We stained inoculated biofilms and performed the Crystal Violet assay. Left in 37 C for 36 hours. We did 4 trials 100uL, 4 trials 200uL, and 4 trials 300uL. Biofilm formation was successful, with varying band thickness. We also replicated this with two 5mL LB petri dish inoculated with 20uL E.coli. We added 5mL of crystal violet each to the petri dishes to cover the whole plate.

We also performed an oxalic acid gradient with 0%, 0.05%, 0.1%, 0.2%, 0.3%, 0.4%, 0.5%, and 2.0% oxalic acid to give a final volume of 6mL. We added 20uL of E.coli to each 6mL tube. Left in 37 C.

We did a 1/100 dilution of E.coli into LB media, and left it in 37 C for (hopefully) 24 hours. We did 4 trials 100uL, 4 trials 200uL, and 4 trials 300uL.

Biofilm produced on 24h 96 well plate.

Testing to see if biofilm will produce on 12h 96 well plate (1 in 100 culture dilution).

And also doing 24h 96 well biofilm formation testing to see what detergents and chemical impacts have on the decrease in biofilm production.

*Biofilms placed at 8:00pm

Oxalic acid test run for 12 hours (0, 0.01, 0.05, 0.1, 0.15, 0.2, 0.4, 0.4, 0.5, 2.0%)

*Oxalic acid placed at 7:30pm

Tested the production of parafilm sacs for in vitro Varroa feeding. (using kimwipes and 2mL tubes as containment methods) .

Performed oxalic acid testing of E.coli growth in LB media at (0, 0.01, 0.05, 0.1, 0.15, 0.2, 0.4, 0.4, 0.5, 2.0%) concentrations. Placed at 6:50pm.

Produced more biofilms.

Had mites and created pouches including Water, LB, Supernatent, E.coli, E.coli RFP. Placed at room temperature and in chris’s drawer.

Took 48 and 24 hour biofilms and performed a bleach treatment on them (0.252%) for 30 minutes. .

Took oxalic acid readings after 23 hours of incubation

Readings are unrealiable as the cultures were left in the shaking incubator for over 23 hours. Still show the same general trend as past gradients.

Make oxalic acid samples again.

Make 24h biofilms.

Objective:Determine if we have successfully cultured mites in vitro using either paraffin pouches or bee pupae.

Table 2. Experimental design showing each of the treatments applied to the mites.

After 24 hours, all mites were found to be dead. Pupae showed some signs of black bacteria growth or browning on body. Will need to repeat or find another method of culturing mites in vitro.

Biofilm test performed on

Fantastic = Biofilm remained

Windex = Biofilm remained

Bleach = didn’t remain

Made overnight cultures for oxalic acid testing

13 hours left in shaker

Objective:Determine if the concentration of oxalic acid that kills bacteria will kill bees.

Procedure:Bees were obtained from the “Mighty Hive” near the penitentiary at 4:00 and placed into 5ml centrifuge tubes. The bees were transported to the University of Lethbridge and stored at room temperature until 8:00. Prior to experimentation, the tubes were supplemented with a sugar water solution to ensure that the bees would not starve. Following feeding, 2ml of each treatment were added to the 5ml tubes containing the bees and set horizontally along the bench. The abdomen of each bee was immersed in the treatment no matter what orientation the bee took. The bees were observed for 1 hour and subsequently euthanized in 70% EtOH.

Table 1. Observations of the bees following 60 minutes of exposure.

Discussion:The bees showed that they could survive well in concentrations of oxalic acid that were above the point where E.coli would die. Further tests must be done to determine the exact contact toxicity. Additionally, the contact toxicity to mites will need to be addressed.

Conclusion:The bees will be able to survive all of the oxalic acid that the E.coli can produce.

Bee: A More Specific Oxalic Acid Gradient

Objective: Determine a more specific concentration of oxalic acid that will prevent cell growth. Already we know that oxalic acid prevents cell growth between 0.05% and 0.2%. Now we want to determine a more specific concentration of oxalic acid that will prevent cell growth.

Cells were left in the shaker for 23 hours accidently. ODs were not taken because they had been left in the shaker too long.

Biofilms: Continued Growth of Biofilms.

