Lab Report on Bacteria and Archaea - that it is possible to identify bacterial species based on their cellular morphology. 3 Labs performed which is to be used for this lab report Lab 1 - Microscopy

Agar plates are not reusable. When you are finished with an agar plate, please place itin one of the autoclave bags at the front of the lab. Once you have put away all of your bacterial samples, spray your bench with disinfectant and then wipe itwith apaper towel. This will clean up any tiny spills that you may not have noticed during the lab. Itisalso agood habit to clean your bench this way at the start of the lab, in case the students in a previous lab section did not clean the bench as thoroughly as they should have. Once you have cleaned up everything else, remove and dispose of your gloves, take off your lab coat and put itin alarge plastic bag with your name and seat number on it. Try to press most of the air out of the bag so that we can fit all of the lab coats in plastic bags into one large bin per lab section. The last thing you should do before you leave the room isto wash your hands again. BIOL2902L Fall 2020 6 Lab 1: Microscopy I This lab will be delivered in-person. A pre-recorded video outlining the experimental procedures will be shown at the beginning of the lab. Please read through this portion of the lab manual before attending lab.

Part 1 Microscopes and magnification Microscopes give us the ability to view individual bacterial cells. Most bacterial cells are 0.5 to 5 micrometres (µm) in length, about one-tenth of the size of aeukaryotic cell. In alight microscope, the process of magnification uses lenses to increase the apparent size of the objects being viewed. The clarity or sharpness of the image isdescribed as resolution. Ifan image islow-resolution, itcan appear blurry and fine details will not be visible. Most cheap microscopes offer high magnification but low resolving power, giving you avery magnified view of hazy blobs. The magnifying power of amicroscope comes from the combination of two sets of lenses. The eyepieces you look through are called the ocular lenses and provide aconstant 10X magnification. The objective lenses are set into arotating nosepiece and include 4X, 10X, 40X and 100X magnifications. The magnification of the two sets of lenses work together so, ifyou are using the 10X ocular lenses and the 4X objective lens, your total magnification will be 40X. As this isamicrobiology class, we can always skip the 4X objective lens because you will not see any bacterial cells with only 40X magnification. The magnification provided by microscopes has standardized over time. As mentioned, low-power (40X total magnification) isnot useful in our labs. Medium power is100X magnification and high power is400X. In order to magnify 1000X, microscope immersion oil isused in order to decrease the refraction that occurs when light passes between glass and air. The oil isamineral oil with the same optical density as glass; when the light passes from the glass slide into the oil and then into the 100X objective lens, itwill not be refracted so the final image will be brighter and sharper than you can achieve without immersion oil. Immersion oil isonly used on the 100X lens (because the amount of refraction isnot significant on the less powerful lenses) and the 100X lens should always be used with immersion oil (because otherwise it’sjust abigger but darker and blurrier version of what you see with the 40X lens). If someone refers to oil immersion microscopy, they are talking about 1000X total magnification. Bacterial staining In non-scientific conversation, the term “stain ”isoften used to describe anything that changes the colour of an object. The scientific definition of astain isareagent that increases the contrast between two different things. For example, cells and water are both, generally speaking, clear. Ifyou were to apply ared chemical that made both the cells and the water red, you wouldn ’tfind itany easier to observe the cells. A more useful reagent would be one that turns the cells red but does not change the colour of the water, or vice versa. The increased contrast between the cells and the water makes it much easier to see them under amicroscope. Stains consist of asolvent, typically water or ethanol, and acoloured molecule called the chromogen. The chromogen iscomposed of acoloured component (the chromophore) and acharged component (the auxochrome). Basic (alkaline pH) stains have positively-charged auxochromes and so are attracted to the negatively-charged bacterial nucleic acids and cell walls. In today ’slab, we will be using simple stains, which are stains in which asingle basic stain, such as crystal violet, isapplied. In order to keep the cells from being washed away when we rinse the stain off the glass slide, the cells must first be fixed to the glass. Historically, this was done by allowing the cells to air-dry onto the glass and then heating them, effectively baking the cells onto the slide. While this iseffective, itcan also cause noticeable distortions in the morphology of some bacterial species. An alternative approach isto cover air-dried cells with methanol for two minutes, which will also adhere the cell to the glass slide but typically without any detectable levels of distortion. Methanol-fixed cells also adhere to the slide better and give more reliable results for Gram staining (Lab 3). BIOL2902L Fall 2020 7 Lab equipment care You must clean your microscope thoroughly after each use. This includes removing used slides, wiping all external surfaces, aligning the mechanical stage, selecting the 4x objective lens or the empty slot between lenses and removing the oil from the oil immersion lens. Workflow Today ’ssimple staining experiment will be performed with alab partner (while social distancing) .Begin by each making six bacterial smears, two of each bacterial species you are provided with .Your partner will be making bacterial smears of the other three bacterial species. You will only need one slide of each species but making new bacterial smears istime-consuming because of the need to wait for the slides to air-dry prior to methanol-fixation. Itisagood idea for you and your lab partner to prepare two bacterial smears of each specimen so that ifone does not work, you will not need to repeat the entire procedure to view another slide. You need to make drawings of the three bacterial species that you observe .You will then take aphoto of your drawings and share this with your lab partner. By the end of the lab you should have drawings of all six bacterial species. A worksheet will be provided for you to record your drawings and accompanying commentary on cell shape and arrangement. When drawing, remember what you are trying to tell your reader in your lab report; do you need to draw every cell you can see or just afew that look representative of the slide? Try to show figures that support your written descriptions. Ifacell arrangement does not fit into acategory, describe itin more detail. Ifyou record something in greater detail now, you can choose not to use those details in your lab report. Ifyou don ’twrite itdown now, you can ’tget additional details later. Microscope calibration (Shown in pre-lab video) 1. Because we will be viewing microbes, start with the 10x objective lens. 2. Take aclean glass slide from the box of slides in your desk. Using agrease pencil, draw acircle or oval on one side of the slide; this will be the “bottom ”of the slide. On the other side of the slide, where you would normally put aloopful of bacteria, draw an Xso that itappears in the middle of the circle. This will be the “top ”of the slide, placed face-up when you put the slide on the microscope. Do not draw this Xwhen you are making slides with bacteria. 3. Place the slide on the stage and clip itinto place. 4. Adjust the ocular lenses so that you can see comfortably with both eyes. 5. Using the 10x objective lens, try to focus on the slide. 6. Start with the stage as close as possible to the lens, and use the focus to slowly move the stage away. Ifyou think you are focused on the slide, use the mechanical stage to move the slide abit. If what you are focused on does not move, you are not focused on the slide. 7. Because all of our slides are approximately the same distance from the lens, they should all be in focus at approximately the same setting. 8. You can rotate the objective lens to 4x and safely remove the slide without changing the focus. Care and maintenance of microscopes 1. Clean the oil immersion lens immediately after you are finished viewing that slide. Apply new oil for each new slide. 2. Double-check that you have removed your slide. 3. Clean all of the objective and ocular lenses with lens paper. 4. Store the microscope on the 4x objective lens or the empty slot between lenses. Bacterial cultures (on tryptic soy agar plates ) Note: Tryptic soy agar, trypticase soy agar and tryptone soy agar are all different names for the same type of growth media. Any of these terms isacceptable and will vary according to which company manufactured the product. TSA isageneral-purpose medium with enough nutrients to allow awide variety of bacteria to grow. BIOL2902L Fall 2020 8 • Bacillus cereus • Pseudomonas flourescens • Micrococcus luteus • Escherichia coli • Staphylococcus epidermidis • Flexibacter canadensis Materials • Microscope slides • Saline • Methanol • Crystal violet EXPERIMENT: Simple stains with crystal violet 1. Hold slides only by the edges to avoid getting smudges or grease on the slides. Label your slides using agrease pencil; the methanol that you will use to fix the slide in step 7will remove permanent marker from glass. Use alabel that will indicate whether the slide isupside-down or right-side-up (e.g. alabel that says “A”looks the same ifthe slide isflipped over, but “BC ”does not). Itisuseful to draw an oval on the underside of the slide, near the middle, to help you find your bacterial smear after it has dried. 2. To make asmear from asolid culture, like an agar plate, place one drop of saline onto the slide. Using asterile inoculating loop, transfer one small loopful of bacteria from the agar into the saline drop on the slide and use the loop to mix the culture into the saline. 3. Using the inoculating loop, spread the bacterial suspension into athin smear. 4. Allow the smear to air-dry. This takes approximately 20 minutes, depending on how much broth or saline you used to make the smear and how far you spread out the smear on the glass slide. Do not blow on the slide or wave itin the air to try to dry itfaster. The slides can be dried more quickly by resting them on awarm slide drying bench. A good smear looks like afaint whitish layer or film when dried. 5. Place the air-dried smear slide on your staining tray and use adropper bottle of methanol to cover the surface of the slide for two minutes. 6. Pour off the excess methanol into your staining tray. 7. Rinse the slide with water. 8. Using the dropped included with the bottle, cover the smear with crystal violet for one minute. 9. Wash the crystal violet off the slide using the bottle of water. 10. Blot the slide in your book of bibulous paper. 11. Focus on the slide at 400x magnification with the 40x objective lens. 12. Rotate the objective lenses back to 4x to make room to access the lens. Place asmall drop of oil directly on the slide. 1. Rotate the objective lenses the other direction, so that the 100x oil immersion lens isin place. The drop of oil should be touching the lens. Only use fine focus with oil immersion. Only use immersion oil with the oil immersion (100x) lens. 13. View at 1000x magnification with immersion oil and make adrawing of what you see. 14. After viewing each slide, use aKim-Wipe or microscope lens paper to clean the objective and ocular lenses and to remove any immersion oil from the microscope. Part 2 Bacterial cultures When amicrobiologist isasked to identify amicrobe, the microbe must typically be isolated first. Bacteria often form complex communities in nature, which can make itdifficult to identify individual species. A culture isan artificial setting, such as abroth or plate, that allows organisms to grow. A pure culture contains only asingle species whereas amixed culture contains more than one species. Cultures are grown on or in different kinds of media; amedium such as abroth or an agar plate contains the essential resources for growth. Because not all organisms require the same resources, different media are used for growing different organisms. Incubation conditions such as temperature and gases such as carbon dioxide and oxygen are also important for the growth of different species. BIOL2902L Fall 2020 9 Colony-forming units Colony-forming units (CFUs) are the units we use to quantify bacteria. Because bacteria can reproduce asexually, asingle living cell can give rise to an entire colony of bacteria. Ifbacteria are spread out across an agar plate, each living cell should grow into avisible colony. By counting the number of colonies, we can estimate the number of living bacterial cells that were put onto the agar plate. In making this calculation, we make several assumptions. One assumption isthat each colony arises from asingle living cell. Iftwo cells are very close together, they may appear to be part of asingle colony. We also assume that all living cells will produce colonies. Itispossible for acell to be alive but, for awide variety of reasons, itmay not grow into acolony that isvisible on the agar plate to be counted. Finally, we assume that all cells within acolony are genetically identical to each other and to the original single cell. Because the cells reproduce asexually, each of the first two daughter cells should be identical to the original parent cell, and so forth, until acolony of millions of cells isproduced. Therefore, each of the millions or billions of cells in asingle colony should be identical to all of the other cells in the colony. If you transfer some of these cells into new growth media, all of the transferred cells should be genetically identical to the single cell that originally gave rise to the colony. For our purposes, these are reasonable assumptions to make. Four-quadrant streak plates Streak plating isamethod of isolating organisms by producing individual colonies on an agar plate. The colony can then be isolated and transferred into sterile medium to grow as apure culture. The streak plate method takes abacterial sample and streaks itover an agar plate. The streaking motion results in individual cells being deposited separately, as the cell density reduces. Cells that are isolated will grow, forming pure colonies of multiple cells of asingle organism. The cells are not always deposited as individuals, sometimes there are pairs, chains or clusters of an organism that can grow into acolony; as such, the term colony-forming unit or CFU isused for an accurate descriptor of colony origin. The most common pattern of plate streaking iscalled the four-quadrant method, leaving different densities of cells across the media. Selective media can be used to help in the isolation process, as some media will encourage or inhibit growth of particular organisms. Some selective media can demonstrate differences between organisms, and are thus termed selective and differential. In today ’sexperiments, we will inoculate agar plates with bacteria using afour-quadrant streak plate technique. We will use these plates next week to examine the morphology of bacterial colonies. The streak plate inoculation procedure can separate cells of amixed culture so that discrete colonies can be isolated. This isaform of dilution in which the cells are spread out across aphysical space (the surface of the agar) until they are at alow enough concentration that individual cells can grow into distinct colonies. Bacterial cultures for streak plates (in tryptic soy broths ) • Bacillus cereus • Pseudomonas flourescens • Micrococcus luteus • Escherichia coli • Staphylococcus epidermidis • Flexibacter canadensis Materials (per student) • 3tryptic soy agar (TSA) plates EXPERIMENT: Streak plates for the isolation of pure cultures This procedure isillustrated in Figure 2. 1. Label all of your agar plates with the organisms to be inoculated, the date, your lab section and seat number. 2. Using the four-quadrant streak plate technique described on the following page, inoculate one TSA plate with one of your bacterial cultures. We use bench-top incinerators instead of the Bunsen burners shown in the illustrations. 3. Repeat step 3using your remaining two bacterial cultures. Between you and your lab partner should now have inoculated all six bacterial species. BIOL2902L Fall 2020 10 4. Write your lab section and seat number on apiece of masking tape and use itto label aplastic storage bucket. You and your lab partner will share one bucket. 5. Place the agar plates in the bucket so that the plates are agar-side up. 6. Place the buckets in the 35 °C incubator on the shelf marked for your lab section. CLEAN UP 1. Ensure that all of your agar plates are in alabeled plastic bucket in the incubator on the shelf marked for your lab section. 2. Put microscope slides and cover slips in yellow sharps buckets. 3. Remove any remaining oil from all of the objective lenses. 4. Rotate the objective lens to 4x or the empty space between lenses. 5. Store the microscope under the dust cover, under your bench. 6. Clean your bench with disinfectant and paper towel. 7. Remove your lab coat and put itin its plastic bag. 8. Remove your mask and put itin its plastic bag. 9. Wash your hands. BIOL2902L Fall 2020 11 1. Label the bottom of the plate with the bacterial species or specimen name, the date, your lab section and your seat number. 2. Ifthe bacterial mixture isin a broth culture, suspend the cells by gently swirling the tube. Do not hold the tube by the cap: it might come off. 3. Hold the inoculating loop as you would apencil. Sterilize the inoculating loop by inserting it into the bench-top incinerator on your bench for 5to 7 seconds. The wire does not need to be red-hot. Allow the loop to cool but do not let it touch anything. 4. Hold the culture tube in your other hand. Curl the little finger of the hand holding the loop around the cap and remove it by twisting and pulling with your other hand. Do not set the cap down. 5. Insert the cooled loop into the broth culture and withdraw a bead of culture held within the loop 6. Lift one side of the Petri plate lid just enough so that you can insert the loop and place the bead of culture on the far surface of the agar. You an rest the plate on the bench top. 7. Streak, or spread out, the bacteria in the bead of culture by moving the tip of the loop in aback- and-forth motion. Do this in the first quadrant (1), as illustrated below. Then sterilize the loop in the incinerator. After the loop cools, start the second quadrant by moving the loop tip through the last few streaks of the first quadrant. Repeat for the remaining two quadrants. When you have finished preparing the streak plate, sterilize the loop before you set itdown. Figure 2: Four-quadrant streak plates, adapted from Techniques in Microbiology: A Student ’sHandbook . Laboratory guidelines have since been updated to recommend benchtop incinerators rather than Bunsen burners. BIOL2902L Fall 2020 12 Lab 2: Microscopy II This lab will be delivered in-person. A pre-recorded video outlining the experimental procedures will be shown at the beginning of the lab. Please read through this portion of the lab manual before attending lab.

