Isolation of bacteriophages

Materials
· Overnight culture of E. coli DH5a
· 125 mL Erlenmeyer flask
· 40 mL waste water
· LB medium
· Warm LB agar plates
· Warm top agarose (0.7 %)
· 37 °C incubator with shaker platform
· 37 °C incubator
· 45 °C water bath
 


1. Amplification of bacterophage

• Pipette 5 mL LB medium into flask containing 40 mL of waste water
• Inoculate the waste water in the flask with 5 mL of overnight culture of E. coli
• Incubate culture at 37 °C, shaking for 24 h

Prior to session 2, 2.5 mL overnight E. coli culture was inoculated with 25 mL LB-medium, and incubated at 37 C shaking until OD600 = 0.8.


2. Bacteriophage isolation and plating

• Transfer 10 mL of waste water-bacteria-phage culture into a centrifuge tube, and centrifuge the sample at 2000 rpm for 5 min.
• Pipette the supernatant into a 10 mL syringe barrel fitted with 0.2 micron filter. Storage tube with phages can be stored at 4 °C for several months. Vortex this tube before serial dilutions.
• Prepare a series of microcentrifuge tubes for serial 10-fold dilutions. Label 10 tubes 1-10. In each tube, pipette 0.9 mL of sterile LB.
• Serial dilution: Transfer 0.1 mL of phage suspension into tube 1, and vortex. Using the same pipette, transfer 0.1 mL of the sample from tube 1 into tube 2, and vortex. Repeat this process, transferring 0.1 mL from tube 2 to tube 3, and so on, mixing each time.
• Get ready 0.7 % top agarose in a falcon tube and let it be "cooled" to 45 °C in water bath. (Turn on water bath and wait until 45 °C)
• Label 11 new small falcon tubes with the number 1-10 ("1-10" phage+DH5a and "Control" containing only DH5a). Distribute 0.5 mL of log-phase E. coli in each of the small falcon tubes.
• To each tube of bacteria, add 0.1 mL of corresponding phage dilution. Note: if you work from the most diluted to the least, you can use the same pipette. Mix gently by inverting (NO VORTEX).
• Incubate at 37 °C for 10 min.
  
• In the meantime, label 11 warm, agar plates. Keep the plates in the 37 °C incubator until you are ready to use them.
• Collect pre-warmed plates from incubator. Pipette 3 mL of top agarose to the cell-phage tube. Cap the tube, and mix gently inverting it 3 times. Quickly pour the mixture onto warmed plate. You can tip the plate slightly to spread the top agarose. Push plate aside, but do not pick it up until agarose solidifies.
• Air dry with open lid for 15 min in the sterile hood.
• When the top agarose has solidified, incubate the plates, inverted at 37 °C for 3 h.

3. Examiniation of bacteriophage plates, phage storage

• Count plaques on plates (See lab journal day 22)
• Pick an isolated plaque for long-term storage: pipette 1 mL of LB into a small Eppendorf tube, and add 1 drop of chloroform. Use a small pipette tip and pierce the agar surrounding plaque, and pick out the agar containing plaque. Place the agar with plaque into 1 mL of LB. Store the plaque in refrigerator (4 °C).

Supercompetent cell production

Day 1

1. Make an E. coli DH5α culture in a small flask with 10 mL Psi-medium. Inoculate culture at the end of the day (4 pm), leave in 37 °C shaking incubator overnight.


2. Prepare transformation buffers (TFB) for day 2.

TFB 1 (200 mL)

• Add the following to a 250 mL flask:
○ Potassium acetate (0.588 g)
○ Rubidium chloride (2.42 g)
○ Manganese chloride (2.00 g)
○ Glycerol (30 mL)
• Fill up with MilliQ water to a total volume of 180 mL
• Adjust pH to 5.8 with a pH meter by adding diluted acetic acid (0.2 M)
• Fill up with MilliQ water to a total volume of 200 mL

TFB 2 (100 mL)

• Add the following to a 250 mL flask:
○ MOPS (0.21 g)
○ Rubidium chloride (0.121 g)
○ Calcium chloride (1.1 g)
○ Glycerol (15 mL)
• Fill up with MilliQ water to a total volume of 90 mL
• Adjust pH to 6.5 with a pH meter by adding diluted NaOH (0.2 M)
• Fill up with MilliQ water to a total volume of 100 mL