Objective: Continue to determine what cleaners will destroy/degrade biofilms. Instead of using crystal violet dye, we are using E.coli DH5α cells with RFP (PSB4K5). 2 μL of KAN was added to 2 mL centrifuge tubes along with 1980 μL LB and 20 μL of culture.

300 μL of mix was pipetted into wells of 96 well plate. Biofilms left to grow 24 hours in 37˚C incubator.

Results: The biofilms did not grow on any of the plates as there was no red ring. Using fresh RFP cultures might fix this issue because the cultures we had used were prepared earlier in the month.

Objective: Due to the oxalic acid gradient from yesterday being left in the shaker too long, the gradient was repeated again.

Cells were left in the shaker for 15 hours and OD’ed the next morning.

Biofilms: Continued Growth of Biofilms

Objective: The biofilms made yesterday did not work, so fresh RFP cultures were prepared tonight for future biofilm testing. Colonies were picked from a streaked plate of RFP and placed in 5 mL of LB. 5 μL of KAN was added to each tube. Two picked colonies were prepared and left overnight in the 37˚C shaker.

Prepped bee pouches for in vitro mite/bee feeding.

Performed Bleach and Fantastic testing on RFP biofilms and did one with crystal violet to check for biofilm formation.

Made biofilms for July 3.

Objective: Determine if bees can ingest E.coli in liquid culture form with supplementation of sugar. Also, can mites survive on papers that have water and RFP liquid culture?

Procedure: Bees were obtained from the “Mighty Hive” near the penitentiary at 6:50 after a heinous struggle and lack of proper equipment. They were then fed RFP producing E.coli that had been supplemented with both glucose and sucrose sugars. The bees were then observed to determine if they were drinking the solution.

Mites were added to petri dishes following a sugar shake. The dishes had different paper derivatives soaked in treatment; H2O and RFP E. coli. They were then placed in the 27C iGEM incubator. They will be observed in the morning.

The bees were euthanized on EtOH, then surface sterilized in bleach, opened using a razor to expose the gut, and placed in LB to shake at 37C overnight. Problem: 50ul of kanamycin was added instead of 5ul. I suppose that’s what happens when you work a 13 hour day in the sun.

Results:

Table 1. Observations of the bees following 60 minutes of exposure.

Discussion: UPDATE TOMORROW

Conclusion: UPDATE TOMORROW

Objective: To continue growing biofilms and determine if certain cleaners will prevent further growth of the biofilms

Procedure: Same protocol used as June 30. Instead of using crystal violet dye, we are using E.coli DH5α cells with RFP (PSB4K5). 2 μL of KAN was added to 2 mL centrifuge tubes along with 1980 μL LB and 20 μL of culture.

300 μL of mix was pipetted into wells of 96 well plate. Biofilms left to grow 24 hours in 37˚C incubator.

Bees: Preparing Solutions for Oxalic Acid Gradient using Potassium Permanganate.

Objective: Solutions were prepared to be used for the oxalic acid assay that will see KMnO4 being oxidized by oxalic acid and therefore detect the presence of oxalic acid.

Procedure: To prepare the 1% and 20% Oxalic Acid concentrations, solid oxalic acid was weighed for a solution of 50 mL. 0.5 g of oxalic acid was weighed out and placed in a 50 mL falcon tube. 50 mL of d2H¬2O was added to the falcon tube to dissolve the oxalic acid and create the 1% Oxalic Acid concentration. Two falcon tubes of 1% Oxalic Acid was prepared.

10 g of oxalic acid was then weighed and dissolved in 50 mL of d2H¬2O in a 50 mL falcon tube for the 20% oxalic acid concentration. The 50 mL of 20% oxalic acid was then placed in the 60˚C oven to help dissolve the oxalic acid.

In addition 1 M H2SO4 was created using stock 5 M for the Wieden Lab. Both the 1 M H2SO4 and 1% oxalic acid were stored in the 4˚C fridge in E770.

Objective: To continue growing biofilms for future testing and to determine if biofilms can be recultured from previous ones that had been degraded based on the July 6 biofilms.

Procedure: New Biofilms

New biofilms were created for July 7 testing. Same protocol used as June 30. Instead of using crystal violet dye, we are using E.coli DH5α cells with RFP (PSB4K5). 2 μL of KAN was added to 2 mL centrifuge tubes along with 1980 μL LB and 20 μL of culture.

