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.

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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.

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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.

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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.

Biofilms

Procedure: Degrading the July 17 Biofilms

The biofilms created on July 17 grew successfully. The solutions used to degrade the biofilms were Mr. Clean and Lysol. 3200 μL of solution was added to each Petri dish. 30 mins after the solutions were added, they were disposed of in the biohazard waste.

Two of the Petri dishes had been degraded. The other three from July 17 were washed with water once before being placed back in the 37 ˚C incubator. The goal of these three biofilms is to see if they can still grow and maintain the RFP biofilms.

Procedure: Reculturing July 17 Biofilms

To reculture the biofilms, 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 listed above in the procedure for degradation of the biofilms. Cultures left in the 37 ˚C shaking incubator

Bee: Oxalic Acid Gradient

Procedure: The procedure for the gradient remains the same as July 9. The concentrations of oxalic acid were quickly added to their respective tubes to minimize differences in time of oxidation.

Objective: Make new biofilms

Biofilms: In a 50 mL falcon tube, 21 mL of LB media, 210 µl of RFP, and 21 µL KAN were pipetted into it and the solution was mixed. 3 mL of the solution was then pipetted into 7 35x10 mm Petri dishes.

These were then put into the incubator for 24 hours at 37˚C.

Results: Reculture of one of the July 17, 2015 biofilms –Day 4

The 5 mL tube of LB media labeled “before” when bleach was not added had RFP cultures growing at the bottom of the tube. This showed that we were able to reculture the biofilm. The 5 mL LB media tube labeled “After” which had been washed with bleach contained no biofilm growth indicating that the bleach had degraded the 4 day old biofilms.

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Biofilms: Growing new biofilms

Objective: To growth new biofilms for future testing with various means

Procedure: In a 50 mL falcon tube, 21 mL of LB media, 210 µl of RFP, and 21 µL KAN were pipetted into it and the solution was mixed. 3 mL of the solution was then pipetted into 7 35x10 mm Petri dishes.

These were then put into the incubator for 24 hours at 37˚C.

Biofilms: Bleach Concentration Gradient

Objective: From our work we are able to determine how effective different cleaners are in degrading biofilms. To test the resilience of the biofilms in another mean, we will test how well the biofilms are degraded at different concentrations of bleach

Procedure:

1.Standard solution of 0.252% bleach was made from the stock bottle for a 50 mL volume

2.The following concentrations had been made from the standard solution of bleach and water. Each concentration was diluted by two folds of the previous solution.

• 0.252% • 0.126% • 0.063% • 0.0315% • 0.01575% • 0.007875% • 0.0039375%

3.The dilutions were made for a 15 mL falcon tube using water and standard bleach solution

4.3.2 mL of one of the bleach dilutions were placed in one the Petri dishes containing biofilms from the previous day. This process was repeated for the other six dilutions.

5.The biofilms were then allowed to degrade for 15 mins.

6.The bleach solutions in each of the Petri dishes was emptied and rinsed once with Milli-Q water.

7.LB media tubes were labeled with one of the bleach solutions. 5 µL of KAN was added to each labeled tube.

8.20 µL of LB from one tube was taken out and placed on the edge of one of the Petri dishes.

9.The media was then placed back into the tube it was taken out of.

10.This process (Steps 8-9) was repeated for the remaining six tubes

11.LB media tubes were placed in the 37˚C incubator for 24 hours.

Bee: Oxalic Acid Gradient

Objective: Continue to test the specificity of the oxalic acid gradient using KMnO4

Procedure: The procedure for the gradient remains the same as July 9. The concentrations of oxalic acid were quickly added to their respective tubes to minimize differences in time of oxidation.

Biofilms: Results

The biofilm gradient using different concentrations of bleach was taken out at 8:13 and observed for visible signs of RFP growth. All seven LB media tubes seem to have visible signs of RFP growth. The 1/16 bleach dilution however does not seem to have any RFP growth. Cultures will be left in the 4˚C fridge overnight to allow the cells to precipitate at the bottom.

Biofilms: Bleach Concentration Gradient

Objective: The previous biofilm gradient using the different concentrations of oxalic acid may not have been successful. A repeat of the gradient is needed to verify results of the previous trials.

Procedure: Protocol the same as on July 22, 2015 with the previous concentrations of bleach used.

1.The dilutions were made for a 15 mL falcon tube using water and standard bleach solution

2.3.2 mL of one of the bleach dilutions were placed in one the Petri dishes containing biofilms from the previous day. This process was repeated for the other six dilutions.

3.The biofilms were then allowed to degrade for 15 mins.

4.The bleach solutions in each of the Petri dishes was emptied and rinsed once with Milli-Q water.

