Electronic Laboratory Notebook

Home , Bunsen burner, Inoculation loop, Inoculation needle

SPRING 2017

ELECTRONIC LABORATORY NOTEBOOK

MCB3023 L SECTION 078G

CAROLYNA GUILLEN MEHRSA RAZAVI

Table of Contents Exploring the Microbiome 2 The Art of Microscopy 4 Life in A Drop of Water 5 Streaking for Isolation 8 Micro-pipetting and Graphing 10 Re Streak for Isolation 13 The Gram Stain 14 Bacterial Growth Curve 16 Saliva Enumeration 18 Gram Stain of Unknown: Identification Begins 21 Inoculation of Selective/Differential Media 1 23 Inoculation of Selective/Differential Media 2 26 Inoculation of Selective/Differential Media 3 28 Inoculation of Selective/Differential Media 4 29 Indole Test 30 Oxidase Test 31 Catalase Test 32 DNA Isolation 33 Loading and Running Gel Electrophoresis 36 ______STD Infection Simulation Using ELISA 37 Test for MRSA Carrier 39 pGLO Transformation 40 Antibiogram Actvity 43 Real Life Dilutions 46 Clean Water Lab 48 Fecal Float 50

Exploring the Microbiome January 5, 2017

Purpose: The purpose of this activity is to observe the microbiome that exists in three different areas: the surrounding environment, the air and the human body. Procedure: 1. Label three separate TSA plates with the different sources from which microbial samples will be acquired (i.e air, environment, body). 2. The air sample will be acquired by leaving the labeled petri dish open on the benchtop until the end of the lab period. 3. To sample from the environment, take a sterile cotton swab and dip it in saline solution. Then, take the swab and rub it on an object from the environment. Take the swab and lightly streak the TSA media in the respective petri dish. Close the petri dish and set it aside. 4. Repeat Step 3 for the body sample. 5. Tape all petri dishes closed. 6. Incubate at 37 degrees Celsius for 5 days. Observe colonies after incubation period. Results Figure 1. The picture on the left depicts colonies from microbes in the air. As seen, the culture is mixed. There are colonies that are very small while others are larger. Nearly every colony is flat except for one small yellow one that is convex. The majority of colonies are pale yellow or gold. The margins around most of them are entire, with a few undulate or lobate. The black colony is filamentous and the larger ones are irregular.

Figure 2. The image on the left depicts colonies from a sample taken of the microbes on my inner left forearm. As shown, the majority of colonies are pale yellow or gold, a few orange ones, and they are circular. All of the colonies are flat with entire margins.

Figure 3. The plate on the left is depicting a colony of microbes isolated from the track pad of my laptop. The colony is round, flat and pale yellow with entire margins. The colony measured .6 cm. This colony will be streaked for isolation and analyzed for the unknown project.

The Art of Microscopy January 5, 2017

Purpose: To become familiar with the anatomy of the microscope and acquire certain microscopy skills such as focusing and using oil immersion. Procedure: 1. Acquire a microscope from the cabinet, as well as a prepared microscope slide. 2. Load the slide onto the microscope and secure it. 3. Observe the specimen using the 4X objective lens first. To focus, use the coarse focus knob. Once focus is attained, move up to the 10X lens. Again, focus the image using th coarse focus knob. Once focus is attained, move up to th 40X lens. Only use the fine focus knob now. 4. Once focus is attained on the 40X lens, ask an Undergraduate TA to assist in adding the oil for the oil immersion step. 5. After the immersion oil is added, slowly turn the rotating piece to the 100X objective lens. Use the fine focus knob to acquire a clear and detailed picture.

Life in a Drop of Water January 5, 2017

Purpose: To practice creating wet mounts and, to observe and characterize various eukaryotic microbes in water. Procedure: 1. A wet mount was created using a water sample from Lake Alice. 2. The wet mount was observed under the microscope and organisms were identified. 3. Another wet mount was created using a water sample from the sewage system. 4. This wet mount was also observed under the microscope. Several organisms were identified. Results: Pictures of organisms found in the Lake Alice wet mount.

Figure 4. Bryozoa

Figure 5. Cyanobacteria Pictures of organisms observed in the sewage water wet mount.

Figure 6. Hydra

Figure 7. Rotifer

Figure 8. Gastrotrich

Streaking for Isolation January 10, 2017

Purpose: To isolate a pure culture in order to: (1) polish laboratory techniques and (2) have a pure culture for the Unknown Project. Procedure: 1. Decide which colony from which isolate is going to be cultured. The colonies are coming from the environmental samples plated in the lab on January 5, 2017. 2. Label the bottom of a fresh TSA plate with pertinent information (initials, date, isolate sample). 3. Draw a T on the bottom of the TSA plate (as shown below). This will make the quadrants where the bacteria will be streaked. 4. Light a Bunsen burner, establishing a steady, contained flame. Be sure to work close to the Bunsen burner in order to decrease the likelihood of contamination of the plate. 5. Grab the inoculation loop and hold it in the flame until the loop burns bright orange. This is to disinfect the loop before you begin streaking. 6. After disinfecting the loop, cool the loop on the EDGE of the agar in the new TSA plate. This way the bacteria will not be fried by the heat. 7. Take the inoculation loop and LIGHTLY scoop a very small amount of the desired colony. 8. (The first streak pattern will be made in the top quadrant of the TSA plate). LIGHTLY drag the inoculation loop over the agar of the new plate creating a zig-zag (depicted below). 9. Disinfect the inoculation loop in the flame. Cool it on the edge of the agar. Now, starting in the bottom left quadrant, streak the agar and create a zig-zag pattern. Make sure that two of the streaks pass into the first quadrant that was streaked. 10. Disinfect the inoculation loop in the flame. Cool it on the edge of the agar. Now starting in the bottom right quadrant, streak the agar and create a zig-zag pattern. Make sure that two of the streaks pass into the bottom left quadrant previously streaked in Step 9. 11. The streak plate is now ready for incubation. Incubate the plate UPSIDE DOWN at 37 degrees Celsius for the next two days.