300 μL of mix was pipetted into wells of 96 well plate. Biofilms left to grow 24 hours in 37˚C incubator.

Procedure: Degrading the July 6 Biofilms

The biofilms created yesterday successfully grew. To degrade the biofilms, bleach, Tilex and Truly cleaner solutions were used that had previously been created. The solutions of biofilms were disposed of in the biohazard waste and the microtiter plates placed in water. They were vigoursly shaken to dry and 300 μL of respective cleaning solution added to each well. Since three plates of biofilms were created, each cleaning solution received its on plate. 15 mins after the solutions was added, they were disposed of in the biohazard waste.

Procedure: Reculturing the July 6 Biofilms

To culture the biofilms from yesterday, 300 μL of LB media was added to one well of a microtiter plate. The media was then pipette up and down to mix with any remnants of the biofilms. The media was then pipetted back into the 5 mL tube of LB media. 5 μL of KAN was added to the 5 mL tube of LB media.

Process was repeated for the remaining two plates that used different cleaners.

Note: Before cultures were degraded using the different cleaners, Kieran had taken a sample of the biofilms and prepared a 5 mL tube of LB to culture the biofilm that would be ensured to grow.

Cultures were left to grow 24 hours in 37˚C incubator.

Objective: Determine if yesterday’s recultured biofilms were successful. In addition continue to degrade biofilms using the household cleaners brought into lab for testing and see if we can successfully reculture the biofilms created yesterday.

Results: Reculturing of the July 6 Biofilms

We were successful in reculturing the biofilms to varying extents based on the overnight cultures of the degraded RFP biofilms. Based on reference of the biofilm that had been cultured before the cleaners were added, the Truly is the most red in colour. Both bleach and Tilex had a more clear appearncec similar to LB.

PHOTO

Procedure: Degrading the July 7 Biofilms

The biofilms created yesterday successfully grew. To degrade the biofilms, Tilex, Truly and Fantastic cleaner solutions were used that had previously been created. The solutions of biofilms were disposed of in the biohazard waste and the microtiter plates placed in water. They were vigoursly shaken to dry and 300 μL of respective cleaning solution added to each well. Since three plates of biofilms were created, each cleaning solution received its on plate. 15 mins after the solutions was added, they were disposed of in the biohazard waste.

Procedure: Reculturing the July 7 Biofilms

To culture the biofilms from yesterday, 300 μL of LB media was added to one well for one of microtiter plate that had been degraded with cleaner. The media was then pipette up and down to mix with any remnants of the biofilms. The media was then pipetted back into the 5 mL tube of LB media. 5 μL of KAN was added to the 5 mL tube of LB media.

Process was repeated for the remaining two plates that used different cleaners.

Note: Before cultures were degraded using the different cleaners, Kieran had taken a sample of the biofilms and prepared a 5 mL tube of LB to culture the biofilm that would be ensured to grow.

Cultures were left to grow 24 hours in 37˚C incubator.

Procedure: New Biofilms

New biofilms were created for July 9 testing. Same protocol used as June 30. Instead of using crystal violet dye, we are using E.coli DH5α cells with RFP (PSB4K5). 2 μL of KAN was added to 2 mL centrifuge tubes along with 1980 μL LB and 20 μL of culture.

300 μL of mix was pipetted into wells of 96 well plate. Biofilms left to grow 24 hours in 37˚C incubator.

Bees: Continued Preparation of Solutions for the Oxalic Acid Gradient Using KMnO4

Objective: Continue preparation of solutions being used for the oxalic acid gradients with KMnO4

Procedure

Solutions of 0.01 M of KMnO4 were prepared from stock. 0.158 g of KMnO4 was dissolved in 100 mL of d2H2O. 0.003 M of KMnO4 was then prepared using the 0.01 M solution.

The oxalic acid gradient was also set up with 10 1.5 mL centrifuge tubes with the following oxalic acid concentrations:0.1%, 0.2%, 0.3%, 0.4%, 0.5%, 0.6%, 0.7%, 0.8%, 0.9%, 1.0%.

A 1 mL volume was created for each concentration using 1% oxalic acid and d2H2O

All solutions and the oxalic acid gradient were set aside for tomorrow on the bench.

