2021_398_Disc_iffusion_Antimicrobial_Week_1_Student_Version (2)

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BME298/398 Antimicrobial Module Anti-Microbial Analyses: Kirby-Bauer Agar Diffusion Assay (note: Material from Antimicrobial Week 1 Prelab Exercise Part B is included here for easy reference. Be sure to complete the exercise in the PreLab 1 Exercise folder, and submit the assignment) Watch this YouTube video from Addgene (4:29), which demonstrates basic benchtop aseptic technique for working with bacteria and for sensitive molecular biology protocols including work with nucleic acids such as cloning. Note that PCR, and quantitative PCR especially, require another level of “clean” not addressed here. https://www.youtube.com/watch?v=wttvhJU9PZM A few clips in this video show mammalian cell culture flasks (pink medium!), which are typically handled inside a biosafety cabinet, where the downflow of HEPA filtered air provides a fully sterile atmosphere. Culture of mammalian cells requires rigorous attention to sterility and handling of all materials, due to the potential for even a single bacterium or fungal spore to contaminate and rapidly overgrow mammalian cell cultures. Benchtop aseptic technique is more common with bacterial, fungal and plant cultures, which are generally less susceptible to overgrowth by undesired organisms. In benchtop aseptic technique, flames are often, although not always, used. A flame serves multiple purposes: > It creates an updraft in the work area, so that spores and contaminated particles in the non-sterile room air are less likely to fall into open plates or bottles of media and other reagents. >It can be used to warm the neck of a flask or bottle, similarly creating an updraft in the vicinity of the opening to reduce risk of contaminants falling in. Note that this brief flaming does not provide enough heat to sterilize the glass. Never touch the necks or threaded areas of flasks or bottles. >It can be used to sterilize metal tools and loops. Heating a loop red hot or dipping tools such as forceps in alcohol and igniting it will sterilize the metal. Be aware that only the tip of the implement will be sterile. Any part of the tool that has been touched, even with gloves and alcohol wiped fingers, should be presumed to be contaminated. Open flames… Alcohol… Flammable paper and plastic materials… BME298-398 Wet Design Lab Module: Anti-Microbial Analyses: Kirby-Bauer Assay (SHB 4-10-2022) Page 1 of 16

What could possibly go wrong? BE VIGILANT!! Watch this ID Laboratory YouTube video (9:05), which shows an example of the Kirby- Bauer disc diffusion assay. Be alert for nine (or more!) mistakes made by the lab worker wearing the white gloves. https://www.youtube.com/watch?v=Np87w5kCL-4&t=112s In the video, the test plates are inoculated with colonies picked directly from a stock plate. In today’s exercise, we will use a diluted overnight liquid culture. Both methods are commonly used. The important similarity is that the bacteria are freshly grown. Do not use a plate or other culture that has been stored, for example, in the refrigerator. Antimicrobial Module Week 1 Anti-Microbial Analyses: Kirby-Bauer Agar Diffusion Assay Biomaterials and Anti-microbial Activity Biomaterials are often used in environments where control of bacterial/microbial growth is important. For example, biomedical implants may acquire biofilms, or surgical/wound sites may become infected. Incorporation of antimicrobial agents into biomaterials can effectively inhibit these processes. The Kirby-Bauer Agar Diffusion Assay is a rapid and convenient method to assess the release of antimicrobial agents from various materials, and also to determine the susceptibility of specific bacterial strains to particular antibiotic/antimicrobial formulations. This laboratory provides an introduction to the Kirby-Bauer Assay and basic microbiological techniques. Various formulations of silver and selected consumer products advertised as having anti-infective properties will be tested for activity against the reference strain E. coli ATCC ® 25922™. Standardized discs preloaded with antibiotics and also blank discs will be used as controls. An example of how this technique could be used to test the antibiotic eluting properties of a biomaterial is illustrated in the 2014 study by Mohiti-Asli et al . posted in LabArchives. A comprehensive review by Zilberman and Elsner (2008) is also posted. Prior to lab please review the Kirby-Bauer reference article and the articles by Zilberman and Elsner and Mohiti-Asli et al. Note that the reference article provides general information on the assay. Specifics of the class laboratory activity may differ. BME298-398 Wet Design Lab Module: Anti-Microbial Analyses: Kirby-Bauer Assay (SHB 4-10-2022) Page 2 of 16