Figure 9. The image on the left depicts the pattern of streaking that is achieved when following the procedures above.

Results:

Figure 10. The image on the left depicts a streak plate of an unknown microbe, sampled from the track pad of my laptop. Isolated colonies were obtained. The colonies are also the same yellow color, shape and of similar size. It is likely that this is a pure culture.

Micro-pipetting and Graphing January 10, 2017

Purpose: To perform a serial dilution in order to establish precise and proper micro-pipetting skills. Procedure: 1. Acquire an ELISA strip and label the end without a notch with your seat number. 2. Using the 200 microliter pipette, fill each of the wells with 90 microliters of water. To DRAW UP liquid, push the plunger down to the FIRST stop and enter the tip into the water. Slowly release the plunger to draw up the liquid. DISPENSE the liquid into the wells making sure to go down to the SECOND stop. This will ensure that every drop of liquid is expelled. Note: NEVER use the pipette without a tip! 3. Using the 10 microliter pipette, fill the FIRST well with 10 microliters of crystal violet dye. Aspirate the well by slowly pressing the plunger up/down. 4. Dispose of the tip. Draw up 10 microliters from the FIRST well and dispense it into the THIRD well. Dispose of the pipette tip. 5. Using a new pipette tip, aspirate the THIRD well and transfer 10 microliters of solution into the FIFTH well. Dispose of the tip. Aspirate. Leave well SEVEN blank. This will serve as a control. 6. Now, to create replicates of the dilutions performed, transfer 10 microliters of crystal violet into the SECOND well. Aspirate and dispose of the tip. Transfer 10 microliters of the solution to well FOUR. 7. Dispose of the tip. Aspirate. Transfer 10 microliters of the solution to well SIX. Aspirate and dispose of the tip. Leave well EIGHT blank. This will serve as another control. The ELISA strip should appear as depicted on the following page.

Figure 11. This photo was added for extra clarity of the procedure

Results:

Figure 12. This figure depicts the completed ELISA strip with all dilutions.

Table 2. Optical density values for each dilution measured at 600nm using a microplate reader. Dilution Optical Density at 600nm 0.01 0.899 0.01 1.138 0.001 0.275 0.001 0.275 Control 0.038 Control 0.038

Average Class OD600 1.8

1.6

1.4

1.2

0.8

0.6

0.4 Optical Densityat 600nm (O600) 600nm Densityat Optical 0.2

0 0.01 0.01 0.001 0.001 control control Dilution

Series1 Series2

Figure 13. Line series 1 represents average optical density measured by ELISA plate reader of students’ serial dilutions of crystal violet. Error bars reflect standard deviation of all optical density values measured at specified dilution. Line series 2 represents the optical density measured by ELISA plate reader of my own serial dilutions of crystal violet. Error bars reflect the standard deviation of my optical density values, along with those of the class.

Re-Streak for Isolation January 12, 2017

Purpose: To create a sub-culture of microbes from the initial streak plate conducted on January 10, 2017, and to provide the microbes with fresh media and nutrients. Procedure: 1. Use the streaking procedures outlined on page 8 to using a sample from the streak plate prepared on January 10, 2017. Results: Figure 14. Below are the results of the streak conducted on unknown microbes on January 10, 2017.

The Gram Stain January 12, 2017

Purpose: To practice conducting the gram-stain procedure as one of the first steps in characterizing bacteria. Procedures: 1. Grab a blank microscope slide and place a VERY SMALL drop of water in the middle of the slide. 2. Using a petri dish of sample bacteria, take a small sample of a colony with an inoculating loop and mix it with the drop of water on the slide. Spread the sample well to facilitate faster drying. Note: Remember to STERILIZE THE INOCULATING LOOP using the Bunsen burner. 3. Let the sample on the slide COMPLETELY air dry before proceeding. 4. Once the sample is completely dry, hold the microscope using a clothes pin and heat fix the sample by passing the slide through the Bunsen burner flame 3x. Note: Do NOT hold the slide in the flame because the glass will break. 5. Now you are ready to gram stain. For the first step, flood the slide with crystal violet (primary stain). Let the dye sit for 1 minute. Thoroughly rinse the slide with water. 6. Flood the slide with iodine (mordant) and let it sit for 1 minute. Thoroughly rinse the slide with water. 7. Flood the slide with ethanol (decolorizer). Let it sit for 10 SECONDS and then rinse very thoroughly. Note: Make sure to have the water on hand to rinse the slide quickly. If the alcohol sits for too long, the sample will be completely decolorized and the gram stain will fail. 8. Flood the slide with safranin (counterstain) and let it sit for 1 minute. Rinse the slide thoroughly. BLOT the slide with bibulous paper to dry it. 9. Observe the slide under the microscope using oil immersion. Results:

Figure 15. Gram stain of E. coli. As shown, the bacteria stained pink/red, meaning the bacteria is gram negative.