Fixing bacterial cells to aglass microscope slide kills the cells, making itimpossible to view changes to the cells over time, such as motility or cell division. Bacterial wet mounts are unstained preparations that can show motility and lacks the distortion common to slide fixation. Because these cells are unstained, they are less easy to view than stained cells. Negative stains Negative staining uses an acidic stain that will not penetrate the cells because of the negative charge on the surface of bacteria. The cells will remain clear, in contrast to the darker stain of the background. The cells will be dried onto the slide and will not show motility but, because the cells have not been fixed to the slide with methanol, they may be less distorted than ifthey had been heat-fixed or methanol- fixed. Some cellular morphologies, such as spirochetes, are particularly susceptible to distortion from fixation to the glass slide and are best viewed using negative stains. For most organisms, there isminimal ifany detectable distortion with methanol fixation. Because these cells have not been methanol-fixed, however, they can remain infectious and should be handled with care. Negative stains use adye solution containing an acidic chromogen that carries anegative charge. The negative charge of the bacteria ’ssurface repels the chromogen, resulting in the cells remaining unstained. The background of the slide, however, takes on the stain and provides the contrast to view the unstained cells. The acidic negative stain we will be using today isnigrosin. Wet mounts As staining and fixing result in the death of the microorganisms, wet mounts can be used to allow observation of the living cells. This enables one to determine motility, natural cell size, arrangement and shape, which can help in the identification process. Wet mount preparations involve putting the specimen in adrop of water on the microscope slide and putting on the cover slip. Itisbest to view with minimal illumination, as there isno stain to provide contrast and the cells are mostly transparent. Ifthe brightness istoo high, itwill not be possible to view the cells on the slide. Motility can be observed in low or high magnification, but must be done relatively quickly to prevent the preparation drying. Water will recede as the slide isbeing viewed: that motion should not be mistaken for motility of the cells: their motion will be independent darting. When observing live cells, itisimportant to note the difference between Brownian motion and true motility. Brownian motion results from cells interacting with water molecules and looks like vibration. Microscopic water currents can also make cells appear to flow or drift, particularly after the slide has been moved. In contrast, true motility appears as independent movement over adistance; cells can often be seen changing direction and speed. Be aware that some organisms that are capable of motility may not display that motility during the brief period that you are observing them. In some cases, cells may be adapting to their new environment or may lack stimuli to promote movement. Your results will be based on whether you observed motion during the time that you were looking at the cells; do your best to differentiate between intentional motility and random Brownian motion. Itcan be difficult to use immersion oil when viewing through acover slip because the cover slip will often adhere more strongly to the objective lens through the oil than to the glass slide through the drop of broth culture. Ifyou choose to use oil immersion with acover slip, be especially careful when focusing and moving the slide. BIOL2902L Fall 2020 13 Phase contrast microscopes use special optics to highlight cellular components that differ only slightly in refractive index from the surrounding water. This allows you to view unstained cells at very high contrast compared to the background, combining the benefits of awet mount with the benefits of staining. However, because of the specialized lenses and condensers involved, they can initially be more difficult to work with than our standard brightfield microscopes. In today ’slab, phase contrast microscopes will be set up as demonstrations. Workflow Today ’sexperiments are all performed with alab partner (while social distancing) .Begin by each making six negative stains, two for each of the bacterial species you are working with .You will only need one slide of each species but the failure rate for first-time negative stains ishigh enough that itis worthwhile to prepare aspare in case the first slide doesn ’twork. While you are waiting for the negative stains to dry, prepare and view wet mounts. Wet mounts become useless ifthey dry out, so you should make one slide, view it, and then proceed to the next species. Because there ismore contrast in negative stains, they are typically easier to focus than wet mounts. You need to make drawings of the three bacterial species that you observe .You will then take aphoto of your drawings and share this with your lab partner. By the end of the lab you should have drawings of all six bacterial species. A worksheet will be provided for you to record your drawings and accompanying commentary on cell shape and arrangement. When drawing, remember what you are trying to tell your reader in your lab report; do you need to draw every cell you can see or just afew that look representative of the slide? Try to show figures that support your written descriptions. Ifacell arrangement does not fit into acategory, describe itin more detail. Ifyou record something in greater detail now, you can choose not to use those details in your lab report. Ifyou don ’twrite itdown now, you can ’tget additional details later. Bacterial cultures (on tryptic soy agar plates) • Bacillus cereus • Pseudomonas flourescens • Micrococcus luteus • Escherichia coli • Staphylococcus epidermidis • Flexibacter canadensis Materials • Microscope slides • Nigrosin • Saline EXPERIMENT: Negative stains with nigrosine 1. Label your microscope slides. Because these slides are not fixed with methanol, you can use a grease pencil or apermanent marker. 2. Put asmall drop of nigrosin onto one end of the microscope slide. Nigrosin dries fairly quickly, so itis best to make negative stains one slide at atime. 3. Using aseptic technique, collect atiny amount of colony growth from an agar plate with your inoculating loop. 4. Mix the bacteria from your inoculating loop into the nigrosin. 5. Place aglass microscope slide or cover slip at a45 °angle into the drop of nigrosin and suspended bacteria and allow the drop to spread along the edge of the applied slide. 6. Push the drop away from the previously spread drop of suspended bacteria, forming athin smear. 7. Air-dry the slide. The slides can be dried more quickly by resting them on awarm slide drying bench. 8. View at 1000x magnifications and make aquick sketch of what you see. They can be viewed using immersion oil ifyou let the negative stain dry first. 9. Repeat for the three bacterial cultures you are working with. BIOL2902L Fall 2020 14 EXPERIMENT: Bacterial wet mounts 1. Label your microscope slides. Because these slides are not fixed with methanol, you can use a grease pencil or apermanent marker. 2. Put asmall drop of saline onto amicroscope slide. 3. Using aseptic technique, collect atiny amount of colony growth from an agar plate with your inoculating loop. 4. Mix the bacteria from your inoculating loop into the saline. 5. Gently place acover slip across the mixture. 6. View at 1000x magnifications and make aquick sketch of what you see. 7. Repeat for the three bacterial cultures you are working with. Figure 1: Negative staining technique with nigrosin, adapted from Cappuccino and Sherman ’s Microbiology: A Laboratory Manual . CLEAN UP 1. Put microscope slides and cover slips in yellow sharps buckets. 2. Remove any remaining oil from all of the objective lenses. 3. Rotate the objective lens to 4x or the ‘empty ’space. 4. Store the microscope under the dust cover, under your bench. 5. Clean your bench with disinfectant and paper towel. 6. Remove your lab coat and put itin its plastic bag. 7. Wash your hands. BIOL2902L Fall 2020 15 Lab 3: Microscopy III This lab will be delivered in-person. A pre-recorded video outlining the experimental procedures will be shown at the beginning of the lab. Please read through this portion of the lab manual before attending lab.

Differential stains A differential staining technique isone that distinguishes between cell types or their structures on the basis of astain that isretained in some structures and washed away in others. There are many differential stain techniques, such as endospore stains that detect endospores within bacterial cells or acid-fast stains that detect bacteria with waxy cuticles like Mycobacterium species. The most important differential stain in bacteriology isGram staining. Gram staining isnamed after its inventor, Hans Christian Gram, which iswhy we capitalize Gram staining but do not capitalize simple or negative staining.

Gram staining Gram staining isatool used to identify Gram-positive and Gram-negative organisms, and isone of the most significant and recurrent microbiological differential stains. Aside from demonstrating the organism ’sGram reaction, the stain also has the benefits of asimple stain in that itincreases contrast to more easily see cell morphology, size and arrangement. Its simplicity and accessibility makes itan effective tool for quick presumptive identification, either confirming or eliminating possible organisms through the results. The Gram stain uses two basic stains with adecolourization step in between. The primary stain iscrystal violet, and iodine isapplied as amordant to enhance the colour of the crystal violet, forming crystal violet-iodine complexes that make the stain more resilient to being washed away. Decolourization with ethanol removes the crystal violent from only the Gram-negative cells. Gram-positive cells have alower lipid-content and thicker peptidoglycan layer than Gram-negative cells. The decolouration stage strips away the lipids from the Gram-negative outer membrane, making itmore porous and incapable of holding the crystal violet-iodine complex. The thicker layer of peptidoglycan in Gram-positive cells traps the crystal violet-iodine complex effectively, resisting the decolourization. Gram-positive cells remain stained purple, and Gram-negative cells are decoloured and can pick up the counterstain, safranin. An effective Gram-staining procedure results in Gram-negative cells appearing reddish-pink and Gram- positive cells appearing purple. Decolourization isthe most crucial step in differential staining. Leaving the ethanol on for the wrong period of time can result in Gram-positive cells being decoloured (ethanol stays on too long) or Gram- negative cells not being decoloured enough (ethanol ison too briefly). The Gram reaction will remain the same regardless of the decolourization step, but the results will be flawed. IfaGram-positive cell is stained incorrectly and appears pink, this isreferred to as afalse-negative because the result appears to be Gram-negative but the organism isGram positive (this isafalse result). IfaGram-negative organism isstained purple, this isreferred to as afalse-positive because the organism falsely appears Gram-positive.