Safety: Safety glasses, labcoats and gloves are required Personal Protective Equipment (PPE) for these exercises, and will be provided to all students. Long pants and closed shoes are required in the laboratory. Loose fitting clothing should not be worn; long hair should be tied back. No eating or drinking in the laboratory. Wash hands before leaving. Follow all safety instructions given by TAs or instructors. Many of the antimicrobial agents being tested are toxic if ingested, inhaled or absorbed through the skin, and may be harmful to the environment. 70% ethanol is flammable and can harm eyes and mucous membranes. Be aware of hazards when handling these materials. The strain of E. coli used in this protocol is not known to cause disease in humans or other animals, or to be hazardous in the environment. However, due to the high concentrations of bacteria used, all contaminated materials and all culture waste are disposed of in biowaste containers for decontamination. Materials: Controls: Standardized commercially prepared discs preloaded with antibiotics, in dispensing cartridges: >Positive Control -Tetracycline 30 microgram Oxoid; Thermo Scientific CT0054B Pack of 250 (5x 50 per tube) >Strain Control - Penicillin G 10 units Oxoid Thermo Scientific CT0043B Pack of 250 (5x 50 per tube) >Negative Control -Blank Antimicrobial Susceptibility Discs – ¼” in dispensing cartridges; Oxoid Thermo Scientific CT0998B Pack of 250 (5x 50 per tube); blank discs are also used as carriers for test agents Alternate items: Blank Paper Disk Remel Thermo Scientific R55054 100/vial Sterile Cloning Discs ¼”/6.4 mm Bel-Art SP Scienceware F37847-0003 Experimental samples: Four different alginate based wound dressings, cut in ¼” circles using a paper punch. >Algicate + Gelling Calcium Alginate Wound Dressing Sterile 10x10 cm; Circle A Medical 261144 (Abbreviation – Alg 1) >Algicate Ag Silver Calcium Alginate Dressing Sterile 10x12.7 cm; Circle A Medical 261545 (Abbreviation – Alg 2) >Areza Calcium Alginate Dressing Sterile 10.8x10.8 cm; Areza Medical 57354 (Abbreviation – Alg 3) >Areza Antibacterial Alginate with Silver Dressing Sterile 10.8x10.8 cm; Areza Medical 57258 (Abbreviation – Alg 4) BME298-398 Wet Design Lab Module: Anti-Microbial Analyses: Kirby-Bauer Assay (SHB 4-10-2022) Page 3 of 16

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Four different creams and gels, which will be applied using blank paper discs >Organa Silver Gel 100 ppm nano; Organa International >Neosporin Cream , 3.5 mg Neomycin Sulfate, 10,000 units Polymyxin B Sulfate, 10 mg Pramoxine HCl per gram; Johnson & Johnson Consumer Inc. > Neosporin Ointment , 3.5 mg Neomycin Sulfate, 5,000 units Polymyxin B Sulfate, 400 units Bacitracin Zinc, per gram; Johnson & Johnson Consumer Inc. >First Aid/Burn Cream, 0.13% Benzalkonium Chloride, 0.5% Lidocaine HCl; First Aid Only, Acme United Corporation Four essential oils: For each oil, 0.5 uL will be applied to a blank disc, followed by 2.0 uL sterile water >Oregano Essential Oil 100% pure; NOW Foods >Organic Lemon Essential Oil; Zongle Therapeutics LLC >Natural Clove Bud Essential Oil 100%; Vinevida >Palmarosa Essential Oil 100% Pure; SVA Three liquid solutions. Two are suspensions of insoluble materials. These will be vortexed well and 2 uL applied to a blank disc >Silver Sulfadiazine Micronized USP; Spectrum Chemical S1093 (2021 Lot# 4JG0022; 1% in 50% glycerol) >Silver-exchanged Zeolite; Aldrich 382280-25G (2021 Lot# MKCK0553; 1% in 50% glycerol); zeolite spheres are crushed in a mortar and pestle >Silver Nitrate AgNO3 Certified ACS, Fisher Chemical S181-25; (2021 Lot# 195447; 1% in 50% glycerol); silver nitrate is soluble in aqueous solution General Reagents: Glycerol Certified ACS; Fisher Chemical G33-1 (2021 Lot#182836) Bacto Tryptic Soy Broth; Fisher BD 211825 Bacto Tryptic Soy Agar; Fisher BD 236950 Consumable Supplies 6” Cotton Swabs Sterile; Fisher 22029681 Transfer pipets, sterile Culture Test Tubes 17x100 mm Fisherbrand 14-956-6D sterile PS, vent caps Microcentrifuge tubes for liquid test agents; screw cap sterile Micropipet tips, various sizes; use Fisher 02707442 reach barrier tips for 0.2-2 uL micropipet; 15x100 mm sterile Petri dishes; Fisherbrand FB0875712 cs 500 25/sleeve Kimwipes; paper towels 70% EtOH for cleaning/sanitizing Specific prepared materials for lab exercise: Sterile water in 15 mL conical centrifuge tube (approx. 10 mL) Tryptic Soy Broth in 15 mL conical centrifuge tube (approx. 10 mL) Tryptic Soy Agar Plates, approx. 8-10 per group. These were poured previously. BME298-398 Wet Design Lab Module: Anti-Microbial Analyses: Kirby-Bauer Assay (SHB 4-10-2022) Page 4 of 16