Figure 16. Gram stain of Staphylococcus aureus. As shown, the bacteria stained purple meaning the bacteria is gram positive. Clusters of cells can also be seen, confirming that it is Staphylococcus.

Bacterial Growth Curve Experiment January 17, 2017

Purpose: To observe and analyze bacterial growth in different mediums and, learn how to calculate generation time. Procedure: 1. For the treatment, Biotene mouthwash was used. 2. Solutions of BHI, and/or bacteria, and/or Biotene (treatment) were mixed in each of the wells. The volumes of each component for each well are listed in the table below.

3. Once each of the wells were prepared, the strips were placed in a microplate and read at OD600 every 20 minutes for 18 hours.

Figure 17. The image to the left demonstrates what the 8-well ELISA strip will look like after all dilutions are prepared.

Figure 18. The image to the left shows a microplate loaded with 8- well ELISA strips. This is how the strips are loaded before the samples can be read by the microplate reader.

Results: Unfortunately the class data had to be discarded. Sample data was used to calculate growth curves and depict data graphically.

Saliva Enumeration January 17, 2017

Purpose: To learn how to enumerate bacteria in a sample using laboratory techniques. Procedures: A. Making Your Saliva Dilutions 1. Initial and date the petrifilms. Label petrifilms with the following dilutions: 10-4, 10-5, 10-6, 10-7, and 10-8 and “control”. 2. Label 10 Eppendorf tubes with the following labels/dilutions: saliva, 10-1, 10-2, 10-3, 10-4, 10-5, 10-6, 10-7, 10-8 and “control”. 3. Fill all of the Eppendorf tubes (EXCEPT the saliva tube) with 1080 microliters of water using a pipette. Note: You CANNOT pipette 1080 microliters with the P1000 pipette. You will have to pipette 2 x 540 microliters. 4. Collect some of your spit (saliva) in the Eppendorf tube labeled appropriately. Ideally, the saliva sample should be about 200 microliters. This is your undiluted sample stock. 5. Conduct a 1:10 serial dilution: Gently vortex the saliva and aseptically transfer 120 microliters of the saliva to the tube labeled 10-1. This is a 1/10 or 10-1 dilution. Note: Saliva is very viscous. Be mindful of this while pipetting. It takes the full volume of saliva a while to be drawn up into the pipette dip and to be completely expelled from the pipette tip. 6. Close the tube and vortex the tube containing the 10-1 dilution. 7. Aseptically transfer 120 microliters of the 10-1 dilution to the next tube, labeled 10-2. This is a 1/100 or 10-2 dilution. Vortex. 8. Using the same pipette tip, repeat this process until you complete the dilution series. Note: Remember to vortex the solution before transferring to the next tube. 9. Do NOT add any saliva/dilution to the tube labeled “control”. Tip: If you forget how to conduct a serial dilution, or need a depiction for more clarity, reference page 10. B. Plating the Saliva Dilutions on Petrifilms

Note: PAY CLOSE ATTENTION TO THE PROCEDURES! Only 5 of the prepared dilutions will be plated on the petrifilms.

1. After the last tube has been diluted, start with the 10-8 dilution tube. Vortex the solution. 2. Place the petrifilm on a flat, level surface. 3. Lift the top film and with the pipette perpendicular to the petrifilm, dispense 1 milliliter (1000 microliters) of the 10-8 dilution onto the center of the bottom film (the one with the grid on it).

4. Drop the top film down onto the sample to prevent bubbles, tusing the spreader to distribute the liquid. Tip: When using the spreader, it helps to LIGHTLY lay it on the sample and tap the sample with the spreader, trying to make an even spread and preventing the sample from overflowing through the sides of the petrifilm. 5. Continue the same process with the petrifilm labeled 10-7. Gently spread out the liquid after you close the petrifilm. 6. Then inoculate the remaining petrifilms, moving from highest dilution to lower dilution, transferring 1 milliliter to the 10-6, 10-5 and 10-4 dilution tubes. You can use the same tip for the procedure. 7. Make sure you inoculate one petri film “control” with your diluent (water) ONLY. 8. Stack the petrifilms, tape the SIDES (not the flap) with 2 separate strips of tape and incubate.

Results:

Figure 19. The image above depicts the colonies formed after plating the saliva and incubating the petrifilms for ~2 days.

Table 3. Saliva enumeration of dilutions conducted. Dilution # of colonies counted CFU/mL Control contaminated N/A 10-4 299 2.99 x 106 10-5 162 1.62 x 107 10-6 129 1.29 x 108 10-7 132 1.32 x 109 10-8 176 1.76 x 1010

CFU/mL = # of colonies counted x dilution factor Dilution factor = 1/total dilution

Gram Stain of Unknown: Identification Begins January 19, 201 Purpose: To carry out the first step of the unknown microbe project by conducting a gram stain to characterize its cellular morphology and arrangement. Procedure: 1. A gram stain was conducted using a pure culture of an unknown environmental microbe streaked on January 17, 2017. Procedures for conducting a Gram stain are outlined on page 13. Results:

Figure 20. Two Gram stains was conducted on a pure culture of an unknown environmental microbe. These are the two resulting microscope slides.