Objective: Determine if yesterday’s recultured biofilms were successful. In addition continue to degrade biofilms using the household cleaners brought into lab for testing and see if we can successfully reculture the biofilms created yesterday.

Results: Reculturing of the July 7 Biofilms

We were successful in reculturing the biofilms to varying extents based on the overnight cultures of the degraded RFP biofilms. Based on reference of the biofilm that had been cultured before the cleaners were added, the Truly is still the most red in colour. Both Fantastic and Tilex was cloudy with culture but not with the RFP used to originally culture the biofilms.

PHOTO

Procedure: Degrading the July 8 Biofilms

The biofilms created yesterday successfully grew. To degrade the biofilms, Fantastic and bleach cleaner solutions were used that had previously been created. Water was used as a control. 300 μL of respective cleaning solution added to each well. 15 mins after the solutions was added, they were disposed of in the biohazard waste.

Procedure: Reculturing the July 8 Biofilms

To culture the biofilms from yesterday, 300 μL of LB media was added to one well for one of microtiter plate that had been degraded with cleaner. The media was then pipette up and down to mix with any remnants of the biofilms. The media was then pipetted back into the 5 mL tube of LB media. 5 μL of KAN was added to the 5 mL tube of LB media.

Process was repeated for the remaining two plates that used different cleaners.

Cultures were left to grow 24 hours in 37˚C incubator.

Procedure: New Biofilms

New biofilms were created for July 10 testing. 2 μL of KAN was added to 2 mL centrifuge tubes along with 1980 μL LB and 20 μL of culture (E.coli DH5α cells with RFP (PSB4K5)).

300 μL of mix was pipetted into wells of 96 well plate. Biofilms left to grow 24 hours in 37˚C incubator.

Bees: Oxalic Acid Gradient Using KMnO4 –July 9

Object: To determine for the presence and concentration of oxalic acid present in solution using the oxidation of KMnO4

Procedure

1.Standard oxalic acid solution (1 mg/mL) was prepared with distilled water

2.100 mL of 0.003 M KMnO4 was prepared from appropriate dilution of 0.01 M KMnO4 in distilled water

3.500 mL of 2 N H2SO4 was prepared in distilled water

4.Assay mixtures contained different concentrations of oxalic acid ranging from 0.1 to 1.0 mg of oxalic acid (we had 1 mL samples of different percentage concentrations of oxalic acid), 5 mL 2 N H2SO4, and 2 mL of 0.003 M KMnO4

5.This mixture was incubated for 10 mins at room temperature (27˚C)

6.After 10 mins absorbance was recorded at 528 nm on the spec in E720

7.Reagent blank was prepared with distilled water

8.The calibration curve obtained is linear in concentration range of 0.1 mg to 1 mg/mL of oxalic acid

9.The regression equation is ∆A = 0.966C -0.027 with correlation coefficient 0.983 where C is the concentration of oxalic acid in mg/mL and ∆A is Ab-As

Data

Objective: Continue to degrade biofilms using the household cleaners brought into lab for testing and to see if we can successfully reculture the biofilms.

Procedure: Degrading the July 8 Biofilms

The biofilms created yesterday successfully grew. To degrade the biofilms, Fantastic and Lysol solutions were used that had previously been created. Water was used as a control. 300 μL of respective cleaning solution added to each well. 30 mins after the solutions was added, they were disposed of in the biohazard waste.

Procedure: Reculturing the July 9 Biofilms

To culture the biofilms from yesterday, 300 μL of LB media was added to one well for one of microtiter plate that had been degraded with cleaner. The media was then pipette up and down to mix with any remnants of the biofilms. The media was then pipetted back into the 5 mL tube of LB media. 5 μL of KAN was added to the 5 mL tube of LB media.

Process was repeated for the remaining two plates that used different cleaners.

Cultures were left to grow 24 hours in 37˚C incubator.

Objective: Determine if yesterday’s recultured biofilms were successful in being cultured again.

Results: Reculturing of the July 10 Biofilms

We were not successful in reculturing the based on the overnight cultures of the degraded RFP biofilms. All overnight cultures remained clear as the control with water using the Lysol and Fantastic cleaning solutions.