Live Bacterial Culture: 200 uL overnight culture E. coli ATCC strain 25922, in a 17x100 mm snap top culture tube. Equipment/Lab Items: Fine point forceps to handle discs Medium tip forceps to handle alginate materials Cell Density Meter: Ultrospec 10 – Biochrom or WPA Biowave CO8000 Micropipet set: 2 uL, 20 uL, 200 uL, 1000uL Inoculation turntable Sharpie markers Rulers General lab equipment for course support: Standard refrigerator/ -20C freezer Biosafety cabinet (recommended but not required) -86C ULT freezer or liquid nitrogen for storage of bacterial stocks Shaking incubator Bacterial incubator DI water should be available Autoclave Analytical Balance Top loading balance Weigh boats, spatulas 500 mL media bottles (4-6; more if pouring plates) 500 mL graduated cylinder 125/250 mL bacterial culture flasks; Nephlo flasks optional, but convenient General bacterial culture items, including Pipet-Aid or similar device, inoculation loops; parafilm Small and large biowaste bags For general information, lab prep instructions are included after the student instructions. Student Lab Activities: All procedures will be performed at the lab bench. Flames will not be used in this exercise. Before starting work, clear a work area and clean the area, pipets and other items using 70% ethanol and paper towels Use aseptic technique to avoid introducing unwanted organisms into the bacterial cultures or contaminating sterile materials and reagents. If bacterial cultures are splashed, dripped or otherwise caused to contaminate the work area or any lab items, promptly decontaminate using 70% ethanol and paper towels. Discard the towels in the large red biowaste can. BME298-398 Wet Design Lab Module: Anti-Microbial Analyses: Kirby-Bauer Assay (SHB 4-10-2022) Page 5 of 16

Change gloves promptly if you suspect they are contaminated. Instructors will supply a tube with approx. 200 uL overnight culture of E. coli ATCC 25922 in Tryptic Soy Broth. For this assay, a bacterial suspension with OD 600 of approx 0.5 will be used. Step 1: Plan your experiment and label the plates. Label the bottom of each plate. Each sample set will be made in duplicate (A and B). Divide each plate in six sectors as shown. Each plate should include a negative control (blank disc), a positive control (tetracycline), and up to four experimental samples. Include your section (298/398 M/W AM/PM) and initials. Also label one plate that will be kept uninoculated (no bacterial culture applied), as a quality control. As a reminder, there are four alginate samples, four “cream/gel” samples, four oils, three liquid/suspension samples and the strain control antibiotic, penicillin. Note: Write close to the edges of the sectors so the writing will not interfere with viewing the sample/disc and zones of inhibition. The image below is an example of how your plates should be prepared. These plates show results of a trial assay, which tested 50% glycerol (negative), 4-7 nm and 0.5-1 nm silver nanoparticles (both also negative), tetracycline (positive control) and penicillin, the “strain control”. The strain control will be discussed in class. BME298-398 Wet Design Lab Module: Anti-Microbial Analyses: Kirby-Bauer Assay (SHB 4-10-2022) Page 6 of 16