Figure 21. The image to the left is an image of the first Gram stain conducted on a pure culture of unknown microbial microbes. As shown in the image, the microbe belongs to the Staphylococcus genus. This can be deduced by the spherical morphology and cluster arrangement of the cells.

Figure 22. The image to the left depicts an image taken of the second Gram stain performed on an unknown environmental microbe. This image depicts the same as the image above. The cells are spherical and arranged in clusters. Because of these observations, it is safe to conclude the microbe belongs to the Staphylococcus genus.

Inoculation of Selective/Differential Media 1 January 24, 2017

Purpose: To use traditional laboratory culturing techniques in order to begin characterizing an unknown environmental microbe. Procedure: 1. Using a pure culture of an unknown microbe, inoculate the following media plates: Blood Agar (BA), MacConkey Agar (MAC), DNAse, and Mannitol Salt Agar (MSA). Label each plate with the date, your initials, media type and isolate source. 2. To inoculate each plate, use a new sterile cotton swab each time. Use the cotton tip of the swab and gently rub the colonies or streaks of the pure culture. Then, lightly drag the swab across the media plates in a zig zag motion. Be sure to discard the swab in biohazard and the paper wrapper in the trash. Note: Remember to inoculate while keeping an open flame. This will prevent contamination. Result:

Figure 23. No growth was observed on the MacConkey Agar. MAC inhibits the growth of Gram positive bacteria. This concurs with the results attained from the Gram stain.

Figure 24. Growth was observed on the Mannitol Salt Agar (MSA). The media also changed color from, red to pink. MSA is selective for Staphylococcus spp. differential for pathogenic vs. non-pathogenic strains of Staphylococcus. Growth on MSA indicates the unknown microbe is a member of the Staphylococcus genus. The color change observed indicates the microbe produced alkaline by-products.

Figure 25. Growth was observed. Alpha hemolysis was observed. BA is used to test for the production of hemolysins by the cultured microbe. The unknown microbe grew on the media and managed to exhibit alpha hemolysis – incomplete lysis of red blood cells.

Figure 26. Growth was observed along with the clearing of the media. Growth on DNAse media indicates the microbe has the ability to hydrolyze DNA for uptake. The unknown microbe exhibits this property.

Inoculation of Selective/Differential Media 2 January 24, 2017

Purpose: To use traditional laboratory culturing techniques in order to begin characterizing an unknown environmental microbe. Procedure: 1. Each row of students will have 1 TSA plate to work with. Divide the plate into 6 slices. Label the tip of a slice with your initials. Write the date once around the perimeter of the plate. 2. Using a sterile cotton swab, sample some of your pure culture and inoculate YOUR slice only. Note: Keep an open flame while inoculating to prevent contamination. 3. Once all media plates have been inoculated, stack the plates in an inverted fashion. Then, tape the sides of the plates together. Plates will be placed in an anaerobe jar with a test strip and an anaerobe pack. These plates will be tested for oxygen requirements. Results:

Figure 27. These are the results of all six students in the row inoculating a TSA plate with their unknown microbes.

Figure 28. This is a close up of the TSA plate inoculated by the entire row of students. The wedge labeled “C.G.” is the wedge inoculated with my unknown environmental microbe. Isolated colonies can be seen. The colonies are all similar in size, shape and color.

Inoculation of Selective/Differential Media 3 January 24, 2017

Purpose: To use traditional laboratory culturing techniques in order to begin characterizing an unknown environmental microbe. Procedure: 1. You should have a tube of phenol red broth with glucose and a Durham fermentation tube – which is the small inverted tube suspended in the broth. Note: You must be INCREDIBLY GENTLE with the tubes! Durham tubes are used to test for the production of gas which is indicated by the formation of a bubble. If you are shaking/moving around the test tube too much, you can incorporate air into it yielding a false positive. 2. Flame your inoculation loop, then allow it to cool in the zone of asepsis. With your sterile loop, pick up some bacterial growth from your streak plate. Then inoculate the glucose/phenol red broth by opening the tube, putting the loop in, and agitating it gently so the growth from the loop becomes suspended in the liquid. Note: Be careful not to hit the Durham tube with the loop. Results:

Figure 29. Color change from red to yellow – glucose fermenter. No gas production. Phenol red broth was used to test for this property. Glucose fermentation will induce a change in the broth from red to yellow. Gas production is indicated by a bubble in the Durham tube placed in the broth. The unknown microbe induced a change in the broth indicating glucose fermentation. However, no gas was produced upon fermentation, indicated by the absence of a bubble in the Durham tube.

