8216;caffeine&#8217;, and &#8216;unknown&#8217;.</div>
		</li>
		<li data-sub-note-item="2-5">
		<div class="lab-step">Now, bring your plate to the common bench. Spot solutions of aspirin, acetaminophen, caffeine, and the unknown that you were assigned on the appropriate marks using a clean capillary for each solution. <strong>Note: </strong>When you are spotting the unknown solution, don&#39;t dip the capillary too far into the solution vial to avoid clogging your capillary with insoluble material from the unknown substance.</div>
		</li>
		<li data-sub-note-item="2-6">
		<div class="lab-step">Discard the capillaries and close the vials before bringing the plate back to your fume hood. Place the plate on a paper towel to finish drying.</div>
		</li>
		<li data-sub-note-item="2-7">
		<div class="lab-step">Next, label a clean 50-mL screw-top jar &#8216;90% ethyl acetate, 10% hexane&#8217;, and bring it in a clean graduated cylinder to the solvent hood.</div>
		</li>
		<li data-sub-note-item="2-8">
		<div class="lab-step">Place 2 &#8211; 3 mL of the 90/10 solution in the jar and return to your fume hood. Confirm that the solvent level will be below the spots on the plate. Then, place a clean filter paper upright in the jar and close it tightly.</div>
		</li>
		<li data-sub-note-item="2-9">
		<div class="lab-step">Once the TLC plate is dry and the filter paper is saturated, carefully place the plate in the solvent. <strong>Note:</strong> It&#39;s essential to hold the plate level when you do this. If the solvent front is diagonal, the solvent distance traveled will be different for each spot, making it impossible to visually compare the unknown to the reference spots.</div>
		</li>
		<li data-sub-note-item="2-10">
		<div class="lab-step">Wait for the solvent front to reach 1 cm from the top of the plate. Then, retrieve the plate, mark it solvent front, and let it dry while lying flat.</div>
		</li>
		<li data-sub-note-item="2-11">
		<div class="lab-step">Once the plate is dry, visualize it with UV light and in the iodine chamber, and outline the spots in pencil.</div>
		</li>
		<li data-sub-note-item="2-12">
		<div class="lab-step">The unknown material could be a mixture of compounds, so look carefully for spots in line with all three reference spots.</div>
		</li>
		<li data-sub-note-item="2-13">
		<div class="lab-step">Then, precisely copy the spots onto the drawing of the plate in your lab notebook. Measure and record the height of the solvent front and the spots relative to the starting line.</div>
		</li>
		<li data-sub-note-item="2-14">
		<div class="lab-step">When you are finished, retrieve the solvent-saturated filter papers from the jars and let them dry in the hood.</div>
		</li>
		<li data-sub-note-item="2-15">
		<div class="lab-step">Pour leftover solvents into the standard organic waste, wash your glassware following your lab&#39;s usual methods, and put away your lab equipment.</div>
		</li>
		<li data-sub-note-item="2-16">
		<div class="lab-step">Throw out used TLC plates in the lab trash or with glass waste, depending on the backing material.</div>
		</li>
		<li data-sub-note-item="2-17">
		<div class="lab-step">Lastly, once the filters are dry, throw them out along with any remaining trash.</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">Calculate the R<sub>f</sub> values for phenol, benzoic acid, and butyl phenyl ether eluted with 100% hexane. For each compound on the TLC plate, measure the distance from the starting line to the center of its spot. Then, divide this value by the distance from the starting line to the solvent front.</div>
		</li>
		<li data-sub-note-item="3-2">
		<div class="lab-step">Only butyl phenyl ether moved a measurable distance from the starting line, which is consistent with it being the least polar. For effective separation, the R<sub>f</sub> values should differ by 0.3 &#8211; 0.7. Thus, we can infer that phenol and benzoic acid are too polar to be separated effectively by an apolar solvent like pure hexane.</div>
		</li>
		<li data-sub-note-item="3-3">
		<div class="lab-step">Calculate the R<sub>f</sub> values for the plate that you ran in a 60/40 mixture of hexane and ethyl acetate. Ethyl acetate is a moderately polar solvent, so the compounds traveled farther and are spaced farther apart on the plate.</div>
		</li>
		<li data-sub-note-item="3-4">
		<div class="lab-step">The separation order shows that benzoic acid, the most polar compound, had the highest affinity for silica gel, whereas butyl phenyl ether, the least polar compound, had the lowest affinity for silica gel. This is consistent with the assumption that less polar compounds travel farther on polar stationary phases.</div>
		</li>
		<li data-sub-note-item="3-5">
		<div class="lab-step">Look at your third plate and calculate the R<sub>f</sub> values for aspirin, acetaminophen, and caffeine. Caffeine, the most polar complex of the three, traveled the least. Acetaminophen traveled the second farthest, while aspirin, the least polar, traveled the most.</div>
		</li>
		<li data-sub-note-item="3-6">
		<div class="lab-step">Finally, identify the compounds in your unknown material by comparing its spot or spots to the three reference spots.</div>
		</li>
	</ul>
	</li>
</ol>

Thin-Layer Chromatography: Experimental Setup and Separation - Procedure

Chromatografia cienkowarstwowa

Source: Lara Al Hariri and&#160;Ahmed Basabrain at the University of Massachusetts Amherst, MA, USA

	Effects of Mobile Phase Composition on Compound ...