Gram stain consistency can also be impacted by the age of the culture, as older Gram-positive cultures can lose their resistance to decolourization, giving afalse-negative result. Bacillus and Staphylococcus cultures should be used within 24 hours to ensure best results. Non-bacterial elements can complicate staining, as old or incorrectly made stain can disrupt the field and crystal violet crystals can be mistaken for bacteria. Direct smears can show host-cells or non-cellular material. Workflow Today ’sexperiments are all performed with alab partner (while social distancing) .Begin by each making six bacterial smears, two of each bacterial species you are working with .Additionally, you will need to create two slides of one of the unknown mixes. You will only need one slide of each species /mix but Gram stains are notoriously difficult at first, so itisagood idea for you and your lab partner to BIOL2902L Fall 2020 16 prepare two bacterial smears of each specimen so that ifone does not work, you will not need to repeat the entire procedure to view another slide. These extra slides can also be used to confirm any ambiguous results that you see on your first slides. You need to make drawings of the three bacterial species that you observe .You will then take aphoto of your drawings and share this with your lab partner. By the end of the lab you should have drawings of all six bacterial species. A worksheet will be provided for you to record your drawings and accompanying commentary on cell shape and arrangement. At this point, you should be fairly familiar with the morphology and arrangement of the six bacterial species. Be aware that the colour of the stained glass slide isnot apredictor of the colour of the cells themselves and that colours can be difficult to discern below 1000X magnification. Ifyou are not sure whether you are seeing dark pink or light purple, ask the lab demonstrator or instructor to confirm. Bacterial cultures for Gram stains (in tryptic soy broth tubes ) • Bacillus cereus • Pseudomonas flourescens • Micrococcus luteus • Escherichia coli • Staphylococcus epidermidis • Flexibacter canadensis • Unknown Mix I • Unknown Mix II Materials for Gram stains • Microscope slides • Methanol • Crystal violet • Gram ’siodine • Ethanol (ethyl alcohol), 95% • Safranin EXPERIMENT: Gram stains 1. Hold slides only by the edges to avoid getting smudges or grease on the slides. 2. Label your slides using agrease pencil; the methanol that you will use to fix the slide in step 7will remove permanent marker from glass. Itisuseful to draw an oval on the underside of the slide, near the middle, to help you find your bacterial smear after ithas dried. 3. When making asmear from aliquid culture, such as abroth, use asterile inoculating loop to transfer one or two loopfuls of liquid media directly onto the slide. Itisnot necessary to add saline. 4. Using the inoculating loop, spread the bacterial suspension into athin smear. 5. Allow the smear to air-dry. The slides can be dried more quickly by resting them on awarm slide drying bench. 6. Place the air-dried smear slide on your staining tray and use adropper bottle of methanol to cover the surface of the slide for two minutes. 7. Pour off the excess methanol into your staining tray and rinse the slide with water. 8. Cover the smear with crystal violet for 60 seconds. 9. Rinse the slide with water. 10. Cover the smear with Gram ’siodine for 60 seconds. 11. Rinse the slide with water. 12. Apply 95% ethanol drop by drop to the slide until run-off isclear (approximately 5-10 drops). 13. Rinse the slide with water. 14. Cover the smear with safranin for 45 seconds. 15. Rinse the slide with water. 16. Blot the slide in your book of bibulous paper. 17. View at 1000x magnification with immersion oil and make aquick sketch of what you see. BIOL2902L Fall 2020 17 18. After viewing each slide, use aKim-Wipe or microscope lens paper to clean the objective and ocular lenses and to remove any immersion oil from the microscope. 19. Repeat for the three bacterial cultures. CLEAN UP 1. Put your microscope slides in ayellow sharps bucket on the bench. 2. Wipe the oil off all of the objective lenses. Rotate the objective lens to 4x or the ‘empty ’space. Store the microscope under the dust cover, under your bench. 3. Use ascour pad to remove the labels from your culture broth tubes. Put the scrubbed tubes in the autoclave racks at the front of the lab. 4. Clean your bench with disinfectant and paper towel. 5. Remove your lab coat and put itin its plastic bag. 6. Wash your hands. BIOL2902L Fall 2020 18 Lab 4 : Unknown Bacteria I This lab will be delivered as self-directed material online through Nexus. A pre-recorded video outlining the experimental procedures will be uploaded to Nexus. You are to watch this video and read through this portion of the lab manual before completing the accompanying assignment. Due to time restraints, there isno “live ”portion for this lab and therefore no scheduled lab time. Ifyou have questions regarding this lab or the associated assignment please contact your instructor. Under normal circumstances the biochemical plate tests would be inoculated in one lab and the results would then be read the following week. Experimental guidelines for inoculating the biochemical tests is outlined in Part 1. Experimental guidelines for reading the plates after inoculation isoutlined in Part 2. Biochemical tests of unknown bacteria Each pair of students will be given three known (control) species and abroth that contains an uncontaminated unknown bacterial species. This unknown will be one of the six bacterial species we have worked with. Because we have atotal of six control species, students must form groups of four in order to compare their unknown to all six possible control species. You are responsible for sharing your results with another pair so that you have data for all six organisms. In today ’slab, we are reasonably confident that the unknown samples are not contaminated but, in practice, itisalways agood idea to perform afour-quadrant streak and isolate asingle colony before inoculating biochemical tests. Selective and differential media Specialized media have been developed to serve awide variety of functions in microbiology laboratories. Selective media incorporate chemicals that specifically inhibit some groups of organisms from growing while allowing (selecting for) other groups to grow. These media are often useful when trying to isolate aparticular type of bacteria out of amixed culture. For example, Phenylethyl Alcohol (PEA) agar inhibits the growth of Gram-negative organisms and selects for the growth of Gram-positive organisms.

Differential media can distinguish between different types of bacteria based on biochemical changes in the media that occur after inoculation and incubation. For example, blood agar differentiates between different types of bacteria based on their ability to destroy red blood cells (called haemolysis) and consume the nutrients released. Some media have both selective and differential properties. For example, mannitol salt agar selects for halophiles (salt-tolerant organisms) and inhibits non-halophiles. Of the salt-tolerant organisms that grow, the medium differentiates those that can ferment the carbohydrate mannitol from those that can ’t ferment mannitol. Chemically defined media are composed of known quantities of chemically pure, specific compounds. These are often relatively simple media and we know, on achemical level, exactly what isavailable to bacteria grown on that medium. We do not know the exact chemical composition of complex or undefined media. They often include plant, yeast or animal extracts with awide variety of amino acids, sugars, vitamins and minerals but the exact quantities of each constituent isunknown. Complex media are generally better at growing awider variety of bacteria with different nutritional requirements. This is not to give the impression that we have no idea what isin complex media; rather, adding aspecific number of grams of each amino acid and carbohydrate isgenerally more effort than it’sworth and referring to amedium as undefined acknowledges that there can sometimes be small variations in the exact composition of yeast extract from one batch to another. When recording your results for selective plates, organisms either grew or did not grow. For differential plates, there are generally two different appearances that may be visible, though some differential tests have more than two possible outcomes. For media that are both selective and differential plates, there are generally three different results you may observe. In the case of mannitol salt agar, ifthe organism is BIOL2902L Fall 2020 19 not salt-tolerant enough to grow on mannitol salt agar, we do not actually learn anything about whether or not itcan ferment mannitol (Table 1). Table 1. Possible observations following the incubation of bacteria on mannitol salt agar. Halophile Non-halophile Mannitol fermenter Growth, agar turns yellow No growth Mannitol non-fermenter Growth, agar remains red No growth Blood agar Blood agar (BA or TSA+5%SB) differentiates bacteria based on their haemolytic traits. Blood agar or sheep blood agar ismade using tryptic soy agar but with the addition of 5% sheep blood. Haemolysis, or the breakdown of red blood cells, can be observed through colour changes in the agar around the bacterial colonies. The bacterial enzymes that destroy red blood cells are called haemolysins. Ifthe bacterial haemolysins are able to damage the red blood cells and reduce haemoglobin to methemoglobin, the agar will develop agreenish colour but remain opaque. This result iscalled alpha haemolysis. Beta haemolysis occurs when the bacterial enzymes are able to completely destroy the red blood cells and metabolize the haemoglobin. The agar will become atranslucent yellow, similar to tryptic soy agar, because the red blood cells have been removed entirely. Ifthe bacteria does not produce haemolysins, there will be no significant change to the colour of the agar around the colonies. This isdescribed as gamma haemolysis. Haemolysis isparticularly useful for differentiating between Gram-positive cocci such as Streptococcus species. Blood agar isalso classified as an enriched medium because itcontains amuch higher level of nutrients than most agars. Some organisms are described as fastidious because they have specific nutrient requirements that are not met by most growth media and so they can only be grown on enriched media like blood agar. Eosin methylene blue agar Eosin methylene blue (EMB) agar selects for Gram-negative organisms and differentiates them based on the ability to ferment the carbohydrate lactose. Itcontains peptone and lactose as nutrients and the dyes eosin Yand methylene blue to inhibit growth of Gram-positive organisms. Lactose can be fermented into lactic acid, an organic acid that can accumulate and change the pH of the agar plate. The dyes act as apH indicator because changes in pH result in dyes being absorbed into the colonies, changing their colour. Under acidic conditions produced by vigorous fermentation, the dyes can produce adark purple colour and the colonies are often covered with ablackish-green metallic sheen. The dark purple colour or metallic sheen on the colonies indicates vigorous lactose fermentation expected with thermotolerant coliforms, aspecific group of Gram-negative bacilli that often indicate faecal contamination. Lower acidic production, as in cases of slow lactose fermenters, will result in pink colouration of the growth instead of dark purple. Non-fermenters remain their normal colour or the colour of the agar. EMB agar also contains peptone, aprotein digest that provides nutrients for organisms that are unable to ferment lactose. This agar isoften used to isolate faecal (thermotolerant) coliforms and can be streaked for bacterial isolation or used with membrane filtration to detect faecal contamination in water. MacConkey agar MacConkey (Mac) agar issimilar to EMB agar in that they both select for Gram-negative organisms and differentiate them based on lactose fermentation. On this agar, most Gram-positive organisms and many Gram-negative organisms are inhibited by the combination of crystal violet and bile salts. Enteric bacteria, bacteria that are often found in the intestines of warm-blooded animals, are typically best able to grow on MacConkey agar. Italso contains peptone, lactose and the pH indicator neutral red; it isred at pH of 6.8 and below and remains colourless above apH of 6.8. Increasing acidity from lactose fermentation turns the dye red and istypically an indicator of coliform bacteria such as E. coli .Non- fermenting coliforms such as Shigella (which can cause dysentery) and Salmonella (which can cause typhoid) will remain their normal colour or the colour of the medium, often increasing the plate ’spH by metabolising the peptone and making the plate less red. BIOL2902L Fall 2020 20 Mannitol salt agar Mannitol salt (MS) agar has ahigh saline content (7.5% sodium chloride), making ithostile to most bacteria. The most common clinically-relevant salt-tolerant organisms or halophiles are Gram-positive staphylococci, such as Staphylococcus aureus ,but itispossible for some other organisms to grow on this medium as well. Itcontains the pH indicator phenol red, which isyellow below pH 6.8, red at pH 7.4 to 8.4 and pink at pH 8.4 and above. Italso contains the carbohydrate mannitol, which can be fermented to produce organic acids that lower the plate ’spH. Yellow halos in the agar around colonies indicate mannitol fermenters while no colour change (reddish pink agar) around the colonies indicates non- fermenters of mannitol. Some laboratories use mannitol fermentation on MS agar as an indicator of pathogenic staphylococci but this isnot always accurate and should be confirmed with additional tests. Phenylethyl alcohol agar Phenylethyl alcohol (PEA) agar isaselective media that has abacteriostatic effect on Gram-negative and some Gram-positive organisms; these organisms are not killed by the medium but are unable to replicate as long as they are on the agar. Particularly heavy inoculation can sometimes look like growth but, ifthe growth isminimal compared to the same species on TSA or blood agar, the organism is probably Gram-negative. The phenylethyl alcohol increases membrane permeability, allowing potassium to leak out of the cell and disrupting DNA synthesis. This medium isoften used to isolate Gram- positive organisms out of amixed culture or swab that iscontaminated with Gram-negative organisms. Some PEA agar ismade with sheep blood to promote the growth of fastidious Gram-positive organisms; itcan also show evidence of haemolysis but this haemolysis isnot always as reliable as what can be observed on blood agar. In some cases, aGram-negative organism will grow long enough to cause limited haemolysis but then the colony will die from phenylethyl alcohol-related inability to replicate. Exoenzymes Many nutrients, particularly large molecules such as proteins, lipids and polysaccharides, are difficult to transport across acell membrane. Bacteria can secrete exoenzymes (extracelluar enzymes) to degrade these nutrients into smaller components that can be transported into the cell. These reactions are typically hydrolytic; water isintroduced into covalent bonds in order to break large molecules into smaller building blocks that can be taken up by the bacteria. Ifthe organism produces exoenzymes that degrade the macromolecules found on the plate, there will appear to be aprominent halo around the bacterial colonies. The casein hydrolysis test, lipid hydrolysis test and starch hydrolysis test are all examples of exoenzyme tests. In Lab 6, the gelatin hydrolysis test isalso an exoenzyme test. Casein hydrolysis test The casein hydrolysis test isused to detect bacteria that can hydrolyze the protein casein, the major protein in milk and the component that gives milk its white colour. Some bacteria produce and secrete avariety of exoenzymes into their environment to catalyze the break-down (hydrolysis) of proteins into smaller peptides or individual amino acids. This hydrolysis makes these peptides and acids accessible for uptake through the cellular membrane for use in bacterial protein synthesis or as an energy source. The enzyme responsible for breaking down casein iscalled casease. In theory, this agar could be modified to use any other protein to test for bacterial exoenzymes capable of hydrolyzing that particular protein. Milk agar contains yeast extract, dextrose, powdered milk and pancreatic digest of casein. When the milk agar isinoculated with acasease-positive organism, the casease secreted will create azone of clearing where the casease diffuses and the casein ishydrolyzed and becomes transparent. There will not be clearing zones around casease-negative organisms. Lipid hydrolysis test The lipid hydrolysis test or lipase test identifies lipolysis, the hydrolysis of lipids. which isaclearing of medium around the bacterial growth. The medium contains peptone, yeast extract, agar and some lipid substrate, such as corn oil, olive oil, soybean oil or high-fat dairy products. Lipids, ageneral term for fats, can hydrolysed by bacterial exoenzymes called lipases. Tributryrin oil isthe most common for lipase- testing media; itisthe simplest triglyceride in natural oils and fats, found in butter and margarine. Lipid agar made with tributyrin isoften referred to as tributyrin agar. BIOL2902L Fall 2020 21 Triglycerides are composed of three fatty acid chains and glycerol. Tributyrin istoo big to enter the cell without being broken down, so hydrolysis catalyzed by lipase isnecessary for cellular uptake. Tributyrin agar isan opaque emulsion that will show zones of clearing around the bacterial growth ifthe organism islipase-positive. Ifthe agar around the bacterial colonies remains opaque, the organism islipase- negative.