Inoculation of Selective/Differential Media 4 January 24, 2017

Purpose: To use traditional laboratory culturing techniques in order to begin characterizing an unknown environmental microbe. Procedure: 1. Sterilize your inoculation needle. Allow it to cool, then pick up some bacterial growth from your streak plate. 2. Stab into the agar, about ¾ of the way to the bottom. Note: Make it a CLEAN VERTICAL LINE. Do NOT gouge the SIM medium by twisting the loop while it is in the tube. Results:

Figure 30. No motility was observed. No hydrogen sulfide was produced. Presence of indole was negative. Because no lateral movement was observed, the microbe is not likely motile. The production of hydrogen sulfide is indicated by the presence of a black precipitate. No black precipitate was observed. Kovac’s reagent was applied to test for the presence of indole (indicated by color change from yellow to red upon the addition of Kovac’s). No color change was seen.

Indole Test Procedure January 26, 2017

Purpose: To test for the production of tryptophanase by our unknown microbe. Procedure: 1. Take container of +Kovac's reagent and dispense 1 to 2 drops of the reagent on top of the SIM medium. 2. Observe the liquid on the agar surface to turn bright red IMMEDIATELY. The appearance of a red color after the addition of 2 drops of Kovac’s Reagent to the SIM tube within 15 sec indicates a positive reaction for indole production. A yellow color is indicative of a negative reaction. Results:

Figure 31. Kovac’s reagent was applied to test for the presence of indole (indicated by color change from yellow to red upon the addition of Kovac’s). No color change was seen.

Oxidase Reaction January 26, 2017

Purpose: To test for oxidase production by the unknown microbe. Procedure: 1. Touch the tip of a sterile swab to the top of a colony, from an actively growing culture, on the surface of the agar medium and gently rub. Preferably use growth from the BA or DNase media plates. 2. Dispense 1 to 2 drops of Oxidase Reagent onto the tip of the cotton swab. 3. Observe the cotton swab for the development of a purple color within 30 seconds. 4. After 30 seconds this is considered to be a false positive. Results:

Figure 32. The oxidase test is used to test for the presence of cytochrome C – a member of the electron transport chain that passes electrons to oxygen as the final electron acceptor. If cytochrome C is absent, a negative result is obtained. Thus the unknown microbe does not use this particular cytochrome but it may use others. This also means that oxygen may not be the final electron acceptor when this microbe is respiring.

Catalase Reaction January 26, 2017

Purpose: To test for the presence of the catalase enzyme in our unknown microbe. Procedure: 1. Obtain a pure culture of the organism to be tested. 2. Using a sterile cotton swab, pick a well-isolated colony and transfer to a glass slide. Do NOT use the metal loop. 3. Add 1 or 2 drops of the Catalase Reagent (H2O2) to the smear. 4. Examine immediately for the rapid production of gas bubbles. Results: Bubbles were produced – positive for catalase. Production of the catalase of enzyme is measured by production of bubbles upon addition of hydrogen peroxide to a sample of the microbe. Aerobic organisms produce catalase in order to hydrolyze toxic byproducts. Bubbles were produced by the unknown microbe, though not many. This means that the microbe is oxygen-tolerant.

DNA Isolation January 31, 2017

Purpose: To isolate cellular DNA for amplification. Procedure: **ALWAYS LABEL YOUR TUBES** 1. Take a micro-centrifuge tube with 1mL 1x Sterile PBS buffer. 2. Scrape off half of the growth on the TSA plate of your pure culture streak for isolation using a sterile inoculating loop. Suspend the cells in the micro-centrifuge tube with 1mL 1X Sterile PBS buffer. 3. Vortex at max speed for 5 sec. 4. Centrifuge at 13, 000 rpm for 1 min. Look for a cell pellet. The DNA is in the cells inside the pellet. It should be of a substantial size to ensure that you’ll have enough DNA. The pellet should AT LEAST cover the bottom of the micro-centrifuge tube. 5. Remove the liquid supernatant and dispose of it in biohazard. Note: Use the 1000 uL pipette to remove the liquid. DO NOT just pour it off! Also, draw the liquid SLOWLY to avoid sucking up your pellet. 6. Add 0.5 mL 1x sterile PBS buffer to the pellet 7. Re-suspend the pellet by vortexing at max speed until the whole pellet is dissolved in the tube. 8. Place in a hot water bath at 85°C for 5 mins. 9. Transfer the cells to the vial of Chemglass beads. 10. Vortex for 5 min at max speed. 11. Centrifuge for 1 min at 13,000 rpm. 12. Your DNA is now in the supernatant. Transfer supernatant to a new tube & discard the pellet.

1. DNA Purification

a. Procedure: **NOTE: ALWAYS LABEL YOUR TUBES** 1. Transfer 150 μl of the DNA extraction supernatant to a new Eppendorf tube. 2. Add 400 μl of PowerClean DNA Solution 5 to the 150 μl of the DNA supernatant. Vortex at max speed for 5 seconds. 3. Load the 550μl of the mix onto a spin filter. Centrifuge at 13,000 rpm for 1 minute at room temperature. 4. Place the spin filter in a new Eppendorf tube (it is fine if it does not close). Discard the FLOW THROUGH, NOT the spin filter.

