Spore Assay

Day 1

1. Determine the volume of culture you need to plate your samples. Per plate, you will spot 4 groups of 5 20uL spots, meaning that you need 400uL of concentrated cells per plate. Typically, if you are making 3 plates per strain, this means you will need 20mL of initial culture per strain to do this. If you made several smaller liquid cultures (~12mL), combine them as long as they look roughly the same density. Take 10.5mL from each culture and combine in a 50mL falcon tube so that your total volume is 21mL.

2. Remove 1mL, dilute as normal for the spec to determine the OD600. This will leave 20mL of culture left (to make the math easy). Use the OD to cell/ml conversion table on the desktop of the lab computer to determine what volume you will resuspend your sample in to reach 5x10^9 cells/ml. The volume column in the table is how much TPM you would use to resuspend 1mL of culture. Multiply this by 20 if you are using 20mL of culture.

Spinning down: 3. Place the 20mL (or other volume) into a 50 mL falcon tube and spin down in the superlab bucket rotator for 8 minutes at max speed (3900 rpm). Decant and resuspend in 4 mL of TPM. spin down for 5 minutes. Repeat wash step two more times. Resuspend to final volume.

4. Ensure resuspended cells are evenly mixed, and then spot 4 groups of 5 20uL spots onto large TPM plates that have been brought to room temperature and are not wet on the surface. Divide your plate into quadrants and in each quadrant, spot 5 20uL just far apart enough so that your cell spots do not run into each other. Leave plates open until completely dry. This is best done under the hood to prevent spores and dust from falling. WHen dry, place them in the 32C incubator for 5 days.

5. Cover plates, invert them, and store in the incubator for 5 days at 32C.

Day 2 (Read ahead to make sure you have the time set aside)

1. Remove plates from the incubator and image if desired.

2. Turn heat block (or preferably the water bath filled to a volume that will entirely cover 9mL in falcon tubes) to 50-55C in preparation, and check that you have enough sterile water to complete the below protocol. For each plate you prepared, you will need 9mL of sterile MiliQ water in a 15mL falcon tube (pre-filling and pre-labeling these tubes is recommended).

3. Using a bent metal spatula (one should already be prepared for you in the lab, sterilize in between by dipping in 70% ethanol and running through the flame), gently scrape 3 of your 4 groups of cell spots from each plate together into one small area, careful so as not to disrupt the surface of the agar. Use the spatula to transfer the cells to the pre-filled 9mL of sterile water.

4. Repeat step 3 for each of your plates.

5. Important; WEAR EAR PROTECTION WHEN USING SONICATOR. Turn on the sonicator and sterilize the probe with isopropyl alcohol. To do this, dip the probe into the tube of alcohol, switch to 'continuous' to turn on, and then switch back to 'remote' to turn off. Sonicate the isopropyl for a few seconds to sterilize, then rinse the probe in sterile water to remove the alcohol by turning the sonicator on for a few seconds.

6. With the sonicator set to an intensity of 6 (or 7, whatever it is set to when taped over), and using a 'blank' tube of 9mL of sterile water without cells, get a feel for where you need to hold the tube, etc to have a constant output wattage. Then use that same probe placement for all samples. Tip: use the gradations on the falcon tube for this, and try to keep your hand placement on the tubes consistent as well. Hold tube toward the top, to help vibrations travel through sample.

7. Sonicate each sample for 10 cycles each (1 cycle: 10 sec on, 30 sec off) for a total of 100 active seconds. Record the output wattage in your lab notebook, especially if different for each sample. Sonicate just one sample at a time, sterilizing the probe with more isopropyl alcohol in between samples. This is super important so you don't cross-contaminate strains! Have a bucket of ice prepared, and on the 30s 'off' times, place the tube in the ice so your sample doesn't overheat.

8. Incubate samples at 50-55C for 2 hrs.

9. While sonicated samples are incubating in the heat block, prepare your tubes of CTTSA. Each tube that you currently have will be split into three for technical replication. (Ex. if you have three tubes from the sonicator, you will dilute and plate those out on 9 total plates at the end.

10. Melt down the CTTSA in the microwave, make 4mL aliquots for each tube that you need, and keep warm in a water bath or heat block until you need them. You can also prep tubes for serial dilution (see below), and take out the CTTYE plates you will need so they can come to room temperature.

11. Once your samples are done heating, vortex to break up any clumps and you are ready to serial dilute.

12. From your well-mixed tube of collected spores, take 100uL of sample into 900uL of sterile water, a 1:10 dilution. Repeat for another 1:10 dilution, changing tips in between and ensuring that samples are well mixed. Then transfer 100uL into 4mL of CTTSA and pour over a room temp CTTYE plate (with Kan if cells are resistant).

13. Allow CTTSA to solidify for 15min and put plates in incubator at 32C for 5 days. Count colonies and then back calculate from your serial dilution to determine the number of spores recovered.