Starch hydrolysis test Starch agar isaplated medium of beef extract, agar and soluble starch .Starch ismade up of glucose subunits in linear (amylose) and branched (amylopectin) forms. In either form, starch cannot pass through bacterial cell membranes without being broken down; the enzymes α-amylase and oligo-1,6- glucosidase break the linear and branched glycosidic linkages and hydrolyze the starch into shorter polysaccharides and, with the aid of other enzymes, into glucose that can be absorbed by the cells. The starch and sugar subunits are soluble and thus very difficult to see without the use of an additional reagent; afew drops of iodine will be added after incubation (in Lab 6) to detect the starch around the bacterial growth, turning blackish-brown in contact with the starch. Microbial starch hydrolysis will then be visible as awhitish-yellow zone around the colonies. Uninoculated media Uninoculated media refers to media that have not been inoculated with bacteria. Because acolour- change isoften the indicator of abiochemical reaction, itishelpful to know what colour an agar plate or broth tube was before itwas inoculated so that you can see whether or not ithas changed colour. In the lab, we will provide examples of uninoculated plates so that you can always check what colour a test was prior to inoculation. Ifatest isinoculated and there isno visible growth, this result would be described as “inhibited growth ” or “no growth ”.Even though itmay look the same as the uninoculated plate, these results can be very important in identifying an organism. Workflow The following experiment was performed for you and adata set has been uploaded to Nexus for you to analyze. Although you did not perform the experiments yourselves, you are still required to understand the methods used and the theory behind the background information presented. Data has been provided to you for 4known species of bacteria and 1unknown. You will use the results from the known bacteria to determine the identity of the unknown bacteria. Bacterial cultures (per pai r) • Bacillus cereus • Escherichia coli • Staphylococcus aureus • Klebsiella pneumonia • Unknown bacteria Materials (per pair)  5blood agar (TSA w/5% SB)  5eosin-methylene blue agar (EMB)  5mannitol salt agar (MS)  5MacConkey agar (Mac)  5phenylethyl alcohol agar (PEA)  5tryptic soy agar  3casein (milk protein) agar (M)  3lipase (tributyrin) agar (T)  3starch agar (S) BIOL2902L Fall 2020 22 Part 1 EXPERIMENT: Selective, differential and enriched media 1. For this experiment, you will inoculate one organism per agar plate. Label your plates with the organism to be inoculated, the date and your lab section and seat number. 2. Using the four-quadrant streak plate technique, inoculate each of the agar plates with one organism per plate (blood agar, eosin-methylene blue agar, mannitol salt agar, MacConkey agar and phenylethyl alcohol agar). EXPERIMENT: Extracellular enzymatic activity (for casein hydrolysis, lipase and starch hydrolysis) 1. For this experiment, you will inoculate two organisms per agar plate. Label your plates with the organisms to be inoculated, the date and your lab section and seat number. 2. Divide each plate in half. 3. In the middle of each half, inoculate the casein agar, lipid agar and starch agar each with asingle streak of bacteria using an inoculating loop. Check for contamination of your bacterial samples (part 1) 1. Itispossible that you may have contaminated your bacterial samples while inoculating other agar plates during this lab. To detect any contamination that may have occurred, the last step of the lab will be to inoculate plates of tryptic soy agar with each of your organisms, one organism per plate. If these plates show more than one distinct colony morphology next week, itsuggests that you contaminated your bacterial broths. CLEAN UP 1. Remove the labels from culture tubes and inoculating suspensions and place them in the autoclave racks. 2. Put your agar plates into the five buckets provided for you and your partner and put them into the incubator. 3. Clean your bench with disinfectant and paper towel. 4. Remove your lab coat and put itin its plastic bag. 5. Wash your hands. Part 2 After 48 hours of incubation the results can then be read from the biochemical plates. The following methods are conducted to test for contamination and to read the results of the starch hydrolysis test. Check for contamination of your bacterial samples (part 2) 1. Observe your TSA plate sfrom Part 1.Ideally, itshows only one kind of colony morphology, suggesting that your inoculating broth has not been contaminated. 2. Ifyour plate shows more than one form of colony morphology (i.e. iscontaminated) and you can identify which organism isthe contaminant, please proceed with the dominant colony type. 3. Ifyour plate shows more than one form of colony morphology and you cannot determine which colony type isdominant or was most likely to be the original sample, you must choose one to identify. 4. Ifyou proceed with steps 2or 3from above, be aware that this means that itispossible for all of your results for the Selective, Differential and Enriched Media experiment to be inaccurate. EXPERIMENT: Starch hydrolysis test (part 2) 1. Add afew drops of iodine directly to the agar plate, on the margin of the bacterial colony. Ifstarch ispresent in the agar, itwill turn brownish-black with the addition of iodine. Ifthe starch has been hydrolysed by exoenzymes produced by the bacterial colony (a positive test result), there will no longer be starch in the media directly next to the bacterial colony and there will be ahalo of agar around the colony that will not turn black with the addition of iodine. BIOL2902L Fall 2020 23 CLEAN UP 1. Put your agar plates into the five buckets provided for you and your partner and put them into the incubator. 2. Clean your bench with disinfectant and paper towel. 3. Remove your lab coat and put itin its plastic bag. 4. Wash your hands. BIOL2902L Fall 2020 24 Lab 5 : Unknown Bacteria II This lab will be delivered as self-directed material online through Nexus. A pre-recorded video outlining the experimental procedures will be uploaded to Nexus. You are to watch this video and read through this portion of the lab manual before completing the accompanying assignment. Due to time restraints, there isno “live ”portion for this lab and therefore no scheduled lab time. Ifyou have questions regarding this lab or the associated assignment please contact your instructor. Part 1 Under normal circumstances the biochemical tube tests would be inoculated in one lab and the results would then be read the following week. Experimental guidelines for inoculating the biochemical tests is outlined in Part 1. Background theory information regarding 1-part biochemical tests isalso outlined in Part 1. Carbohydrate fermentation tests While most bacteria are able to obtain energy through some form of fermentation, the particular substrates or carbohydrates that different bacteria can ferment depends on that bacteria ’s complement of enzymes. The fermentation of carbohydrates requires an organic molecule to act as an electron donor and the final product(s) to accept that electron. Glucose fermentation begins with the organism producing pyruvate, typically through glycolysis in bacteria. The products of pyruvate fermentation are various acids and alcohols, and hydrogen or carbon dioxide, depending on the organism and the substrate fermented. Carbohydrate fermentation broths contain peptone, apH indicator and one fermentable carbohydrate. While any pH indicator can be used, our tests rely on phenol red, which isyellow below pH 6.8, red between pH 6.8 and 7.4 and pink or magenta above pH 7.4. Any carbohydrate functions for the test, but glucose, lactose and sucrose are frequently used. A small inverted tube, called aDurham tube, isput into each larger test tube to capture any gas produced through fermentation. Carbohydrate fermentation produces an organic acid and decreases the pH below the neutral range for the indicator, turning the broth yellow. Ifthe organism metabolizes the peptone through deamination, the resulting ammonia released into the broth increases the pH and turns the broth pink. Gas production can be visible by the accumulation of abubble in the Durham tube where the broth is displaced.