5. Add 500μl of PowerClean™ DNA Solution 6 and centrifuge again at 13,000 rpm for 30 seconds at room temperature. 6. Repeat step 4. 7. Centrifuge at 13000 rpm for 1 minute at room temperature. 8. Repeat step 4. 9. Add 50μl of PowerClean DNA Solution 7 to the center of the white filter membrane. 10. Centrifuge the spin filter at 13,000 rpm for 30 seconds at room temperature. 11. Discard the SPIN FILTER. The DNA is in the Eppendorf tube. It is now ready for PCR! 12. UTA’s quantified the DNA products for the class using a microplate reader prior to beginning PCR. This is done to quantify the amount of DNA extracted and is later used for calculations for PCR. PCR Calculation: Because the class data yielded extremely varying concentrations of DNA product, we did not follow through with the conventional calculations. Normally, 20 µL of PCR master mix is combined with a volume of DNA and water (if necessary) to arrive at a reaction mixture of 25 µL. The volume of DNA needed for the reaction mixture is derived using the C1V1=C2V2 equation, C1= the concentration of DNA in your sample obtained from the microplate reader, V1= 25 µL, C2- 250 ng/µL and V2= the volume of DNA that will be added to the 20 µL of PCR mix.

2. Polymerase Chain Reaction

a. Procedure:

1. Label your PCR microtube on the SIDE AND TOP clearly with your seat number! Be sure not to rub the µLoff with your fingers. 2. Go up to the ice bath and have one of the UTA’s pipette 20 µL of master mix directly into your PCR microtube. 3. Transfer 5 μL of your DNA to the PCR microtube containing the master mix. Flick the tube to mix. 4. Put the PCR tube in the PCR machine. Fill out the spreadsheet next to the PCR machine so your tube can be located in case the label is wiped off. 5. PCR was conducted by Maddie. 6. Data was sequenced by a third party on campus.

Results:

Figure 33. This is the sequence obtained for the 16s rRNA gene of the unknown environmental microbe.

As depicted, the sequence is short. Discrepancy in size could have been due to the fact that a very minute amount of DNA (9.89 ng/µL) was initially extracted. Because the sequence so short, qualitative results were not obtained when searching the BLAST database. The sequences only showed between 15-16 percent similarity. The results were also extraneous. Instead, the Gideon database results were used to definitively identify the unknown microbe as Staphylococcus epidermidis.

Loading and Running Gel Electrophoresis February 2, 2017

Purpose: To perform gel electrophoresis using purified DNA from our unknown microbe in order to identify it. Procedure: 1. Gels were poured for the class. 2. Maddie poured 1X TAE buffer into the electrophoresis trays. 3. Maddie loaded the DNA ladders for the class. 4. Prepare samples for the gel run. Take 0.5 μL of Loading Dye and mix with 2.5 μL of the PCR product. Aspirate the solution gently to mix. 5. Load your samples in the gel wells using the pipette. The density of the solution should help dispense the DNA into the well and prevent it from going everywhere. Note: Be careful and make sure the pipette tip is DIRECTLY ABOVE the well. DO NOT stab the gel. 6. Students loaded their samples and the UTA’s ran the gels for the class. Maddie imaged the results.

Results:

Figure 34. Electrophoresis results of PCR purification of the 16s rRNA gene of the unknown microbe in question. The red circle is used to denote which lane to observe. The picture on the right is a labeled image of the ladder used for electrophoresis. As depicted on the image of the ladder -- indicated by bolded text – the DNA products should fluoresce at about 3 kb. The product fluoresced at this position on the gel

STD Infection Simulation Using ELISA February 28, 2017

Purpose: To gain a better understanding for the importance of practicing safe sex by demonstrating the ease of and extent to which an STD can spread. Procedure: Indirect STD ELISA 1. Label an Eppendorf tube with “bodily fluid” with your initials. 2. Share an 8-well microplate strip with your partner (you use 4 wells, and your partner uses 4 wells). 3. Transfer 50 microliters out of the “bodily fluid” tube to your well 1. This will serve as your control since you have not exchanged bodily fluids with anyone. 4. Pair up with a random student in. This is your first sharing partner. *Note: Record who your sharing partners are and their correct order. 5. Transfer 700 microliters of your “bodily fluid” into your Sharing Partner 1’s tube. Aspirate gently. 6. Transfer half of this mixture (~700 microliters) back to your empty sample tube. 7. Transfer 50 microliters of this mixture to your well #2. This is your Sharing Partner #1 well. 8. Pair up with another student. This is your Sharing Partner 2. Repeat steps 5-7. You should end up filling another well and it being your Sharing Partner #2 well. 9. Pair up with a third student and repeat steps 5-7. This is your Sharing Partner #3. You should end up filling your last well. 10. TA’s prepared positive and negative control. 11. Wash 3x. a. Fill wells with 200 microliters wash buffer. b. Tip the strip over onto paper towels and gently tap the strip to expel the liquid. c. Add 200 microliters of wash buffer to each of the wells again. d. Tip the strip over onto CLEAN paper towels and tap gently. e. Repeat c and d. f. Throw away the towels. 12. Repeat the wash one more time. The wells should be empty now. 13. Add 50 microliters of primary antibody to each of the 8 wells. This will bind to the antigen. 14. Incubate at room temperature for 5 mins to allow the antibodies to bind to the antigen. 15. Wash away unbound antibodies by washing the wells twice (using Wash Buffer with Tween). 16. Add 50 microliters of the secondary antibody into each of the 8 wells. This antibody is conjugated to horseradish peroxidase. 17. Incubate at room temperature for 5 mins to allow the secondary antibodies to the primary antibodies. 18. Wash away the unbound antibodies by washing 3x with PBS WITHOUT Tween 19. The wells should be empty now.