5.LB media tubes were labeled with one of the bleach solutions. 5 µL of KAN was added to each labeled tube.

6.20 µL of LB from one tube was taken out and placed on the edge of one of the Petri dishes.

7.The media was then placed back into the tube it was taken out of.

8.This process (Steps 8-9) was repeated for the remaining six tubes

9.LB media tubes were placed in the 37˚C incubator for 24 hours.

Biofilms: Making New Biofilms

Objective: to make new biofilms to continue testing.

Procedure: In a 50 mL falcon tube, 21 mL of LB media, 210 µL of RFP culture, and 21 µL of KAN were pipetted into it. 3 mL of the solution was then pipetted into 7 35x10 mm Petri dishes.

The Petri dishes were then put into the incubator at 37˚C for 24 hours.

Biofilms: Reculturing 6 Day sample

Objective: testing to see if there are any bacteria left

Procedure: A before and an after tube were made with the 6 day sample. In the before tube, a sample was taken and put into an LB media tube to test if there are any bacteria alive on it. In the after tube, a standard solution of bleach was put on the Petri dish for 15 minutes.

The tubes were then put into the incubator at 37 ˚C for 24 hours.

Biofilms:

Objective: To make new biofilms to be able to continue lab work, reculture old biofilms to continue to see if we can successfully reculture and degrade them, and try a new method of testing using different antibiotics.

Procedure: New Biofilms

In a 50 mL falcon tube, 21 mL of LB media, 210 µL of RFP, and 21 µL of KAN was pipetted into it. 3 mL of the solution was then pipetted into 7 35x10 mm Petri dishes.

The Petri dishes were then put in the incubator for 24 hours at 37 ˚C.

Procedure: Reculturing Biofilms

The Day 9 biofilm was recultured by pipetting a proper amount of LB media onto the side of the Petri dish. This was then pipetted back into the 5 mL LB media tube along with 5 µL of KAN.

This was then incubated in the shaker for 24 hours at 37 ˚C.

The biofilms that were washed with the cleaners Tilex, Mr. Clean, Fantastik, and water was used as a control were recultured 24 hours after the cleaner had been removed. They were recultured in the same manner as mentioned above.

These were then also incubated in the shaker for 24 hours at 37˚C.

Bees:

Objective: Continue trials of the oxalic acid gradient.

Procedure:

The same procedure was implemented that had been used in previous trials.

Results:

Objective: New biofilms were created so that testing of various sorts could be continued. To make sure the RFP biofilms that have been recultured in the past are actually from the plates, different antibiotics were used to reculture the biofilms. If RFP persisted in the different types of antibiotic we can ensure that the recultures are not our cultures –and therefore no contamination.

Procedure: New Biofilms

In a 50 mL falcon tube, 21 mL of LB media, 210 µL of RFP, and 21 µL of KAN was pipetted into it. 3 mL of the solution was then pipetted into 7 35x10 mm Petri dishes.

The Petri dishes were then put in the incubator for 24 hours at 37 ˚C.

Procedure: Reculturing Biofilms

The Day 9 biofilm was recultured by pipetting a proper amount of LB media onto the side of the Petri dish. This was then pipetted back into the 5 mL LB media tube along with 5 µL of KAN.

This was then incubated in the shaker for 24 hours at 37 ˚C.

The biofilms that were treated with the antibiotics KAN (x2), AMP, TET, CAM, IPTG, and water which was used as a control, were recultured 15 minutes after the antibiotics had been removed. They were recultured in the same manner as mentioned above.

These were then also incubated in the shaker for 24 hours at 37˚C.

Biofilms Results:

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Bees

Objective: Continued with oxalic acid gradient testing to determine accuracy

Procedure: the same procedure was used that has been done previously.

Results:

Objective: to use the Typhoon to have quantitative data to see if the biofilms have grown.

Procedure:

The biofilms made on July 30, 2015 were washed out 3 to 4 times. 5 biofilms were put into the Typhoon that had not been treated with cleaners.

TYPHOON SETTINGS: 532 Green Laser. 580 BP. 100 Microns.

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Bee: Oxalic Acid Gradient

Objective: To continue with oxalic acid gradient testing to test accuracy of test before using the assay for construct testing.

Procedure:

Procedure was taken from previously on July 9, 2015.

Results:

Figure 1 Different concentrations of oxalic acid at 5 minutes of incubation at RT

Biofilms: Using the Typhoon

Objective: to use the Typhoon to have quantitative data to see if the biofilms have grown.

Procedure:

The biofilms made on July 30, 2015 were washed out 3 to 4 times. 5 biofilms were put into the Typhoon that had not been treated with cleaners.

TYPHOON SETTINGS: 532 Green Laser. 580 BP. 200 Microns.