Day 2

1. Start culture as early as possible. Transfer 1 mL culture to a new shaking flask with 100 mL Psi-medium. Grow in shaking incubator at 37 °C until OD600 reaches 0.4.
2. Place the culture on ice for 15 minutes, along with TFB1 and TFB2.
3. Centrifuge the culture for 5 minutes at 4000 rpm at 4 °C. Pour off supernatant. (Safe break point, if you leave cells on ice.)
4. Resuspend pellet carefully in 40 mL TFB1. Let sit for 5 minutes on ice.
5. Centrifuge for 5 minutes at 4000 rpm at 4 °C. Pour off supernatant. Resuspend pellet carefully in 3 mL TFB2.
6. Mark as many 1.5 mL Eppendorf tubes as you will need with cell strain name and the date.
7. Aliquot 100 uL of resuspended cells into the 1.5 mL Cryo tubes. Leave all tubes on ice until finished aliquoting. Flash freeze for 5-10 seconds in liquid nitrogen, store at -80 °C.

Transformation

1. Thaw supercompetent cells on ice for approximately 10 minutes.
2. Using a pipette, transfer 1 μL of mini-prepped plasmid into the cells. Pipette up and down while stirring the pipette tip gently around the tube for 5-10 seconds.
3. Incubate the cells on ice for 20 minutes (can be shortened to 5 if you are in a hurry).
4. Heat shock the cells for 45 seconds at 42 °C. Immediately put cells on ice for 2 minutes after.
5. Add 900 uL LB medium to the cells. Incubate at 37 °C, 200 RPM for 1 hour (can be shortened to 45 minutes if you are in a hurry).
6. Plate out 100 uL of the transformation mixture onto agar plates with the appropriate selection agent

DNA purification (MiniPrep)

1. Centrifuge 1 mL of bacterial culture in a clear 1.5 mL tube at full speed (16,000 g) for 20 seconds in a microcentrifuge. Discard supernatant.
2. Add 200 μL of P1 buffer (red) to the tube and resuspend the pellet completely by pipetting.
3. Add 200 μL of P2 buffer (green), mix by inverting the tube 2-4 times, and incubate for 1-2 minutes. Do not vortex.
4. Add 400 μL of P3 buffer (yellow) and mix gently but thoroughly. Allow the lysate to incubate at room temperature for 1.2 minutes. Do not vortex.
5. Centrifuge samples for 4 minutes at 16,000 g.
6. Plave a Zymo-Spin IIN column in a collection tube and transfer the supernatant from step 5 into the column. Avoid distrurbing the pellet.
7. Centrifyge the Zymo-Spin IIN assembly for 30 seconds at 16,000 g.
8. Discard the flow-through and retyrn the column to the same collection tube.
9. Add 200 μL of Endo-Wash Buffer to the column and centrifuge for 30 seconds at 16,000 g.
10. Add 400 μL of Plasmid Wash Buffer to the column and centrifuge for 1 minute at 16,000 g.
11. Transfer the column into a clean 1.5 microcentrifuge tube and then add 30 uL of DNA Elution Buffer directly to the column matrix and incubate for one minute at room temperature.
12. Centrifuge for 30 seconds at 16,000 g to elute the plasmid DNA.

Store the plasmid at -20 degrees.

Restriction digestion of DNA

Materials

• Up to 1 μg DNA (0.5 ug)
• Appropriate buffer (2 μL)
• Restriction enzyme (0.5-1 μL)
• Distilled water

Method

1. Mix buffer and DNA, and fill up to 20 μL with distilled water. Mix well.
2. Add enzyme, pipette up and down with a pipette several times to mix.
3. Incubate for 2 hours or overnight at 37 ˚C in a heat block.

NB! Enzymes should always be kept in an ice rack when outside of the freezer. Always wear gloves when handling enzymes.

Gel electrophoresis

1. Pour agarose gel (recipe below) containing the appropriate* DNA binding dye into a gel tray of appropriate size. Make sure the tray is tightly put into the gel tub.
2. After the gel has set, add TAE buffer until the gel is completely submerged in buffer.
3. Add DNA ladder with dye to at least one well in the gel.
4. Add your DNA samples to be separated into separate wells.
5. Connect the gel tub to a power supply, and run the gel at 80-110 V until the color bands are approximately 75 % to 80 % down the length of the gel.
6. Disable the power supply and put the gel into the imaging apparatus.
7. On the computer open the ImageLab program.
8. Select the appropriate* protocol, position gel correctly and take image.
9. Print image and save the image file to the iGEM2016 folder (C://). Use a descriptive name for the file.
10. If running gels to excise DNA fragments, attach the UV screen to the drawer.
11. Put on orange safety goggles.
12. Turn on light (top button)
13. Excise the desired bands using a scalpel. Clean the scalpel before and between excisions. Transfer the excised gel bands to clean 1.5 mL Eppendorf tubes, and label them clearly. Store in freezer.
14. Dispose of the remaining gel in the blue barrel on the floor next to the imaging apparatus.
15. Rinse gel tray and gel tub in water and leave to dry.