The detection of acid production in the medium largely depends on incubation time and the fermenter ’sability to produce excess acid compared to the ammonia resulting from deamination. At 24 hours, apink colour in the phenol red broths shows that the organism did not ferment the substrate and has deanimated the peptone amino acids. Readings after 24 to 48 hours may be unreliable: ifthe medium shows acid production, the reading has no problem; however, lack of colour change or indicators of alkalinity could result from reversion. A reversion isthe result of the organism performing deamination due to complete consumption of the carbohydrate. The resulting shift from acid to alkaline can mask indicators of fermentation, because the broths shifting to pink or purple makes itimpossible to tell between areversion and atrue non-fermenter. Decarboxylase broth Decarboxylase broths detect the presence of different decarboxylase enzymes based on how the organism breaks down any one of the many amino acids used in the media. A decarboxylase enzyme can remove the carboxyl group (–COOH) from aspecific amino acid, producing carbon dioxide and amines that cells can use to synthesize other molecules. The base medium for decarboxylase broth can be used with one of many amino acid substrates to identify different decarboxylase enzymes. Our lysine decarboxylase broth contains peptone, glucose and the pH indicator bromocresol purple, which is purple at pH 6.8 and up and yellow at apH of 5.2 or lower. As anote of interest, the amine produced from the decarboxylation of lysine iscadaverine, afoul-smelling diamine produced by the putrefaction of animal tissue. BIOL2902L Fall 2020 25 After the broth isinoculated, an overlay of mineral oil isapplied to prevent oxygen from reaching the broth, making an anaerobic environment to promote fermentation. Ifthe organism isable to ferment glucose, organic acids accumulate and turn the pH indicator yellow. The bacterial gene that encodes for decarboxylase enzymes isturned on in the presence of low pH and specific amino acids in the medium. Ifthe organism isable to make adecarboxylase enzyme, itwill decarboxylate the available amino acids, resulting in alkaline products that will raise the pH and turn the broth purple. Ifthe organism isaglucose fermenter, but does not produce decarboxylase, the medium turns yellow and stays yellow. Ifthe organism does not ferment glucose, there will be no colour change. In our experiments, we use two different tubes for lysine decarboxylation testing. One tube contains the base medium described above and the amino acid L-lysine (labeled LD+). The other tube contains the same base medium but does not contain any amino acids (labeled LD −).The only positive result isifthe LD+ tube ispurple and the LD −tube isyellow. In this case, fermentation of glucose occurred, making both tubes acidic (yellow) but the decarboxylation of lysine caused the pH to rise in the LD+ tube, reverting itto purple. The LD −remains yellow because, even though the organism iscapable of making alysine decarboxylase enzyme, there isno L-lysine in that tube. A negative reaction occurs when both LD+ and LD −are yellow, glucose fermentation occurred but the organism was unable to produce alysine decarboxylase enzyme, so the broths remain acidic. All of the conditions for lysine decarboxylase enzyme production occurred in the LD+ tube (acidic environment and presence of amino acid) but there was no rise in pH, suggesting the organism isincapable of producing that enzyme. Ifboth LD+ and LD −are purple, itislikely that glucose fermentation did not occur. Ifthe broth never became acidic, the organism would not have produced any decarboxylase enzyme, so we do not know whether or not itcarries that gene. This result isdescribed as inconclusive. Urease test Urea results from decarboxylation of particular amino acids. Itcan be broken into ammonia and carbon dioxide by bacteria containing the enzyme urease. Many bacteria can metabolize urea but asmaller number of them metabolize itrapidly enough to give apositive broth test result. Urea broth contains urea, yeast extract, potassium phosphate and phenol red. The yeast extract isa nutrient source, the potassium phosphate acts as apH buffer to resist alkalinisation and the phenol red pH indicator isyellow or orange below pH 8.4 and red or pink above that pH. Hydrolysis of urea to ammonia will overcome the buffer and change the medium from orange to pink, indicating arapid urease-positive organism, while orange or yellow isconsidered anegative result for this test. Citrate utilization test The citrate utilization test determines the organism ’sability to use citrate as its sole carbon source. This is part of apanel of tests called the IMViC (Indole, Methyl Red, Voges-Proskauer and Citrate) tests, commonly used to differentiate Gram-negative bacilli, particularly members of the family Enterobacteriaceae, which includes the coliform bacteria. Citrate, or citric acid, isproduced when acetyl coenzyme A reacts with oxaloacetate at the beginning of the Krebs cycle. Bacteria possessing citrate-permease can take in citrate and convert it enzymatically to pyruvate, which can be transformed into avariety of different products. Simmons citrate agar isadefined medium that provides sodium citrate as the only available source of carbon and ammonium phosphate as the only source of nitrogen. The pH indicator, bromothymol blue dye, turns green at pH 6.9 and blue at pH 7.6. Bacteria that can use citrate as carbon will also convert ammonium phosphate to ammonia (NH 3)and ammonium hydroxide (NH 4OH) to get nitrogen. These both alkalinize the agar and as the pH goes up, the medium shifts from green to blue, indicating a positive citrate test result. Citrate-positive organisms can occasionally grow on aSimmons citrate slant without avisible colour- change. This absence of colour change istypically because of incomplete incubation. Even in the BIOL2902L Fall 2020 26 absence of avisible colour-change, growth on the agar slant demonstrates citrate utilization and isa positive result. Triple sugar iron agar Triple sugar iron agar (TSIA) isadifferential test based on fermentation of the carbohydrates glucose, lactose and sucrose, and sulfur reduction. The agar contains 1% lactose, 1% sucrose and 0.1% glucose (dextrose). The medium also contains beef extract, yeast extract and peptone to provide carbon and nitrogen, and sodium thiosulfate as areducible sulfur compound. Phenol red isthe pH indicator and iron in the ferrous sulfate indicates the level of hydrogen sulfide. The medium isprepared in atube on aslant but filled with more medium than most agar slants, providing an aerobic environment along the slant and an anaerobic environment at the bottom of the agar (called the butt of the tube). Itisinoculated using astab-and-streak technique described in the experimental procedures. Carbohydrate fermentation needs 18 to 24 hours of incubation and up to 48 hours for hydrogen sulfide reactions. There are various combinations of reactions that can take place. A glucose-only fermenter will produce acid and lower the pH, turning the medium yellow within afew hours. This reaction consumes the small amount of glucose, and the organisms in the aerobic slant will hydrolyse the amino acids and produce NH 3,raising the pH. The process, called reversion, takes 18 to 24 hours only occurs in the aerobic portion of the medium. A TSIA with ared slant and yellow butt ispositive for glucose-fermentation and negative for lactose-fermentation and sucrose-fermentation. Organisms fermenting lactose and/or sucrose turn the full medium yellow; because of the concentrations in the TSIA, amedium with ayellow slant and butt at 24 hours indicates the organism ferments at least one of these sugars. Because the medium isalready yellow, this result does not indicate which of the two sugars turned the entire agar yellow and whether or not the organism isalso capable of fermenting glucose. In some cases, carbon dioxide gas produced by carbohydrate fermentation will lift the agar off the tube or cause fissuring. Ifthe organism isunable to ferment lactose, sucrose or glucose, the tube will show ared alkaline slant and an orange-red butt (for organisms that can only catabolize peptone aerobically) or ared alkaline butt (indicating aerobic and anaerobic utilization of peptone). Timing reading the test can impact the result, as an early reading might result in glucose fermentation falsely looking like lactose or sucrose fermentation when the glucose in the agar has not yet been exhausted. Reading after the lactose and sucrose are depleted, allowing reversion to occur, would show ayellow butt and red slant, falsely suggesting the organism only ferments glucose. Bacteria can anaerobically reduce sulfur to produce hydrogen sulfide (H 2S) either by reducing thiosulfate or by hydrolysing the amino acid cysteine, present in peptone, into pyruvate. In either case, H2Siscolourless so we detect itwhen itreacts with iron ions in the medium to form ferrous sulfide (FeS), a black precipitate, typically in the butt of the agar. Acidic conditions must exist for this reaction, so the black precipitate indicates sulfur reaction and fermentation. The black precipitate can obscure the colour reading in the butt, meaning the carbohydrate fermentation needs to be read by the colour reaction in the slant. Motility test Motility test agar isasemisolid; its concentration of agar isreduced to 0.4%, enough to preserve its form while permitting movement of motile bacteria. Peptone and beef extract provide nutrients for growing bacteria. Inoculation isdone with astraight transfer needle; growth radiating out from the straight stab line demonstrates motility. To enhance the visibility of the bacteria, 2,3,5-triphenyl tetrazolium chloride (TTC) can be added to help read results. As the growing bacteria use TTC as an electron acceptor, it becomes red and insoluble, indicating where the bacteria are in the tube. Red TTC precipitate radiating out from the inoculation line shows motility whereas red only along the stab line isanegative result.

SIM medium isan agar designed to detect sulfur reduction, indole production and motility. Because SIM lacks TTC, motility in this medium can be harder to see. BIOL2902L Fall 2020 27 Sulfur reduction test The sulfur reduction test isperformed using SIM agar, amultifunction medium that can be used to detect sulfur reduction, indole production and motility. The test uses SIM medium, formulated with casein and animal tissue, amino acids, an iron-containing compound and sodium thiosulfate for sulfur. The mechanisms of detecting sulfur reduction to hydrogen sulfide (H 2S) in SIM agar are the same as in triple sugar iron agar. Bacteria can anaerobically reduce sulfur either by reducing thiosulfate or by hydrolysing the amino acid cysteine, present in peptone, into pyruvate. Either reaction produces hydrogen sulfide, which reacts with iron ions in the medium to form ferrous sulfide (FeS), avisible black precipitate. No visible blackening in the medium isanegative result. Workflow The following experiment was performed for you and adata set has been uploaded to Nexus for you to analyze. Although you did not perform the experiments yourselves, you are still required to understand the methods used and the theory behind the background information presented. Data has been provided to you for 4known species of bacteria and 1unknown. You will use the results from the known bacteria to determine the identity of the unknown bacteria. Bacterial cultures (per pair) • Bacillus cereus • Escherichia coli • Staphylococcus aureus • Klebsiella pneumonia • Unknown bacteria Materials (per pair) • 5glucose fermentation broths with Durham tube (yellow dot) • 5lactose fermentation broths with Durham tube (red dot) • 5sucrose fermentation broths with Durham tube (blue dot) • 5lysine decarboxylase broths with L-Lysine (LD +) • 5lysine decarboxylase broths without L-Lysine (LD –) • 5MRVP broths for methyl red (MRVP) • 5nitrate reduction broths (N) • 5urease broths (U) • 5MRVP broths for Voges-Proskauer (MRVP) • 5citrate agar slants (C) • 5phenylalanine deaminase agar slants (PD) • 5triple sugar-iron agar slants (TSI) • 5motility deep agars (M) • 5sulfur-indole-motility deep agars (SIM) • 5tryptic soy agar (TSA) Table 2: Biochemical tube tests performed during Lab 6. Label each tube with anumber (1-14) and an organism (BC, EC, SA ,and KP ). Media Contents Tube (Label) Inoculation Broths Glucose fermentation (with Durham tube) 1(yellow dot) 2drops Lactose fermentation (with Durham tube) 2(red dot) 2drops Sucrose fermentation (with Durham tube) 3(blue dot) 2drops Lysine decarboxylase with L-Lysine 4(LD +) 2drops Lysine decarboxylase without L-Lysine 5(LD –) 2drops Methyl Red 6(MRVP) 2drops BIOL2902L Fall 2020 28 Nitrate reduction 7(N) 2drops Urease 8(U) 2drops Voges-Proskauer 9(MRVP) 2drops Agar slants Citrate 10 (C) Streak Phenylalanine deaminase 11 (PD) Streak Triple Sugar-Iron (TSI) 12 (TSI) Stab-and-streak Deep agar Motility 13 (M) Stab Sulfur-Indole-Motility (SIM) 14 (SIM) Stab Preparation of broth suspension 1. Using your inoculating loop, collect abacterial colony from the provided bacterial cultures on TSA plate s. 2. Suspend itin tryptic soy broth. 3. All of your inoculations for today ’sexperiments will be done using this broth suspension. EXPERIMENTS: Inoculation of broth cultures (for glucose fermentation, lactose fermentation, sucrose fermentation, lysine decarboxylase, methyl red, nitrate reduction, urease and Voges-Proskauer tests) 1. Collect abacterial colony from your TSA plate using your inoculating loop. 2. Suspend itin tryptic soy broth. 3. Inoculate your other liquid cultures by adding tryptic soy broth using Pasteur pipettes. 4. Please use 2drops for each broth tube. Note: Inoculation of lysine decarboxylase (LD+ and LD –)broths 5. After all of your tubes have been inoculated, add alayer of mineral oil to restrict oxygen and prevent false alkalization in the LD tubes. Add the mineral oil while holding the tube in aslanted position. Try not to let the tip of the Pasteur pipette touch the walls or inoculated broth. Add enough mineral oil to cover the surface of the broth with avisible layer of oil. EXPERIMENTS: Inoculation of agar slants (for citrate, phenylalanine deaminase and triple sugar-iron tests) 1. Dip the inoculating loop in your broth suspension. 2. Touch the inoculating loop to the agar slant near the bottom of the slant and drag itup the slant towards the mouth of the tube, streaking itslightly side-to-side. 3. Sterilize the inoculating loop. Note: “Stab-and-streak ”inoculation of triple-sugar iron (TSI) agar slants 1. Dip the inoculating needle in your broth suspension. 2. Insert to the bottom of the agar tube, like when inoculating deep agar. 3. Draw straight up out of the agar until itbreaks the surface and drag itup the slant towards the mouth of the tube, streaking itslightly side-to-side. EXPERIMENTS: Inoculation of deep agar (for motility and sulfur-indole-motility tests) 1. Dip the inoculating needle in your broth suspension. 