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Using the Cell Density Meter: The Cell Density Meter is a simplified spectrophotometer that measures optical density at 600 nm. OD 600 is a common measure of cell density for yeast and bacteria and can be used to estimate the number of bacteria and standardize the number between samples and experiments. An OD 600 = 1.0 very roughly correlates with 8 x 10 8 cells/mL for E. coli . If the number of Colony Forming Units (CFU; one CFU is one living cell) is important, it will be necessary to plate a dilution series of the culture to make a standard curve that represents the specific situation. Many factors, including the instrument used to make the OD 600 measurement, the number of living vs. dead cells and the degree of clumping of cells in the culture can substantially influence the relationship between OD 600 and CFU. For precision measurements, cuvettes should be used due to their optical clarity and controlled dimensions. However, the Cell Density Meter can also be used directly with clear culture tubes, typically polystyrene or glass, or with a nephlo flask , a specialized culture flask that has a test tube arm on the side. The flask can be tilted so that the liquid culture will fill the tube, which can then be inserted directly into the Cell Density Meter. After measurement, the tubes or flasks can be returned to the incubator for continued growth if desired. The option for direct measurement in either tubes or flasks allows convenient monitoring of the growth of the culture, without the need to transfer to cuvettes. Tips for obtaining the best readings from the Cell Density Meter: 1. The Density Meter will accept tubes with diameters from 10-17 mm. Be sure the tube used to blank the Density Meter is of the same diameter and type (brand and product number) as the tube being read. 2. Check the User Manual for the minimum volume of liquid necessary for the diameter of tube. For example, a 10 mm tube requires at least 0.9 mL liquid. A 16 mm tube requires 2.2 mL. The manual does not list 17 mm diameter tubes, but we use a minimum of 2.5 mL. 3. Unlike cuvettes, culture tubes are not optically perfect. When taking multiple readings in the same tube, make a mark on the tube and insert it in the same orientation each time you take a measurement. When blanking the instrument, always use a tube of the same type being used for the culture readings. 4. Plastic and glass tubes can be scratched by the clamps inside the Density Meter. Insert the tube straight into the opening and do not twist from side to side either when inserting or removing, especially if multiple measurements will be made in the same tube. This is particularly important if using nephlo flasks. They are expensive and not disposable, and if scratches accumulate on the tube, their utility is lost. BME298-398 Wet Design Lab Module: Anti-Microbial Analyses: Kirby-Bauer Assay (SHB 4-10-2022) Page 7 of 16

5. OD 600 readings taken of the same culture in different diameter tubes differ only slightly, despite the difference in pathlength. However, always use the same type of tube within an experiment and between experiments, if you will need to compare data at any point. The lab has two styles of Cell Density Meter. The instruments are identical, but the touch pads are labeled differently. The instructions are for the WPA Biowave CO8000. The “T” button on the Biowave corresponds to the button with the tube/arrow symbol on the Ultrospec 10; The Biowave “R” button is labeled OA/100%T on the Ultrospec 10. “T” (tube/arrow symbol) = Test “R” (OA/100%T) = Reference Step 2: Prepare the suspension of E. coli 1. Position the Density Meter conveniently on your bench, plug it in and turn it on. If the battery is charged, the plug in may not be needed. Please plug in when finished to recharge the battery. 2. Prepare the Reference tube: Aseptically transfer a minimum of 2.5 mL (more is fine) TSB to a 17x100 mm polystyrene culture tube. If the tube is not already labeled, mark the tube “R” or “Reference” near the top. 3. Insert the R tube into the meter, press the R button. It should read 0.00 Abs. Using a Sharpie marker, make a vertical line on the front of the tube so you can insert it in a consistent orientation for future readings. 4. Without twisting the tube, pull it out, then reinsert it. Press T. The reading should be 0.00 Abs. If it is not, repeat step 3, then step 4. If you are unable to obtain a reading of 0.00 with T, consult instructor. 5. Obtain a tube with 200 uL overnight bacterial culture from instructor. Using a sterile transfer pipet (small wrapped plastic pipet), add 3 mL TSB and mix. Insert the tube into the Density Meter, Press T, and record the result in your lab notebook. 6. Adjust the OD600 of the culture to 0.5 by adding additional TSB and mixing (a transfer pipet works well for this) until the desired OD is obtained. If the tube gets too full, discard some in the waste beaker and continue adding TSB. Record the total amount of TSB required (it may be approx. 5 mL) Part 3: Inoculate the plates Inoculate all your plates with the diluted E. coli culture before placing any samples. 1. Place an agar plate face up on the inoculating turntable. BME298-398 Wet Design Lab Module: Anti-Microbial Analyses: Kirby-Bauer Assay (SHB 4-10-2022) Page 8 of 16