*If excising bands use GelGreen. Otherwise use GelRed.

Always use gloves and safety goggles when working with gels.


Agarose gel recipe (0.8 %):

1. Add TAE buffer (400 mL) to 400 mL glass bottle
2. Add agarose (3.2 g) to the buffer
3. Boil in the microwave oven for approximately 5 min at medium power
4. Add GelRed or GelGreen (20 μL) (gelGreen if you want to cut out the band)

Loading of samples

• 2 μL ladder
• 20 μL sample + 4 μL loading dye (mixed)
• Run for 45 minutes
• Take picture in the ChemiDoc and print out

TECAN infinite M200 PRO

Two scripts are utilized for OD-measurements.

Script No.1

• Hold Temp: 37˚C
• Performed: N-cycles
• Each cycle is measured at absorbance 600 nm in 300 uL culture�25 flashes per reading
• Shaking:
• Duration: 300 second
• Mode: Orbital
• Amplitude: 2.5 mm
• Frequency: 244 rpm

Script No.2

• Hold Temp: T
• Performed: 24-cycles
• Fluorescence intensity scan with settings
• Scan selection: Emission scan
• Mode: Top (from top)
• Gain: manual = 255

Integration

• Lag time: 100 μSec
• Integration time: 20 uSec
• 25 flashes per reading
• Excitation wavelength: 390 nm
• Emission wavelength: 450-540 nm
• Step size: 10 nm
• Z-position: 18000 μm

Shaking:

• Duration: 300 second
• Mode: Orbital
• Amplitude: 2.5 mm
• Frequency: 244 rpm

PCR (Polymerase chain reaction)

Kit: EHIFI-RO Roche "Expand High Fidelity PCR System"
Procedure: The general instruction-sheet has not been followed.
Changed made are described below:

1. Briefly vortex and centrifuge all reagents before starting.
2. Prepare two mixes in a sterile microfuge tubes (on ice):

Mix 1 (for one reaction)

Reagent	Amount
Sterile double-dist. Water	8 μL
Deoxynucleotide mix, 10 mM of each dNTP)	1 μL
Upstream primer	2 ng/L
Downstream primer	2ng/L
Template DNA	1 μL, 20 ng

Mix 2 (for one reaction)

Reagent	Amount
Sterile double-dist. Water	18 μL
Expand High Fidelity buffer, 10×conc. with 15 mM MgCl2	5 μL
Expand High Fidelity enzyme mix (polymerase)	0.75 μL
dMSO	1.25 μL

Combine Mix 1 and Mix 2 in a thin-walled PCR tube (on ice).
Gently vortex the mixture to produce a homogeneous reaction, then centrifuge briefly to collect sample at the bottom of the tube.

DNA assembly (NEBuilder)

The mass of each fragment can be measured using NanoDrop. Recommended amount of fragments used for assembly:

	2-3 fragment assembly*	4-6 fragment assembly**	Positive control
DNA ratio (vector : insert)	1 : 2	1 : 1	-
Total amount of fragments	X μL	X μL	10 μL
DNA Assembly Master Mix	10 μL	10 μL	10 μL
Deionized water (dH2O)	10-X μL	10-X μL	-
Total volume	20 μL	20 μL	20 μL

*Optimized cloning efficiency is 50-100 ng of vectors with 2 fold excess of inserts. Use 5 times more inserts if size is less than 200 bps. Total volume of unpurified PCR fragments in the assembly reaction should not exceed 20%.
**To achieve optimal assembly efficiency, it is recommended to design >20 bp overlap regions between each fragment with equimolarity.

1. Thaw and vortex master mix thoroughly and keep on ice.
2. Set up the following reaction (see table) in a PCR tube on ice:
   vector (backbone) + insert (plasmid) + dH2O + master mix
3. Incubate samples in a thermocycler at 50 C for 15 minutes (2-3 fragments) or 60 minutes (4-6 fragments).
4. Store samples on ice or at -20 C for subsequent transformation.