PHOTO

Objective: To determine the concentration of oxalic acid present in solution using the oxidation of oxidation of KMnO4

Procedure: same procedure as on July 9

Data:

Biofilms: Creating Biofilms

Objective: to create biofilms to continue testing on them with household cleaners brought into the lab.

Procedure: Creating Biofilms

Biofilms were created in 5 35x10 mm Petri dishes and in 4 wells of a 28-well plate by putting 3000 µL of LB media, 3 µL of KAN, and 20 µL of RFP into each dish and well.

These were left in the 37 ˚C incubator for 24 hours in order for the biofilms to grow.

Objective: Continue to degrade biofilms using household cleaners brought into to the lab to see if we can successfully reculture the biofilms.

Procedure: Degrading the July 13 Biofilms

The biofilms created on July 13 grew successfully. The solutions used to degrade the biofilms were Mr. Clean, Lysol, Tilex, and Fantistik. Water was used as a control. 3200 μL of solution was added to each well and Petri dish. 30 mins after the solutions were added, they were disposed of in the biohazard waste.

Procedure: Reculturing July 13 Biofilms

To reculture the biofilms from yesterday, 200 μL of LB media was added to two of the wells in the 28-well plate and to the Petri dishes that had been degraded with cleaner. The media was moved around the rim to where the biofilm had formed. Then the media was pipette back into the 5 mL tube of LB media. 5 μL of KAN was added to the 5 mL tube of LB media.

This process was repeated for each of the cleaners and the control listed above in the procedure for degradation of the biofilms.

The Petri dishes and the 28-well plate were left to air out on a sheet of paper towel.

Procedure: Creating Biofilms

Biofilms were made in 5 35x10 mm Petri dishes and 4 wells in a 28-well plate by putting 3000 μL of LB media, 3 μL of KAN, and 20 μL of RFP in to each Petri dish and well.

These were left in the 37 ˚C incubator for 24 hours so that they could grow.

Objective: Continue to degrade biofilms using the household cleaners brought into the lab for testing and to see if they can be successfully recultured.

Results: Reculturing the July 13 Biofilms

The recultured biofilms did not produce any RFP cultures as the cultures were more yellow than red. Water and Lysol were both cloudy but could have been contamination. The remaining cleaning soltuons of Mr. Clean, Tiklex and Fantastik were clear.

PHOTO

Procedure: Degrading the July 14 Biofilms

The biofilms created on July 14 grew successfully. The solutions used to degrade the biofilms were Mr. Clean, Lysol, Fantistik, Tilex, and GreenWorks cleaner. Water was used as a control. 3200 μL of solution was added to each well and Petri dish. 30 mins after the solutions were added, they were disposed of in the biohazard waste.

Procedure: Reculturing July 14 Biofilms

To reculture the biofilms from yesterday, 200 μL of LB media was added to one well for each Petri dish that had been degraded with cleaner. The media was moved around the rim to where the biofilm had formed. Then the media was pipette back into the 5 mL tube of LB media. 5 μL of KAN was added to the 5 mL tube of LB media.

This process was repeated for each of the cleaners and the control listed above in the procedure for degradation of the biofilms.

The Petri dishes were left to air out on a sheet of paper towel.

Procedure: Creating Biofilms.

Biofilms were made in 5 35x10 mm Petri dishes by putting 14.85 mL of LB media, 15 μL of KAN, and 150 μL of RFP into a 15 mL falcon tube. 3 mL of this solution was then pipetted into each Petri dish.

These were left in the 37 ˚C incubator for 24 hours so that they could grow.

From the overnight cultures of the July 14 biofilms, the ones recultured after the use of water and Tilex were red confirming we had recultured the two cultures. The one recultured from GreenWorks was also successful. Although not very red, it had an orange tinge indicating the presence of RFP. The Lysol reculture was more yellow in culture colour but might have been successful. The same results as previous for Fantastik showed no culture growth.

PHOTO

Biofilms

In one 15 mL falcon tube, 15 mL of LB media, 150 µL of RFP, and 15 µL of KAN were added. Then 3 mL of the solution was pipetted into 5 separate 35x10 mm Petri dishes.

The Petri dishes were then incubated in the 37 ˚C incubator.

Bee: Oxalic Acid Gradient