2. Insert to the bottom of the agar tube and remove itin one smooth motion. 3. The gelatin tubes will be incubated at 35 °C for five days. They will then be refrigerated to attempt to solidify the gelatin. Ifthe gelatin has been hydrolyzed, itwill no longer solidify. Check for contamination of your bacterial samples 1. Itispossible that you may have contaminated your bacterial inoculating broth suspension during this lab. To detect any contamination that may have occurred, the last step of the lab will be to inoculate plates of tryptic soy agar with each of your organisms, one organism per plate. Ifthese plates show more than one distinct colony morphology next week, that suggests that you contaminated your bacterial broths. 2. Label your tryptic soy agar plates with the organism to be inoculated, the date and your lab section and seat number. BIOL2902L Fall 2020 29 CLEAN UP 1. Place your TSA control plates in racks on the side bench. Each plate must be labeled with the species and your lab section and seat number. 2. Place all of your remaining inoculated test tubes in abucket and label the bucket with your lab section and seat number. Itispossible to carefully fit all 30 tubes into asingle bucket. Carefully place the bucket in the incubator on the shelf marked for your lab section. 2. Dispose of the agar plates from Lab 5in the autoclave bins at the front of the room. 3. Clean your bench with disinfectant and paper towel. 4. Remove your lab coat and put itin its plastic bag. 5. Wash your hands. Part 2 Under normal circumstances the biochemical tube tests would be inoculated in one lab and the results would then be read the following week. Experimental guidelines for reading the biochemical tube tests and performing the rapid tests isoutlined in Part 2. Background theory information regarding multi-step biochemical tests and rapid tests isalso outlined in Part 2. Catalase test During aerobic respiration, the electron transport chains of aerobic, facultatively anaerobic and microaerophilic bacteria use oxygen as aterminal electron acceptor. While most of this oxygen isused to produce water, avery small percentage of itproduces reactive oxygen species such as superoxide radical (O 2−)and hydrogen peroxide (H 2O2).To keep these toxic products from oxidizing and damaging proteins, these organisms produce the enzymes superoxide dismutase and catalase hydrolyze these products into oxygen and water. Superoxide dismutase, which hydrolyzes the superoxide radical into hydrogen peroxide, isso important that itisfound in almost all aerobic cellular life. Because hydrogen peroxide ismuch less toxic than superoxide, not all aerobes possess the enzyme catalase, which hydrolyzes hydrogen peroxide into oxygen and water. The inability of strict anaerobes to produce these enzymes may explain why oxygen istoxic to these microbes. The addition of the hydrogen peroxide to acatalase-positive organism will result in immediate and vigorous bubbling. Ifthere are only afew bubbles. The absence of bubbling, or afew number of bubbles after asignificant delay, should be interpreted as acatalase-negative result. Clinically, the catalase test isoften used to differentiate catalase-positive Staphylococcus species from catalase- negative Streptococcus species. Oxidase test Oxidases play avital role in the electron transport chain (ETC) during aerobic respiration. The two major functions of the electron transport chain are to transport electrons to lower energy levels and eventually to aterminal electron acceptor and to generate proton motive force by removing H+from the cell to drive production of ATP from the ion gradient. Organisms can have multiple forms of ECTs and many bacteria have ECTs that resemble mitochondrial ECTs. These chains, consisting of enzymes named Complexes I,II, IIIand IV, contain molecules that transfer electrons and use the energy released in the reactions. The last enzyme, Complex IV, also called cytochrome coxidase, makes the final electron transfer from cytochrome cto the terminal electron acceptor, oxygen. The oxidase test specifically detects cytochrome coxidase, which can oxidize cytochrome cand can catalyze the reduction of cytochrome cby p-aminodimethylaniline oxalate (DPD). Ifthe bacteria are added to afilter paper saturated with DPD, apositive result will show acolour change to bluish-purple within 30 seconds, while alack of change or achange to any other colour indicates that cytochrome c oxidase isnot present. Because apurple colour isapositive test result, itisbest not to transfer from bacteria grown on amedium that contains purple dyes, such as EMB or MacConkey agars. Some species can use oxygen as aterminal electron acceptor in their electron transport chains but, because they do not contain the enzyme cytochrome coxidase, these species are oxidase-negative. Potassium hydroxide (KOH) Gram reaction test BIOL2902L Fall 2020 30 Potassium hydroxide provides ameans of confirming Gram reaction without staining: emulsifying part of acolony in KOH for one minute will result in either aviscous, stringy and adhesive suspension (Gram- negative) or no change to the emulsion (Gram-positive). However, there are many species that can provide ambiguous or conflicting results, so this test isnot considered to be as reliable as aconventional Gram stain. Methyl Red test The methyl red test detects the ability to produce acid end products from mixed-acid fermentation of glucose. Itispart of apanel of tests called the IMViC (Indole, Methyl Red, Voges-Proskauer and Citrate) tests, commonly used to differentiate between Gram-negative bacilli, particularly members of the family Enterobacteriaceae, which includes coliform bacteria. MRVP broth isused for both the methyl red and Vogues-Proskauer tests. Itcontains glucose as a fermentable carbohydrate, peptone as aprotein and phosphate as abuffer to resist pH changes. The difference isthat, following incubation, the methyl red test uses the pH indicator methyl red instead of Barritt ’sreagents used in the Voges-Proskauer test. While many bacteria can use glucose for energy production, the end products of this process vary based on the specific enzyme pathways that the bacteria possess. The methyl red test detects organisms that can accomplish mixed-acid fermentation, overcoming the potassium phosphate and lowering the pH. Acids produced by these bacteria, such as lactic acid, acetic acid and formic acid, are stable and maintain apH of approximately 4in the tube. Methyl red-negative organisms convert these acids into non-acidic end products such as 2,3-butanediol and acetoin, resulting in an elevated pH of approximately 6. Adding methyl red indicator after incubation confirms the mixed acid fermentation; methyl red isred at pH 4.4, indicating apositive result, yellow at pH 6.2, indicating aclear negative, and orange for an inconclusive result. Nitrate reduction test Some aerobic and facultative anaerobic microbes are able to reduce nitrates in the absence of molecular oxygen. In these organisms, anaerobic respiration isan oxidative process in which inorganic compounds like nitrates (NO 3−)or sulfates (SO 42−)can act as terminal electron acceptors. In the case of nitrate, itisreduced with electrons and hydrogen ions by the enzyme nitrate reductase to produce nitrite (NO 2−)and water. Some bacteria possess enzymes to perform amulti-step process called denitrification to convert nitrate to ammonia (NH 3+)or molecular nitrogen (N 2). Nitrogen broth isan undefined medium containing beef extract, peptone and potassium nitrate (KNO 3). The medium ismade semisolid by the addition of 0.1% agar to reduce the diffusion of oxygen and facilitate the anaerobic environment necessary for nitrate reduction. No colour indicator isin the broth, as colour reactions are visible from the nitrate reduction reagents added after incubation. Following incubation, sulfanilic acid (nitrate reagent A) and α-naphthylamine (nitrate reagent B) are added to the broth. Ifnitrite ispresent, itwill form nitrous acid (HNO 2)in the water of the broth and will react with nitrate reagents A and Bto produce ared water-soluble compound, indicating that the single-step reduction of nitrate to nitrite has occurred. Ifno colour change occurs after the reagents have been added, this could be because the nitrate was not reduced or itwas rapidly reduced beyond nitrite to ammonia or nitrogen. To determine which of these reactions has occurred, zinc isadded to the broth already containing reagents A and B. The zinc will act as an non-biologic catalyst to reduce nitrate to nitrite. This reduction will result in the earlier test result of the medium turning red, indicating that the organism did not reduce the nitrate. Ifno red colour isproduced after the addition of zinc, this indicates that the organism broke down the nitrate to ammonia or gaseous nitrogen. The most common challenge in interpreting this test istrying to think of red indicating apositive reaction or anegative reaction but this test has three possible results: nitrate reduction to nitrite, nitrate reduction to ammonia or gaseous nitrogen, or lack of nitrate reduction. The medium turning red indicates a different result based on whether you have added zinc. Because the broth isquite thick, the colour- BIOL2902L Fall 2020 31 change reaction can be slower than some of our other biochemical reactions. Ifyou cannot tell ifa reaction isoccurring in the tube, set itaside for five to ten minutes and check on itat that point. Ifthe tube isbecoming red, itwill continue to get redder, making iteasier for you to interpret the result. Voges-Proskauer test The Voges-Proskauer test identifies organisms that ferment glucose and transform the organic acid products to acetoin and 2,3-butanediol. This test ispart of the IMViC (Indole, Methyl Red, Voges- Proskauer and Citrate) panel of tests, commonly used to differentiate Enterobacteriaceae and other Gram-negative bacilli. MRVP broth isused for both the methyl red and Vogues-Proskauer tests. Itcontains glucose as a fermentable carbohydrate, peptone as aprotein and phosphate as abuffer to resist pH changes. The difference isthat, following incubation, the Voges-Proskauer test uses Barritt ’sreagents. Barritt ’sreagents (5% α-naphthol in ethanol and 40% potassium hydroxide) oxidize any acetoin that may be present following incubation into diacetyl. The diacetyl can then react with guanidine nuclei from the peptone to produce areddish-pink complex that indicates apositive result. No colour change when the reagents are added isanegative result. An interaction between the reagents can produce a copper colour, which isnot the same as ared positive result. To help you to interpret this test, positive and negative comparison controls are provided on the side bench. Phenylalanine deaminase test Deaminase-positive organisms are able to use oxygen and water to remove the amine group (–NH 2) from amino acids, producing aketo acid and ammonia. In the case of the amino acid phenylalanine, the specific enzyme phenylalanine deaminase produces phenylpyruvic acid and ammonia. After incubation, a10% solution of ferric chloride (FeCl 3)isadded to the agar slant because α-and β-keto acids produce acolour reaction with ferric chloride solutions. Ifthe organism ispositive for phenylalanine deaminase, the ferric chloride reacts with phenylpyruvic acid and rapidly turns dark green. A negative result shows the yellow colour of the ferric chloride solution. The positive dark green reaction will fade over time, so you should read the test within 30 minutes of adding the ferric chloride. Indole test The indole test distinguishes bacteria that produce indole by hydrolyzing the amino acid tryptophan using tryptophanase. Itispart of apanel called the IMViC (Indole, Methyl Red, Voges-Proskauer and Citrate) tests, commonly used to differentiate Gram-negative bacilli, particularly members of the family Enterobacteriaceae. This test istypically performed using SIM agar, asemi-solid medium made of casein and animal tissue for amino acids, an iron-containing compound and sodium thiosulfate for sulfur. Indole production results from bacteria possessing tryptophanase hydrolyzing the tryptophan in the casein and animal protein into pyruvate, ammonia and indole. The indole isdetected by adding Kovac ’sreagent, which isasolution of dimethylaminobenzaldehyde (DMABA) and hydrochloric acid in amyl alcohol. Kovacs ’reagent will form aliquid layer on top of the solid SIM agar the tube. This liquid layer will turn red ifthe DMABA reacts with any indole, undergoing adehydration reduction to produce aquinoidal red-violet compound that isindicative of apositive result. Ifno red compound isproduced, the organism isindole-negative. Workflow The following experiment was performed for you and adata set has been uploaded to Nexus for you to analyze. Although you did not perform the experiments yourselves, you are still required to understand the methods used and the theory behind the background information presented. Data has been provided to you for 4known species of bacteria and 1unknown. You will use the results from the known bacteria to determine the identity of the unknown bacteria. BIOL2902L Fall 2020 32 Materials (per pair) • Microscope slides • Hydrogen peroxide • p-aminodimethylaniline oxalate (note that several different redox indicator can be used for this test) • Methyl red • Sulfanilic acid (nitrate reduction solution A) • α-naphthylamine (nitrate reduction solution B) • Powdered zinc and applicator sticks • Barritt ’sreagent A (5% α-naphthol in ethanol) • Barritt ’sreagent B(40% potassium hydroxide) • Ferric chloride solution (10%) • Kovac ’sreagent • 3% KOH solution Check for contamination of your bacterial samples (part 2) 1. Observe your TSA plate sfrom Part 1.Ideally, itshows only one kind of colony morphology, suggesting that your inoculating broth has not been contaminated. 2. Ifyour plate shows more than one form of colony morphology (i.e. iscontaminated) and you can identify which organism isthe contaminant, please proceed with the dominant colony type. 3. Ifyour plate shows more than one form of colony morphology and you cannot determine which colony type isdominant or was most likely to be the original sample, you must choose one to identify. 4. Ifyou proceed with steps 2or 3from above, be aware that this means that itispossible for all of your results for the Selective, Differential and Enriched Media experiment to be inaccurate. EXPERIMENT: Catalase Test 1. Transfer acolony of bacteria from your TSA contamination plate to aglass microscope slide. 2. Add one drop of hydrogen peroxide (H 2O2). 3. Examine for the presence of immediate and vigorous bubbling or foaming. EXPERIMENT: Oxidase Test 1. Transfer acolony of bacteria from your TSA contamination plate to afilter paper wet with N,N- Dimethylbenzene-1,4-diamine. 2. Examine for colour change to purple in 10-30 seconds. EXPERIMENT: KOH Gram reaction 1. Place adrop of 3% KOH to aclean glass slide. 