PHOTO

Objective: Create new biofilms, Perform transformations on all 3 constructs into pSB1C3, use typhoon to quantify results of biofilms undergone cleaner treatment.

Methods: 1) Biofilms – 7000uL of total volume (6230uL LB Media, 700uL RFP-Culture, 70uL Kanamycin). Was distributed into 200uL wells for a total of 35 samples.

2) Transformations were performed see (insert transformation protocol).

3) Biofilms were tested with Cleaners.

Objective: To produce biofilms and remove old biofilms.

Procedure: Making the biofilm.

Added total volume 8000μL consisting of 7912μL of LB media, 80μL of cell culture and 8μL of antibiotic (KAN).

Made 200μL aliquots 35 wells on a 96 well plate.

Placed in 37 C incubator.

Objective: Quantify biofilm formation on 96 well plates, using OD600 readings followed by crystal violet/cleaner treatment. Then final quantification using OD550.

Results: 35 wells filled with 200uL culture + LB + Kan, left for 24 hours. OD600 readings follow for the 35 wells.

Objective: to test biofilms with antibiotics and to make new biofilms to continue testing on them.

Procedure:

Antibiotics Testing

Biofilms that were incubated for 24 hours were cleaned. In 12 2 mL tubes, 1650 µL of double distilled water was pipetted into each of them. Into 2 2 mL tubes, 3.3 µL of AMP was pipetted into them. This process was repeated for 4 other tubes with CAM and TET. A control with water was also used.

2 of the 2 mL tubes were then placed into a 35x10 mm Petri dish. This was done with 5 other 35x10 mm Petri dishes. 3 of them were rinsed after 15 minutes while the other 3 and the control were rinsed after 30 minutes. These were then recultured by pipetting 100 µL of LB media into the rim of the 35x10 mm Petri dish. It was the put back into the tube along with 5 µL of KAN.

These were then put into the incubator for 24 hours at 37˚C.

New Biofilms

In a 50 mL falcon tube, 21 mL, 200 µL of E.coli with RFP, and 20 µL of KAN was combined. 3 mL was then pipetted into 7 35x10 mm Petri dishes.

These were then put into the incubator for 24 hours at 37 ˚C.

We did a regular Biofilm test using antibiotics instead of cleaning solutions.

This matched our first test showing that biofilms can be recultured after using a treatment of antibiotics.

Objective: Taking 48 hours biofilms and cleaning them we are using water. We are using the Typhoon to find RFP concentrations.

Objective:

Determine a concentration of oxalic acid that will prevent cell growth.

Procedure:

The same procedure was used as has been in previous trials.

Results:

Biofilms: Making biofilms

Incubated new biofilms at 37 degrees for 24 hours. Protocol used was the same as has been in other trials.

Biofilms: Detergent Treatments

On seven biofilms, one was controlled with water, two were treated with Mr. Clean, two were treated with Fantastic, and two were treated with Tilex. Detergents remained in dishes for fifteen minutes, then were removed and rinsed twice with water.

Biofilms

Objective: To clean biofilms and to make new ones to continue testing.

Procedure:

Cleaning Old Biofilms

August 25th biofilms were cleaned using 3.2 mL of Mr. Clean, GreenWorks Cleaner, and Lysol in 2 35x10 mm Petri dishes each. Water was put into a seventh Petri dish of the same size to be used as a control. The cleaning solutions were left on for 15 minutes and then were rinsed.

The Petri dishes were then put into the refrigerator for 24 hours at 4˚C.

Making New Biofilms

New biofilms were made by putting 21 mL of total volume of LB media, RFP culture, and KAN in a 50 mL falcon tube. The RFP culture was added in a 1/100th dilution and the KAN was added in a 1/1000th dilution. 3 mL of this solution was then pipetted into seven 35x10 mm Petri dishes.

These were put in the incubator for 24 hours at 37˚C.

Objective: to test the effectiveness of our constructs and to use the typhoon to get quantitative data.

Procedure:

12 overnight biofilms in 35x10 mm Petri dishes were rinsed twice with distilled water and then dried. These were then put into the Typhoon.

Colonies from dextranase, nuclease, and PDP were made into cultures by extracting a colony from each and then putting it into a tube of LB media along with CAM.

Objective: to test the effectiveness of our constructs.

Biofilms:

In a 50 mL falcon tube, 4 mL of LB media, 40 µL of RFP, and 4 µL of KAN were added together. 300 µL of this solution was then pipetted into 15 wells of a 96 well plate. This was repeated 3 more times with dextranase, nuclease, and a combination of dextranase and nuclease replacing RFP. However, 4 µL of CAM was used instead of KAN.

This was then put into an incubator for 24 hours at 37˚C.