2. Dip a 6” cotton swab into the bacterial suspension, then press it against the side of the tube to allow excess solution to drain away. 3. Holding the lid of the agar plate in a partially lifted position, move the swab back and forth over the agar to entirely cover the surface. 4. Turn plate 60°, continue swabbing, turn another 60°, swab some more to obtain an even distribution of bacteria across the entire plate. 5. The same swab can be used for multiple plates, however, if it starts to unravel, get a fresh one. Part 4: Apply the samples General note: Keep the materials as clean as possible. >The paper discs are all sterile, however, some of the materials, for example the oils, suspensions and Organa nano silver gel have not been sterilized. >The alginate wound dressings are inside sterile packaging, however, the punching process may allow the outside of the package material to contact the alginate dressing. >In general, keep tubes and containers closed as much as possible and handle the gels and alginate circles inside sterile Petri dishes. >Wipe the forceps with 70% ethanol between each sample. 1. Obtain the experimental samples from central location: >Blank paper discs and antibiotic discs are in cartridges. Groups may need to share. >The oils and silver solutions/suspensions should be in screw cap microcentrifuge tubes. >For the creams/gels, use a marker to divide an empty Petri dish into four sections and label one section for each cream/gel. You may write inside the dish but only allow the pen to touch the surface, don’t stick your fingers in there. Apply a SMALL amount, approx. the size of a dried split pea, of each material to the dish. Instructor will have a demo. >For the alginate wound dressings, label four empty Petri dishes on the bottom, (inside or outside, but as above, don’t stick your fingers in the dish), one for each dressing. The alginate dressings will be punched from a sterile envelope into the Petri dishes, using a regular ¼” paper punch. The dressings are very fragile and require the support of the paper/plastic envelope in order for a clean punch to be made. The envelope materials will also be in the Petri dish. When applying the samples to the plates, be sure to only apply the dressing material, not the paper or plastic envelope pieces. (note: punches of alginate dressing may already be prepared) 2. Apply the samples to the plates . Use the sharp pointed forceps and wipe with ethanol between each sample. Gently press each disc into the surface of the agar, but do not damage the agar or the disc. DO NOT move the disc around on the agar, even a BME298-398 Wet Design Lab Module: Anti-Microbial Analyses: Kirby-Bauer Assay (SHB 4-10-2022) Page 9 of 16

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tiny bit. This can disrupt the formation of the ROI. If it falls in the wrong place, leave it there. If necessary, you can always prepare another plate. >> FIRST apply all the negative control discs , then all the positive control discs to all the plates, followed by the penicillin strain control discs (only on one set of plates). >>For all samples , apply the duplicates at the same time. >>For the liquid samples , first apply the paper disc to the inoculated plate, then apply the sample (2 uL in all cases) to the disc. >>For the lemon, oregano and other essential oils , first apply 2 uL of sterile water to the center of the disc, then pipet 0.5 uL oil directly to the disk. The water seems to help keep the volatile oil from rapidly covering a large area of the plate. >>Z eolite and silver sulfadiazine are insoluble and must be mixed into suspension just before application. Vortex, then apply 2 uL to the disc, before the particles have a chance to settle. The silver nitrate should be soluble at this concentration, but vortex it anyway. >>The alginate dressings are applied directly, no blank disc. When applying the samples to the plates, be sure to only apply the dressing material, not the paper or plastic envelope pieces. The dressings are fluffy and fragile. If available, use broader tipped forceps to handle them. Gently apply all the dressings to the plates, carefully pressing them down. Then apply 20 uL sterile water to each dressing, and again gently press the fibers into contact with the agar surface. >> To apply a cream/gel to the inoculated plate, pick up a small amount of the material on both sides of a sterile paper disc. The amount should cover the disc, but not be so much that it squeezes out when the disc is gently pressed to the plate. When all samples are applied, place the plates face up in the bacterial incubator, along with the uninoculated quality control plate. Although bacterial plates are normally cultured upside down, in this case there is a chance the samples could fall off. Tomorrow , your instructors will remove your plates from the incubator and put them in the refrigerator. Next week , you will measure your ZOIs, and make some conclusions! Clean up Clean up!!! >Any liquid culture should be poured in the bleach bucket in the sink, and all contaminated disposable items discarded in the biowaste containers. >Thoroughly wipe down your work area, pipets, and any materials that could have possibly been spattered with bacteria with 70% ethanol. BME298-398 Wet Design Lab Module: Anti-Microbial Analyses: Kirby-Bauer Assay (SHB 4-10-2022) Page 10 of 16