20. Add 50 microliters of the substrate SUB to each of the 8 wells. This is the substrate for horseradish peroxidase. 21. Incubate at room temperature for 5 minutes to allow the reaction to occur. 22. Observe the results. Blue = positive result; clear = negative result. Which of your partners were infected?

Results: All of my sharing partners were infected. All 3 of my sharing partners exchanged bodily fluids with me and each other, basically infecting one another. The point of infection was the exchange with my Sharing Partner #1 who was originally infected.

Figure 35. STD ELISA results. Well 1-4 from left to right were my bodily fluid samples. Wells 2, 3, 4 fluoresced meaning that those are infected. All 3 of my sharing partners were infected.

Test for MRSA Carrier February 28, 2017

Purpose: To explore if anyone in the class was a MRSA carrier. MRSA is relevant because of its recent discovery and its dangerous pathogenicity. Procedures: 1. Light a Bunsen burner and be sure to work within the zone of asepsis. 2. Swab your upper nose with a sterile cotton swab. Insert the swab into the tip of one nostril and swab it for 30 seconds. Swab the other nostril for 30 seconds using the same swab. 3. Rub swab over half (because you are sharing the plate with a partner) of the MRSA Chromagar plate. 4. Incubate for 48 hours. If purple colonies are observed, you are a MRSA carrier.

Results: I am a MRSA carrier.

Figure 36. The left half of the plate is my sample. Purple colonies can be seen growing on the agar, which indicates a positive result.

39 pGlo Transformation March 14, 2017

Purpose: To learn how to perform a successful transformation Procedures: 1. Label one microcentrifuge tube +pGLO and the other –pGLO. 2. Open the tubes. Using a sterile pipette tip and aseptic technique, transfer 250 microliters of the transformation solution into the tube. *Note: You basically need to have your Bunsen burner on and be working in the zone of asepsis for the entire experiment. 3. Place the tubes on ice for 5 minutes. 4. Use a sterile PLASTIC inoculating loop to pick up a single colony from the E.coli plates provided. Immerse the loop in the transformation solution at the bottom of the tube labeled +pGLO. Make sure to dispense the entire colony in the transformation solution! Place the tube back on ice. 5. Using a NEW sterile plastic loop and aseptic technique, repeat this for the –pGLO tube. 6. Immerse a new plastic sterile loop into the plasmid DNA stock tube. Withdraw in a loopful – there should be a film of solution across the ring. The TAs will do this for you. Insert the loop into the cell suspension of the + pGLO tube and mix. Close the tube then return it to the ice. DO NOT do this for the –pGLO. 7. Incubate the tube on ice for 10 mins. Make sure to push the tubes ALL THE WAY down in the ice to make sure the solution is submerged. 8. While the tubes sit on ice, label the agar plates on the bottom as shown below.

9. Heat shock: Transfer both the + and - pGLO tubes onto the heating block and wait 50 seconds. Then put both tubes back on ice and incubate for 2 mins. 10. Add 250 microliters of LB nutrient broth to both tubes using a new, sterile pipette tip each time. 11. Let both tubes incubate for 10 mins at room temperature 12. Tap the closed tubes with your finger to mix. Using a new sterile pipette tip for each tube, add 100 microliters of the transformation and control suspensions onto the appropriate plates. 13. Use a new sterile “L” disposable spreader for each plate. Using the short part of the L, spread the suspensions evenly around the surface of the agar. 14. Stack your plates and tape them together. Label the tape with your initials and put it in the incubator until the next day.

Results:

Figure 37. A figure of all of the plates after incubation. All of the plates had growth but there is no indication – yet – that the transformation was successful.

Figure 38. The transformation was successful. The plate depicted to the left is the +pGLO plate with LB/amp/ara. This was the only plate that should have fluoresced.

The only plate to fluoresce was the +pGLO with LB/amp/ara, which makes sense because the sample was exposed to the plasmid that carried an antibiotic resistance gene for ampicillin and a GFP gene regulated the araC regulator. Ampicillin and arabinose were both substrates that were present in the agar which is how the microbe was able to grow on the medium and fluoresce.

Antibiogram Activity March 16, 2017

Purpose: To explore the effectiveness of various common antibiotics vs. other antimicrobial agents (i.e. acne cleanser, essential oils, detergents/soaps). Procedures: 1. Label the bottom of each Mueller-Hinton agar plate with your initials, date and test microbe. 2. Using aseptic technique, pipette 200 microliters of the bacterial suspension onto the plates. Then spread the liquid across the agar surface using the “L” spreaders (only use one per plate). Dispose of the spreaders in the biohazard. 3. Let the plates sit for ~30 seconds, allowing excess moisture to absorb. 4. Choose a plate to use for this first CLASSIC antibiogram. 5. Choose 5 different antibiotics. 6. Sterilize your tweezers by gently dipping the tip into the ethanol bottle and then passing it through the Bunsen burner. *Note: Make sure you hold the tweezers at a slant with the tips angled downward to avoid burns. 7. Gently remove an antibiotic disk from the cache. Using aseptic technique, place it onto the agar. Make sure they are evenly spaced out. Do not place them near the edge. 8. Sterilize your tweezers between each disk as before. DO NOT AGITATE THE PLATES. It can cause the disks to move. 9. Carefully tape plates in a stack RIGHT SIDE UP and place in metal bins. 10. To conduct the second ALTERNATIVE antibiogram, choose 5 antiseptics/disinfectants and make note of them. 11. Take a sterile empty filter disk with STERILE tweezers. 12. Using aseptic technique, dip the disk into the solution of choice. Try to shake off excess moisture. 13. Gently place the disk onto the agar plate you have not used. Be sure that you have recorded what substance you have used, and where it is located on the plate. 14. You can place up to 5 disks in a circle on your plate. Do not place them close to each other or near the edge of the agar. 15. Tape the plates in stacks RIGHT SIDE UP and place in metal bin for incubation.