2. Transfer aloopful of bacteria from your TSA contamination plate the previous lab and mix the cells into the KOH using the inoculating loop constantly for 60 seconds. 3. Examine the consistency of the bacterial suspension. Ifitisstringy, the culture isGram-negative. Ifno stringiness isobserved, the culture isGram-positive. Note that using too much KOH can produce a false-negative result (the mixture iswatery, suggesting that the organism isGram-positive) and using too little KOH can produce afalse-positive (the mixture isviscous, suggesting that the organism is Gram-negative) result. EXPERIMENT: Methyl Red test (part 2) 1. Add 5drops of methyl red. 2. Observe the colour. EXPERIMENT: Nitrate reduction (part 2) 1. Add 5drops of solution A (sulfanilic acid). 2. Add 5drops of solution B(α-naphthylamine). 3. Observe the colour. 4. Ifatube did not turn red after step 2, wait five to ten minutes. Ifitstill has not turned red, insert a wooden stick into the zinc dust and then stir that stick into the tube. The amount of zinc that will remain on the stick issufficient to complete the reaction. BIOL2902L Fall 2020 33 5. Observe the colour. Ifitdid not turn red, wait an additional five to ten minutes before making your final observation. EXPERIMENT: Voges-Proskauer test (part 2) 1. Add 5drops of Barritt ’sReagent A. 2. Add 5drops of Barritt ’sReagent B. 3. Gently mix the tube for 1minute. 4. Observe the colour after 30 minutes but within 1hour. EXPERIMENT: Phenylalanine deaminase (part 2) 1. Add 5drops of 10% ferric chloride solution and mix gently. 2. Observe the colour. The results will fade over time so the ferric chloride solution can be reapplied. EXPERIMENT: Indole test (part 2) 1. Observe ifthe deep tube has become blackened, which isan indication of sulfur reduction. 2. Add 10 drops of Kovac ’sreagent to deep tube and agitate gently. 3. Observe results of indole test. Record all remaining results for tubes inoculated during Lab 7. CLEAN UP 1. Remove all labels from all test tubes. 2. Sort tubes by type and place each type in aseparate autoclave rack at the front of the lab. • Broths with Durham tubes (1, 2, 3) • Broths (4, 5, 6, 7, 8, 9, 10) • Agar slants and deep agars (11, 12, 13, 14, 15) 3. Dispose of all agar plates in the autoclave bags at the front of the lab. 4. Clean your bench with disinfectant and paper towel. 5. Remove your lab coat and put itin its plastic bag. 6. Wash your hands. BIOL2902L Fall 2020 34 Lab 6 : Bacterial Growth I This lab will be delivered as self-directed material online through Nexus. A pre-recorded video outlining the experimental procedures will be uploaded to Nexus. You are to watch this video and read through this portion of the lab manual before completing the accompanying assignment. Due to time restraints, there isno “live ”portion for this lab and therefore no scheduled lab time. Ifyou have questions regarding this lab or the associated assignment please contact your instructor. Serial dilutions Chemistry and biology have different notation when writing about dilutions. Both of the following are instructions to make a10% solution. As this isa(micro)biology class, we will use the biology notation. Be aware that you will occasionally find biologists and biology textbooks that format the numbers like a ratio but mean the numbers as afraction (e.g. 1:10 instead of 1/10for a10% dilution), which can be confusing. Biology dilution notation for a10% solution “one-in-ten dilution ” 1/10 Chemistry dilution notation for a10% solution “one-to-nine dilution ” 1:9 (part solute) dilution (total parts) (parts solute) dilution (parts solvent) Serial dilution issimply adilution procedure performed repeatedly. Ifyou take 1mL of ethyl alcohol, add itto 9mL of water and mix the two liquids, you will get a10% ethyl alcohol solution. Ifyou then take 1mL of that 10% solution, add itto another 9mL of water and mix, you will get a1% alcohol solution. The procedure can be repeated subsequently to produce a0.1% solution, a0.01% solution, a0.001% solution, a0.0001% solution (only one part per million ethyl alcohol) and as many smaller dilutions as are needed. The major advantage of serial dilutions isaccuracy; ifyou wanted to make a0.0001% ethyl alcohol solution in asingle step, you would either need to combine such extreme volumes as 1mL ethyl alcohol to 99.999L of water or 0.1 μLethyl alcohol to 9.9999mL of water, and itwould be easy for asmall error in measurement to result in asolution that ismuch stronger or weaker than the one desired. Serial dilution allows for the accurate preparation of solutions using convenient volumes. When performing aserial dilution, itisessential to mix each dilution thoroughly before proceeding to the next dilution. When possible, this should be done with careful vortexing. There isasmall volume of liquid left behind inside apipet after the contents have been dispensed. Ifthe same pipet isused for each of the serial dilutions, the carryover from the first dilution can be asignificant source of error in the second and subsequent dilutions. Therefore, whenever performing a1/10or greater dilution, change your pipet between dilutions. Ifyou are performing a1/2dilution (a 50% dilution), the carryover inside the pipet will not be asignificant source of error and itisnot necessary to change the pipet. In general, itismore accurate to change the pipet between dilutions but not changing pipets is not always aproblem, depending on the magnitude of the dilutions. Imagine that you draw liquid up into apipet from Tube 1and transfer itinto Tube 2; abit of residual liquid remains in the pipet (up to 2%). When the liquid from Tube 2isdrawn up to be transferred into Tube 3, itmixes with the residual liquid that was left inside the pipet. Because that residual liquid isthe concentration of Tube 1, itwill increase the concentration of the liquid being transferred. Ifthe difference in concentration between Tube 1and Tube 2isrelatively small, this increase in concentration will be insignificant. Ifthe difference in concentration between Tube 1and Tube 2islarge, the residual liquid in the pipet can significantly reduce the dilution. Example of 1/10dilution using 1mL transfer volume. When transferring from Tube 2to Tube 3, the pipet contains 1mL of 10% +0.02mL of 100% =12%. Tube 3will be 20% more concentrated than itshould be (10% vs. 12%). This isasignificant source of error. Example of 1/2dilution using 1mL transfer volume. When transferring from Tube 2to Tube 3, the pipet contains 1mL of 50% +0.02mL of 100% =52%. Tube 3will be 4% more concentrated than itshould be (50% vs. 52%). While this isobviously still asource of error, it BIOL2902L Fall 2020 35 generally falls within what isacceptable or at least within the margin of what can be caused by human pipetting error. Spectrophotometry Spectrophotometry isthe measurement of light of aspecific wavelength to learn about the substance through which the light has travelled. Absorption of light isused as an indicator that there issomething blocking that light (i.e. bacteria). The more light isblocked, the more bacteria are present. Spectrophotometers can provide output in the form of either “transmission ”or “absorbance ”.For our experiments, we will use absorbance of light at 600 nm wavelength. Absorbance ismeasured in units A or AU. An absorbance of 0means that 100% of the light gets through, compared to ablank or reference sample. An absorbance of 1means that 10% of the light gets through and an absorbance of 2means that only 1% of the light gets through. Workflow The following experiment was performed for you. Although you did not perform the experiments yourselves, you are still required to understand the methods used and the theory behind the background information presented. Bacterial cultures (in saline, per pair) • Escherichia coli Materials (per pair) • 5small tubes containing 3mL tryptic soy broth for spectrophotometry • 1small tube containing 5mL tryptic soy broth for spectrophotometry • 10 large tubes containing 9mL sterile saline for pour plates • 250mL molten agar for pour plates, kept in awater bath at 55 C until needed • 8empty Petri plates EXPERIMENT: Quantifying bacteria with spectrophotometry 1. Perform aserial dilution as shown in Figure 3. Do not change pipettes during these dilutions. They are each 1/2dilutions, so the margin of error from carryover inside the pipette issmall. 2. Use the spectrophotometers to measure the absorbance of light at 600nm wavelength to indicate bacterial growth. 3. Begin by using the “blank ”sample of sterile media. 4. Ensure that the cuvette lid issecure and vortex the cuvette. 5. Wipe the outside of the cuvette with aKimWipe to remove any smudges. 6. Place the cuvette in the spectrophotometer and indicate that this isyour reference sample (REF). Remove the blank cuvette. 7. Repeat steps 4and 5using dilution tubes containing bacteria. Place the cuvette in the spectrophotometer and record the absorption value. The more bacteria are in the sample, the higher this number will be. 8. Periodically place the blank back into the spectrophotometer. As long as itcontinues to read approximately 0.00, itisnot necessary to re-blank the machine. BIOL2902L Fall 2020 36 Dilution 1/2 1/4 1/8 1/16 1/6 1/12 1/24 Figure 3: Workflow for preparing cuvettes to quantify bacteria using spectrophotometry. EXPERIMENT: Quantifying bacteria with pour plates 1. Perform aserial dilution as shown in Figure 4. Change pipettes between each dilution; 1/10isa sufficiently large step that carryover can be asignificant source of error. 2. Label eight empty Petri plates. 3. Starting with the lowest dilution tube, add 1mL of your eight lowest dilutions (10 -10 through 10 -3)to agar plates. 4. You will have 200mL tryptic soy agar (TSA) in the water bath. Add approximately 20mL per plate, or about one-third of the height of the plate. 5. Set the plates out to solidify. At the end of the lab, place them, inverted, in abucket in the incubator. BIOL2902L Fall 2020 37 Figure 4: Workflow for preparing cuvettes to quantify bacteria using agar pour plates. Itisessential to change pipets between each dilution. CLEAN UP 1. Remove all labels from all test tubes and place them in autoclave racks at the front of the lab. 2. Ensure that all agar pour plates are incubated inverted at 37 °C. Dispose of all other agar plates in the autoclave bags at the front of the lab. 3. Clean your bench with disinfectant and paper towel. 4. Remove your lab coat and put itin its plastic bag. 5. Wash your hands. BIOL2902L Fall 2020 38 Lab 7 : Bacterial Growth II This lab will be delivered online through Nexus as a“live ”Zoom meeting. An invite to join your lab section ’sZoom meeting will be sent to your Nexus email the day before your lab. The Zoom meeting will be scheduled during your regular lab time slot. The Zoom meeting will include apre-lab presentation of theory and experimental procedures for this lab, as well as an overview of any associated assignments. At the end of the Zoom meeting you will have the opportunity to ask questions and interact directly with your instructor. Bacterial growth When bacteria are transferred to agrowth medium under optimal conditions, they will rapidly reproduce. The dynamics of this growth can be charted as abacterial growth curve. There are four stages that are typically seen in abacterial growth curve. Figure 5: Bacterial growth curve. Lag phase When the cells are first introduced to the medium, they will need time to adjust to their new environment. Metabolism will increase, proteins will be manufactured and the cells will grow in size but no cell division will occur during this phase. Logarithmic (log) phase During this phase, the number of cells will grow exponentially under optimal conditions. As each cell undergoes binary fission, the population will double regularly until amaximum number of cells has been reached. This isalso known as the exponential growth phase. Stationary phase During this phase, cells are being produced but are also dying, resulting in anet stability of the total population. Some essential metabolites are becoming depleted from the media and toxic byproducts are beginning to accumulate. Decline phase BIOL2902L Fall 2020 39 During this phase, the bacteria are dying at arapid and uniform rate due to the accumulation of toxic metabolic byproducts and the continued depletion of nutrients. The decline of the population closely mirrors the growth of the logarithmic phase. The length of time required to double the concentration of bacteria iscalled the doubling time. The length of time required between instances of cell division iscalled the generation time. These terms are sometimes used interchangeably because, during exponential growth, the doubling time and the generation time are the same. However, these terms are not actually identical, as we can see from the stationary phase where cell division can occur but the total population isno longer increasing. Bacterial standard curve The bacterial standard curve isagraph that compares the absorbance of light, measured with a spectrophotometer, with the number of CFU of bacteria within asample. The absorption of light isused as an indicator that there issomething blocking that light (i.e. bacteria), so the more light isblocked, the more bacteria are present, the higher the Absorbance measurement. Once you have constructed a standard curve, you can use itto estimate the number of CFU at agiven OD 600 for your bacterial species. This curve isunique to the experimental conditions used. Ifthe experimental conditions change (e.g. organism, media, atmosphere), the curve isno longer valid. For most bacteria, moderate changes in temperature do not affect the standard curve. The term “curve ”isabit of amisnomer. For our purposes, the relationship between absorbance and CFU/mL isastraight line. We also know that when there are zero CFU/mL in the tube, the absorbance of that tube should also be zero (a blank sample). This means that one point on our line will be the origin (zero on both the X-axis and the Y-axis) of the graph. Bacterial standard plate count (SPC) In order to construct acurve, we need to have at least one additional data point where we know both the number of CFU/mL and the absorbance. We will begin by using the agar pour plate data to calculate the number of CFU/mL in the undiluted sample. On astandard-sized Petri plate, the ideal range of colonies isbetween 30 and 300. Ifthe plate has more than 300 colonies on it, chances are that one of the colonies might actually be two different colonies that have grown so close together as to be indistinguishable or uncountable. Ifthe plate has fewer than 30 colonies, this count isunacceptable for statistical reasons. Ifthe serial dilution has been performed properly, there should be aten-fold difference in the number of colonies on each plate between dilutions. This also means that you are likely to only have one plate with between 30 and 300 colonies. Ifyou divide this number by the dilution for the plate, you can determine the number of CFU in the original