>Replace all items where they belong. >Tidy your workbench. >Check with instructor to see if there is anything else you need to do before leaving. > Remove you gloves. Store your labcoat in a gallon Ziploc bag. Wash your hands and place bag in your locker. Week 1 Notebook questions (30 pts): Please answer these questions in your lab notebook following the Week 1 activity: 1. What is the Biosafety Level of the bacterial strain E. coli ATCC ® 25922™? (1 pt) The biosafety level of the bacterial strain E. coli ATCC® 25922™ is BSL-1 (Biosafety Level 1). 2. What are standard laboratory practices for this level (PPE, containment etc.)? (3 pts) Standard laboratory practices for Biosafety level 1 include routine disinfection of work table and bench, the use of proper aseptic technique when pipetting, minimizing splashing throughout the experiment, all pipetting must be done mechanically, and appropriate biohazard signage must be used. PPE for Biosafety level 1 include lab coats, gloves, long pants, close toed shoes, and eye protection. 3. Why is it important to avoid contaminating laboratory surfaces and equipment? (3 pts) Avoiding contamination of laboratory surfaces and equipment is essential to ensure the accuracy and validity of experiments, protect laboratory personnel from exposure to hazardous microorganisms, and comply with regulatory requirements. 4. What is the Zone of Inhibition? (1 pt) The zone of inhibition is the circular area around an antimicrobial disk or a spot where a microorganism is unable to grow due to the presence of an inhibitory substance. The zone of inhibition is a useful tool for evaluating the potency of antimicrobial agents and for determining the susceptibility of microorganisms to these agents. 5. One important determinant of the size of the ZOI is the test compound’s rate of diffusion in agar. With that in mind: a. How can you estimate how quickly a molecule or particle will move through a gel substrate with no agitation (only due to Brownian motion)? What are at least two physical parameters you need to know? (3 pts) Hint: look up the Stokes-Ei Polyla BME298-398 Wet Design Lab Module: Anti-Microbial Analyses: Kirby-Bauer Assay (SHB 4-10-2022) Page 11 of 16

nstein equation. One possible resource here: https://chem.libretexts.org/Bookshelves/Physical_and_Theoretical_Chemi stry_Textbook_Maps/ Supplemental_Modules_(Physical_and_Theoretical_Chemistry)/Kinetics/ 09%3A_Diffusion b. What happens over time in terms of the concentration of antibiotic in the agar after you place, say, one disc with a reservoir of antibiotic on the surface of a Petri dish with an even distribution of bacteria? Describe in 1-2 sentences and/or sketch. (Hint: think about food coloring diffusing through water – it doesn’t instantaneously equilibrate to an even concentration throughout the solution.) (2 pts) c. Will all of the bacteria experience equal concentrations of your antibiotic or will it vary with distance from the center of the disc? (1 pt) d. Say there is a cutoff point at which the antibiotic is no longer concentrated enough to kill the bacteria (e.g., 10% concentration relative to a 100% antibiotic solution). Given that and your answers to b and c, what does the size of the ZOI reflect? (2 pts) 6. The dose of penicillin G in the standard discs is 10 units. How many micrograms is this? (4 pts) 10 units of penicillin G is 6 micrograms. 7. What polymer(s) was/were used to make the fibers tested in the 2014 paper by Mohiti-Asli et al.? (5 pts) 8. The Elsner and Zilberman (2009) Research Article (not the review article) shows an illustration of the Kirby-Bauer test. Which antibiotic is illustrated? How many different strains of bacteria were tested? (5 pts) BME298-398 Wet Design Lab Module: Anti-Microbial Analyses: Kirby-Bauer Assay (SHB 4-10-2022) Page 12 of 16