Results:

Figure 39. The photo on the left depicts the alternative antibiogram. E. coli is shown to be susceptible to all agents except Windex. The other agents include Eucalyptus oil, Clean and Clear Acne Cleanser, Neutrogena Acne Cleanser and dish liquid.

Figure 40. The photo to the left depicts the classic antibiogram. Here E. coli is shown to be susceptible to ampicillin, tetracycline and vancomycin but resistant to oxacillin and trimethoprim sulfamethoxazole.

Real Life Dilutions (with Amanda) March 28, 2017

Purpose: To apply laboratory techniques we practiced throughout the course to food microbiology. Procedures: 1. Label each set of petri films with the appropriate labels denoted on the quantitative testing sheet. 2. Label all petri plates as instructed. 3. Emulsify the sample of strawberries by squishing and pitching the plastic bag. Do not be so rough that the bag bursts or contents are leaked. 4. Arrange petri films and petri plates from top to bottom on lab bench with the lowest dilution closest to you and the highest dilution farthest away from you. 5. Portion out your sample as instructed so you have enough to complete the dilutions necessary. 6. Dilute your samples and homogenize them. 7. Pipette your samples onto the appropriate films and plates. 8. Tape the plates and incubate them at 37 degrees Celsius for 2 days. Incubate petri films under appropriate conditions. Each petri film has different growing conditions.

Results:

Figure 41. This photo depicts some of the petri films we were able to observe that were positive for growth.

The RYM films and the petri dishes were not found. We only observed the APC films. Dilution one was too numerous to count and dilution 4 was too few to count. Dilution 2 had 249 colonies and dilution 3 had 58 colonies. These samples came from the strawberries used for the activity.

Clean Water Lab March 30, 2017

Purpose: To test the effectiveness of current water treatments. Procedures: 1. Label petri films at the top with initials, date, origin of water sample, treatment and dilution. 2. Mix part of your water sample with part of the dirty class water sample. 3. Make a 1:10 dilution of the sample by mixing 1 microliter of the contaminated water with 9 microliters of clean water in a falcon tube. Vortex the solution. 4. Dilute the water sample 1:10 to 106. 5. Plate 1mL of the 10-3, 10-4, 10-5, 10-6 dilutions onto an RYM, CC and Enterobacter film. You should end up with 12 films. 6. Filter 150 mL of your original contaminated sample through a sari cloth. a. Make the filter by folding the cloth over twice. Place the cloth in the funnel and make sure you push it well down into the funnel. Pour the water into the funnel and filter out 150 mL. 7. Dilute the filtered water sample 1:10 to 10-6. 8. Plate 1mL of the 10-3, 10-4, 10-5, 10-6 dilutions onto an RYM, CC and Enterobacter film. You should have 12 films. 9. Chemical treatment: Dissolve an iodine tablet in the filtered water. Do not dilute the sample. 10. Plate 1 mL onto an RYM, CC and Enterobacter film.

Results:

Figure 42. The photo to the left depicts the petri films after incubation. The purple films are the Enterobacter films and the red one is the CC film.

The top Enterobacter film was the 10-6 dilution of the sari filtrated water. This film had 2.56 x 10-4 CFU/mL. The bottom Enterobacter film was the 10-4 dilution of the water prior to being filtered or treated. This film had 1.04 x 10-2 CFU/mL. The CC film in the middle was the 10-6 dilution of the sari filtered water and it had a CFU/mL = 6.7 x 10-5.

Fecal Float March 30, 2017

Purpose: To observe the various microorganisms that could be inhabiting our waste. Procedures: 1. Take a small scoop of the fecal matter with a wooden tongue depressor and mix with about 2 mL of flotation solution in a weighing boat to make a homogeneous suspension. 2. Strain the dispersed fecal matter through the cheese cloth into a 10 mL conical tube. 3. Fill the tube up to the top with the fecal solution using a transfer pipette. Make sure the solution has a convex surface. 4. Put the cover slide on top of the tube/solution. The cover slip has to contact the solution surface completely so it may spill a little. 5. Let it sit for 10 mins. 6. The eggs will rise to the top and stick to the glass. 7. Lift up the cover slip vertically and put onto a glass slide. 8. Look for parasite eggs, cysts and other life forms with the 10x magnification.

Results:

Figure 43. The photo to the left depicts what was observed under the microscope after performing the fecal float procedure. No eggs, cysts or parasites are depicted.

No life forms were found in the sample we performed a fecal float on.