sample. We have chosen to use 1mL samples for all of our plates to simplify this calculation; ifyou use adifferent volume, you must also divide by the volume used per plate to find the original concentration per mL. colony count on agar plate dilution of sample × volume of sample plated (1mL) =CFU per mL undiluted sample Workflow The following experiment was performed for you and adata set has been uploaded to Nexus for you to analyze. Although you did not perform the experiments yourselves, you are still required to understand the methods used and the theory behind the background information presented. BIOL2902L Fall 2020 40 Table 3: Quantification of bacteria through agar pour plates. You do not need to fill in the entire table; start with the smallest concentration (10 -10)and count subsequent plates until you find one with 30-300 colonies. You do not need to count any dilutions after that. Dilution CFU/mL (counted) CFU/mL of undiluted sample 10 -10 10 -9 10 -8 10 -7 10 -6 10 -5 10 -4 10 -3 Once you have determined the bacterial concentration of the undiluted sample (CFU/mL), you can combine this information with the absorbance data you acquired through spectrophotometry. The most concentrated dilution that you put into the spectrophotometer was ½ (50% concentration). This means that the number of CFU/mL to produce that absorbance reading should be half of the bacterial concentration that you calculated for the undiluted sample. You can repeat this calculation for each of the dilutions that were put in the spectrophotometer. Table 4: Quantification of bacteria through spectrophotometry. Use the absorbance values from the previous lab. The undiluted CFU/mL (calculated) comes from the dilution in Table 2with 30-300 colonies. Dilution Absorbance (measured OD 600nm ) CFU/mL (calculated) Undiluted 1/2 1/4 1/6 1/8 1/12 1/16 1/24 Now you can graph the results of this table. Put absorbance (OD 600nm )on one axis and concentration (CFU/mL) on the other. Knowing that when the absorbance iszero, the concentration must also be zero, make aline of best fit through your data points. You can now use this graph to convert between absorbance and concentration for this bacterial species growing under these conditions on this media. So, for example, ifyou were aresearcher who needed to start an experiment with abacterial concentration of 10 5CFU/mL, you could dilute your stock culture until itreached the OD 600nm absorbance that you know approximates that concentration. BIOL2902L Fall 2020 41 Bacterial growth curve data A culture of E. coli was split into two side-arm flasks and grown at either 20 °C or 37 °C. Every twenty minutes, the absorbance of light at 600nm wavelength was measured in both flasks and recorded in the table below. Using this data and the bacterial standard curve for E. coli based on your experimental results calculated above, prepare abacterial growth curve. This should be graphed on semi-log paper with the Y-axis scaled logarithmically. The growth curves for both temperatures should appear on the same graph paper. Calculate the generation time for E. coli at each temperature. Table 5: The change in concentration of colony forming units per mL of E. coli in two identical sidearm flasks measured through absorbance of light at 600nm wavelength over time at incubation temperatures of 20 °C or 37 °C. Concentration can be calculated using the standard curve for E. coli. Time (minutes) Absorbance (OD 600 nm)at 20 °C Concentration (CFU/mL) at 20 °C Absorbance (OD 600 nm)at 37 °C Concentration (CFU/mL) at 37 °C 0 0.030 0.030 20 0.031 0.031 40 0.036 0.046 60 0.052 0.071 80 0.073 0.111 100 0.103 0.173 120 0.138 0.207 140 0.181 0.223 160 0.200 0.229 180 0.203 0.232 BIOL2902L Fall 2020 42 Lab 8 : Bacterial Growth III This lab will be delivered online through Nexus as a“live ”Zoom meeting. An invite to join your lab section ’sZoom meeting will be sent to your Nexus email the day before your lab. The Zoom meeting will be scheduled during your regular lab time slot. The Zoom meeting will include apre-lab presentation of theory and experimental procedures for this lab, as well as an overview of any associated assignments. At the end of the Zoom meeting you will have the opportunity to ask questions and interact directly with your instructor. Environments can significantly reduce or completely inhibit the growth of some bacterial species while having minimal impact on other species. Two examples that we will look at are the effects of antibiotics and the effects of temperature. These experiments have been performed for you and are on display in the lab, so you can observe and record the results. Organisms can be classified as psychrophiles, mesophiles or thermophiles based on their optimal temperature for growth and replication. Psychrophiles can grow at 0-5 °C and grow optimally at -5°C to 20 °C. Mesophiles usually grow between 20 °C and 45 °C; by definition, they can grow at 37 °C and will not grow above 45 °C. Mesophiles that grow optimally at 35 °C to 40 °C usually grow in the bodies of warm-blooded organisms. A distinction can be made between facultative thermophiles, which can grow at 37 °C and grow optimally at 45 °C to 60 °C, and obligate thermophiles, which can only grow above 50 °C and grow optimally above 60 °C. Workflow The following experiment was performed for you and adata set has been uploaded to Nexus for you to analyze. Although you did not perform the experiments yourselves, you are still required to understand the methods used and the theory behind the background information presented. EXPERIMENT: Effects of temperature on bacteria An agar plate was streaked so that four different species of bacteria (Bacillus stearothermophilus, Escherichia coli ,Pseudomonas fluorescens and Serratia marcescens )were in four separate quadrants. This procedure was repeated to produce four identical plates, each with four species on the TSA plate. One plate was incubated at 4°C, one at 20 °C, one at 37 °C and one at 60 °C. By comparing these plates, itispossible to estimate the temperature ranges of these four species. 1. Demonstration plates have been provided for you. Qualitatively estimate the level of growth in each quadrant for each species and record your observations in Table 6. 2. Based on your observations, classify each species as apsychrophile, mesophile, facultative thermophile or obligate thermophile. Table 6: Qualitative levels of bacterial growth after incubation at different temperatures on TSA. Bacterial growth was classified as no growth (−),slight growth (+), moderate growth (++) or abundant growth (+++). Organism 4°C 20 °C 37 °C 60 °C Bacillus stearothermophilus Escherichia coli Pseudomonas fluorescens Serratia marcescens BIOL2902L Fall 2020 43 EXPERIMENT: Effects of antibiotics on bacteria The effects of an antibiotic on bacteria are assessed by placing apaper disk impregnated with antibiotic onto an agar plate that has been inoculated with bacteria. Ifthe antibiotic isable to inhibit the growth of the organism, this will be visible as aclearing or zone of inhibition around the disk. Each disk is6mm in diameter, so the size of the zone of inhibition can be quantified by measuring the diameter of the clearing in which there isno bacterial growth. This size isthen compared to aset of known standards to classify the organism as susceptible, intermediate or resistant to the antibiotic. Ifthe growth comes all the way up to the edge of the antibiotic disk, record this as zone of inhibition as 0mm. This test isknown as the Kirby-Bauer antibiotic susceptibility assay .Itisperformed on an enriched medium called Mueller-Hinton agar with abacterial OD 600 of between 0.08 and 0.1. 1. Demonstration plates have been provided for you. Measure the diameter of the area around the antibiotic disk in which there isno visible bacterial growth and record this measurement, in millimeters, in Table 7. 2. Using Table 8, determine ifthe organisms are sensitive, intermediate or resistant to each of the six antibiotics. Table 7: Zone of inhibition diameters observed during Kirby-Bauer antibiotic sensitivity testing, recorded in millimetres.

Organism Chloram- phenicol (C 30) Gentamicin (GM 10) Penicillin (P10) Streptomycin (S10) Tetracycline (Te 30) Vancomycin (Va 30) E. coli P. fluorescens E. faecalis Table 8: Zone diameter interpretive standards. Adapted from Clinical and Laboratory Standards Institute Performance Standards for Antimicrobial Disk Tests, Tenth Edition, 2008. Disk Antibiotic Zone diameter (mm) Resistant Intermediate Sensitive C30 Chloramphenicol ≤12 13 -17 ≥18 GM 10 Gentamicin ≤12 13 -14 ≥15 P10 Penicillin ≤14 — ≥15 P10 Penicillin for E. faecalis ≤28 — ≥29 S10 Streptomycin ≤11 12 -14 ≥15 Te 30 Tetracycline ≤14 15 -18 ≥19 Va 30 Vancomycin for E. faecalis — — ≥15 BIOL2902L Fall 2020 44 Epidemiology Communicable disease are caused by disease-producing microorganism called pathogens. These pathogens can be spread, either directly or indirectly, from one host to another. Some microorganisms cause disease only ifthe body isweakened or ifthe predisposing event, such as awound, allows them to enter the body. Non-communicable diseases tend to be chronic conditions, such as cardiovascular diseases (like heart attack and stroke), chronic respiratory diseases (like asthma), diabetes and cancer. The science that deals with when and where diseases occur and how they are transmitted in the human population iscalled epidemiology. Sporadic diseases are those that only occur occasionally in a population; an example isthe current outbreak of meningitis. Endemic diseases, such as pneumonia or influenza, are constantly present in the population. When many people in agiven area acquire the disease in arelatively short period of time, itisreferred to as an epidemic disease. Influenza often achieves epidemic status. Infectious or communicable diseases can often be transmitted by direct contact between hosts. Droplet infection, when microorganisms are carried on liquid drops from acough or sneeze, isamethod of direct contact. Disease-causing pathogens can also be transmitted by contact with contaminated inanimate objects, or fomites. Drinking glasses, bedding and towels are all examples of fomites that can be contaminated with pathogens from faeces, sputum or pus. Some disease-causing pathogens are transmitted from one host to another by vectors. Vectors are living organisms that transmit an infectious pathogen to another organism. In mechanical transmission, the vector transmits the pathogen without ever being infected by it. An example of mechanical transmission istyphoid fever, which can be transmitted on the feet of insects and the pathogen can be transferred to aperson ’sfood. Transmission of apathogen by an infected vector iscalled biological transmission. An arthropod such as amosquito can ingest apathogen while biting an infected host. The pathogen can multiply or mature in the arthropod and then be transmitted to anew, healthy host in the arthropod ’ssaliva or faeces. Examples of biological transmissions are malaria, Lyme disease and West Nile fever. The continual source of an infection iscalled the reservoir. The reservoir isoften agroup or species that can be regularly or continuously infected with minimal symptoms; Humans who harbor pathogens but do not exhibit any signs of the disease are called carriers. An epidemiologist compiles data on the incidence of adisease and its method of transmission and tries to locate the source of the infection in order to decrease the incidence. Epidemiology scenario Thirty-two people attended abrunch (11:00 to 15:00) on Sunday, May 22. Fourteen of the individuals developed symptoms of food poisoning after ingesting various food items. All fresh food items were delivered to the cooking area at the morning of May 22. The menu consisted of omelettes, bacon, fruit salad, and pastries. Omelettes were prepared from Grade A eggs from alocal farm. They were beaten, mixed with milk, salt, pepper, and diced onions. The egg batter was fried in small batches and served from warming trays heated by alcohol burners. Bacon was purchased from ameat wholesaler. The bacon was fried on alarge griddle and served from warming trays. Fruits salad was made fresh from apples, bananas, and oranges; the fruit was cut up and mixed with commercially prepared peaches and pears. Muffins and other breads were purchased from abakery and served with butter. What isthe most likely cause(s) of the outbreak? Are there any outliers (individuals who also become sick but who are unrelated to the outbreak)? BIOL2902L Fall 2020 45 Table 9: Epidemiological data collected for thirty-two people attending abrunch, some of whom developed symptoms of food poisoning. Foods and beverages consumed, and symptoms developed, are indicated by .Foods listed are: O, omelette; B, bacon; F, fruit salad; P, pastry. Beverages listed are: C, coffee; T,tea; M, milk; J, orange juice. Symptoms listed are: D, diarrhoea; N, nausea; V, vomiting; A, abdominal cramps. Case Foods Beverages Time of meal Symptoms Onset of symptoms O B F P C T M J D N V A Day Time 1        11:00     Sun 23:00 2     12:00 3      12:00   Mon 02:00 4      12:00    Mon 06:00 5   11:00 6     12:00 7     13:00 8    13:00 9      11:00     Mon 11:00 10   12:00 11   13:00 12      13:00    Mon 14:00 13   12:00 14      15:00 15     15:00    Tue 01:00 16    11:00 17     11:00 18    12:00  Sun 13:00 19    12:00     Mon 17:00 20     12:00 21     13:00    Mon 02:00 22   14:00  Tue 08:00 23    14:00 24      11:00   Mon 15:00 25     11:00    Mon 11:00 26    12:00 27   11:00 28      12:00   Mon 02:00 29    13:00 30   14:00 31    11:00 32     12:00   Mon 12:00 CLEAN UP 1. Clean your bench with disinfectant and paper towel. 2. Remove your lab coat and put itin its plastic bag. 3. Wash your hands. BIOL2902L Fall 2020 46 Review The review will be delivered online through Nexus as a“live ”Zoom meeting. An invite to join your lab section ’sZoom meeting will be sent to your Nexus email the day before your lab. The Zoom meeting will be scheduled during your regular lab time slot. The Zoom meeting will include areview presentation of theory and experimental procedures covered throughout the term lab. At the end of the Zoom meeting you will have the opportunity to ask questions and interact directly with your instructor. Lab Exam The lab exam will be delivered online through Nexus during the week of November 23 rd.Students will write their exam on their regular lab day (Wednesday the 25 th,Thursday the 26 th,or Friday the 27 th).The exam will open at 9:00 am and close at 11:59 pm each day. Itwill be worth 16 % of your final mark in the course.

The lab exam will consist of practical and theoretical questions .The practical question swill ask you to perform calculations or analyze results .The theoretical question swill be based on information provided through the Nexus website or found in the The Laboratory Manual .Multiple question formats will be presented on the exam including; multiple choice, fill in the blank, true or false, and short answer.