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Prep instructions for Instructor and TA reference: At the beginning of the first lab, each student will be issued a lab coat and safety glasses, which they will store in a gallon Ziploc bag until the end of the module. Gloves are supplied in the lab. Make sure enough labcoats, safety glasses and gloves are available for the anticipated number of students. Students may reuse Ziploc bags from prior classes, so save them at the end of the module. No more than 2-3 weeks before class, streak a fresh TSA plate with E. coli ATCC strain 25922. If existing plates are older, make 2 fresh plates, streaking directly from stocks frozen in 50% glycerol. Stocks are stored at -86C (consult instructor for location). The day before lab, inoculate 20 mL TSB for overnight culture. If there are morning and afternoon sections on the same day, the overnight culture flask can be used in the morning, then kept at RT and used in the afternoon as well. For Monday AM labs, to avoid having to start a culture on Sunday, a flask can be grown over the weekend at ambient temp, 225 rpm. 2 hrs before the beginning of class, add 2 mL culture to 20 mL TSB , incubate at 37C, 225 rpm. Consult with instructor regarding number of students/groups. Students typically work in pairs. Autoclave 1x 500 mL DI water Prepare and autoclave 2x500 mL TSB per label instructions Check availability of TSA plates, which should be left over from prior classes. Each group will need 8-10 plates. There should be extras. Prepare 1 per group plus a few extras; use the label maker for all labels: Sterile water in 15 mL conical centrifuge tube (approx. 10 mL) Tryptic Soy Broth in 15 mL conical centrifuge tube (approx. 10 mL) >Note: The 15 mL polypropylene tubes can be autoclaved and reused. Check for leftover tubes from prior semester. Autoclave on dry cycle with caps removed or very loose. If labels bubble up, wait until tubes are cool and dry, then carefully lift and reseal the labels. Tryptic Soy Agar Plates, approx. 8-10 per group. These were poured previously. Empty labeled round bottom culture tubes (use the label maker), one per group: >200 uL E. coli 25922 (use fresh sterile tubes) >Density Meter TSB Reference – these can be rinsed and reused, if not too scratched. Due to the short time they are in use, sterility is not necessary. Mix up 40 mL 50% glycerol in sterile water, in a 50 mL conical tube BME298-398 Wet Design Lab Module: Anti-Microbial Analyses: Kirby-Bauer Assay (SHB 4-10-2022) Page 13 of 16

Prepare 10 mL of the following, 1% w/v in 50% glycerol: 100 mg/10 mL; solutions should be used within 2 weeks; >Silver Sulfadiazine Micronized USP; Spectrum Chemical S1093 (2021 Lot# 4JG0022; 1% in 50% glycerol) >Silver-exchanged Zeolite; Aldrich 382280-25G (2021 Lot# MKCK0553; 1% in 50% glycerol); zeolite spheres are crushed in a mortar and pestle >Silver Nitrate AgNO3 Certified ACS, Fisher Chemical S181-25; (2021 Lot# 195447; 1% in 50% glycerol); silver nitrate is soluble in aqueous solution Dispense 1 mL into 3-6 (1 tube per bench) microcentrifuge tubes. All sections will share the same tubes. Be sure to keep the particulates suspended. A transfer pipet is good for this purpose; mix well with pipet just before each transfer. Some solutions with darken with time, this is OK. The tubes can be rinsed and reused in future semesters. Note: new item number for round bottom tubes, Falcon 14-959-1B from Fisher. Ordered 1 case 3/25/2022 – see how they are. Prepare 1 tube per bench with approx. 500 uL oregano oil and lemon oil; use screw cap microcentrifuge tubes. Also prepare similar tubes with additional oils as per instructor’s direction. Setup at each student station: >Biowaste bag in 1L plastic beaker >250 mL plastic beaker, labeled for liquid waste >70% Ethanol in spray bottle >Box Kimwipes >Box Wypall L10 paper towels >Organizer with cotton swabs, transfer pipets (min 10 each) >2 inoculation turntables >2 super-fine tip forceps >1 med tip forceps >2 Sharpie markers, extra fine point >set of four micropipets >tips for micropipets: Include Reach barrier tips for 0.2-2 microliter pipet (needed for oils-Fisher 02707442; other tips are less fussy but make sure there are appropriate sizes for all the micropipets) >micro tube rack >Rack for 15 mL tubes/17 mm culture tubes (note-the culture tubes are larger diameter than the 15 mL tubes, and may not fit in all racks for 15 mL tubes) >Vortex mixer >Cell Density Meter Test materials in central location: See student protocol for specifics on the test materials. BME298-398 Wet Design Lab Module: Anti-Microbial Analyses: Kirby-Bauer Assay (SHB 4-10-2022) Page 14 of 16