Buffers

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

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

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

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

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

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/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

Bacterial Transformation

<ol class="note-items"><li data-note-item="1" data-parent-collapse=""><strong>Bacterial Transformation</strong>
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="1-1"><div class="lab-step">IMPORTANT: In addition to wearing the appropriate personal protective equipment, be sure to take care to keep your face away from suspension cultures, and to avoid inhaling reagents. Do not touch your face while performing the experiment. Always wash your hands before and after every experiment.
</div></li><li data-sub-note-item="1-2"><div class="lab-step">When you are ready to safely begin, check to make sure you have a tube or tubes of competent <em>E. coli</em> cells in your ice bucket.
</div></li><li data-sub-note-item="1-3"><div class="lab-step">Then, transfer 50 &#181;L from one of these tubes to each of two new micro-centrifuge tubes, one labeled '&#8211;', and one labeled '+'.
</div></li><li data-sub-note-item="1-4"><div class="lab-step">Next, pipet 2 &#181;L of p-GREEN plasmid into the plus tube and flick the tube gently to incorporate the plasmid throughout the bacterial solution. HYPOTHESES: The experimental hypothesis is that on plates containing no antibiotic, bacteria with or without the plasmid can survive, and so both will grow lawns of bacteria. Conversely, on the plates containing the ampicillin antibiotic, only the bacteria successfully transformed with the resistance plasmid will grow, whereas the minus plasmid plates will contain no bacteria. The null hypothesis of this experiment is that there will be no difference in bacterial growth between the plates.
</div></li><li data-sub-note-item="1-5"><div class="lab-step">Incubate both tubes on ice.
</div></li><li data-sub-note-item="1-6"><div class="lab-step">After 30 minutes submerge both tubes 3/4 of the way into the water of a 42 &#176;C water bath for 30 seconds.
</div></li><li data-sub-note-item="1-7"><div class="lab-step">At the end of the water bath incubation, place the tubes back on ice for 5minutes, then add 950 &#956;L of room temperature SOC media to each tube.
</div></li><li data-sub-note-item="1-8"><div class="lab-step">Incubate the tubes in a shaker at 37 &#176;C for 60 minutes at 250 rpm.
</div></li><li data-sub-note-item="1-9"><div class="lab-step">While the tubes are incubating, label the bottoms of one LB and one LB/ampicillin, or AMP plate, &#8220;+ plasmid&#8221;, and the bottoms of one LB plate and one LB/AMP plate &#8220;- plasmid&#8221;.
</div></li><li data-sub-note-item="1-10"><div class="lab-step">At the end of the incubation, pipet 50 &#956;L from the &#8220;+ plasmid&#8221; cell tube onto both the LB/AMP plus plate and the LB plus plate.
</div></li><li data-sub-note-item="1-11"><div class="lab-step">Then, with a fresh tip, add 50 &#956;L of the bacteria that were not exposed to the plasmid to each of the minus plates.
</div></li><li data-sub-note-item="1-12"><div class="lab-step">Then, spread each plate using a fresh pipette tip.
</div></li><li data-sub-note-item="1-13"><div class="lab-step">Cover the plates with their lids, and then seal them with laboratory film.
</div></li><li data-sub-note-item="1-14"><div class="lab-step">Then store the plates upside down in a 37 &#176;C incubator for 24 hours.
</div></li></ul></li><li data-note-item="2" data-parent-collapse=""><strong>Results</strong>
<button class="expand">Expand</button><ul class="lab-items"><li data-sub-note-item="2-1"><div class="lab-step">The day after the transformation, observe which plates have bacteria, and how the bacteria are distributed over each plate.
</div></li><li data-sub-note-item="2-2"><div class="lab-step">Record the number of colonies and the color of the colonies for different conditions. <a href="https://www.jove.com/CDNSource/lm/labs/11/Table_1_Bacterial_Transformation.xlsx" style="display:block;margin:20px;" target="__blank">Click Here to download Table 1</a>
</div></li><li data-sub-note-item="2-3"><div class="lab-step">To determine the initial mass, IM, of the plasmid, first multiply the plasmid concentration from the stock tube of p-GREEN by the volume transferred into the experimental tube.
</div></li><li data-sub-note-item="2-4"><div class="lab-step">To calculate how much DNA is spread onto each plate, divide the volume of bacterial suspension spread onto each plate by the volume of the suspension in the tube, and multiply this fraction by the initial mass of plasmid used.
</div></li><li data-sub-note-item="2-5"><div class="lab-step">To determine the transformation efficiency, count all of the visibly distinct colonies on the plus plasmid LB/AMP plate, and divide this number by the mass of plasmid spread. NOTE: The unit for transformation efficiencies is cfu/&#956;g of plasmid DNA.
</div></li><li data-sub-note-item="2-6"><div class="lab-step">Record all of the results for each plate in the table and analyze them.
</div></li><li data-sub-note-item="2-7"><div class="lab-step">Observe any differences between the colonies and make hypotheses to explain this observation. Consider the factors that may affect the transformation efficiency of competent bacteria.
</div></li><li data-sub-note-item="2-8"><div class="lab-step">NOTE: You will have also noticed that each plate had a different outcome. The minus plasmid and plus plasmid LB plates should have grown a lot of bacteria across their plates. The minus plasmid LB/AMP plate should have no bacterial growth. And the plus plasmid LB/AMP plate should have grown bacterial colonies that fluoresced green.
</div></li></ul></li></ol>

Bacterial Transformation using Plasmids - Procedure

Transformacja bakteryjna

Bacterial Transformation
ExpandIMPORTANT: In addition to wearing the appropriate personal protective equipment, be sure to take care to keep your face ...

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/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/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 ...