>Micro tube rack with oils, one of each per group of lemon/oregano, one total for other oils >Micro tube rack with silver solutions, one of each per group >Labeled Petri dishes with punches of alginate wound dressings; each group needs two of each type, so 10 per type per dish is plenty for total of 4 sections/4 gps each; They are staticky, so have a few extra. These same punched circles are used in Week 3, four of each type per lab section. >To prepare, use ¼” paper punch, punch through paper/plastic dressing wrap, allow circles of paper/plastic and dressing to fall into dish. Prepare the non- silver dressings (Alg 1 & Alg 3) first, then the silver dressings (Alg 2 & Alg 4). When finished, rinse the paper punch with water and then 70% EtOH to remove any fibers. Allow to dry COMPLETELY before punching any more circles. See additional notes in student protocol. Four creams; one empty Petri dish per group. Disposable plastic spatulas for the Organa gel. Sharpie marker for students to label own plates. Make a demo plate with a small dab of each (see student instructions) Antibiotic discs are stored refrigerated and desiccated, if open. Check with instructor. Remove from refrigerator at least one hour before class. >Tetracycline discs, 50 per cartridge: Each group uses 8, check for four cartridges with a minimum of 10 discs in each. 4 sections x 4 groups x 8 per group = 128 discs; >Penicillin discs, 50 per cartridge: Each group uses 2, but be sure there are 2-3 cartridges to avoid delays >Blank discs, 50 per cartridge: Each group uses 30 assuming they don’t drop any. Put two cartridges per lab station, checking each is reasonably full. Have additional cartridges at the central location. 4x4x30 = 480 for a class. Other setup/cleanup items: Large biowaste bucket to collect class waste; Check with instructor regarding autoclaving of the waste for the class. Staff may be available to take care of this. 1L “slop” bucket in sink with bleach; Have a cover for the bucket/beaker, because the odor can be intense. Wait until near the end of class, then pour in enough bleach so that it will be at least 10% of the final volume after all class liquid is added. Students or TA should pour all excess liquid culture into the bleach bucket and discard disposable tubes in the large biowaste can. After lab, collect the TSB and distilled water tubes; pour out excess, rinse with DI water and place to dry; Check with instructor regarding Reference tube, it may be kept with the TSB already in it for the next section. BME298-398 Wet Design Lab Module: Anti-Microbial Analyses: Kirby-Bauer Assay (SHB 4-10-2022) Page 15 of 16

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After students have left, empty small biowaste bags into large biowaste can. If the waste does not readily fall out of the small bag, dispose of the whole bag and get a new one. Do not touch any of the waste in the small bag to empty it. Organize student stations, replace items where they belong, check for any missed waste. All silver waste (1% glycerol solutions) must be disposed of as hazardous waste. Check with instructor. These will not be disposed of until the end of the module. Save the labeled 50 mL and microcentrifuge tubes for next class, rinsing with DI water and setting to dry. BME298-398 Wet Design Lab Module: Anti-Microbial Analyses: Kirby-Bauer Assay (SHB 4-10-2022) Page 16 of 16

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