King's Broth Preparation
- 1L Distilled Water
- 20g Difco Proteose Peptone #3
- 1.5g K2HPO4
- 1.5g MgSO4•7H2O
- 10mL Glycerol
- 15g Agar (if making solid media)
Instructions
1. Bring the water to a boil.
2. Add in the peptone, and stir until dissolved.
3. Add the other ingredients, and stir until dissolved before cutting the heat.
4. Bottle, and autoclave to sterilize.
So many experiments in microbiology require the creation of media that it is often forgotten in terms of procedures. Kings Broth is an agar and broth recipe important of the growth of Pseudomonas strains, our model pathogen, and the name sake of our team, the Phage Court.
Materials per Liter
Filtrate Creation
- 2 50mL Falcon Tube
- 30mL distilled Water
- 1m of 500mM CaCl2
- 10mL Kings Broth
- 1mL of Overnight Liquid Host Culture
Instructions
1. Add the distilled water to the first falcon tube, and fill the rest with soil from the host's preferred environment.
2. Shake the mixture before letting it settle.
3. In the second tube, prepare the rest of the ingredients, and add 10mL of water from tube A.
4. Incubate the mixture at 28°C for 24 hours.
Phage exist almost everywhere in nature, and anywhere your host may be found it is likely there is also phage to target it. Our first attempt at this process used environmental water and leaf samples did not yield phage. Only on our second attempt when we included soil samples did we produce viable results.
Materials per Sample
Phage Isolation
- 18mL of KB agar
- 200mL 500mM CaCl2
- 100 mL overnight host
- Roughly 4 mL of Phage Buffer
- 10μL of filtrate
- One large plate
Instructions
1. Section the large plate into 6 sections.
2. While the agar is liquid, add the overnight culture alongside the calcium chloride and pour the plate.
3. Set up a row of 5 microtubes, and add 90μL of phage buffer to each.
4. Perform serial dilutions by first adding 10μL of filtrate to the first and then taking 10 from there for the next etc.
5. Once the plate has solidified, add 10μL of each dilution, and the original filtrate thrice in each secion, one for each row.
6. Incubate the plates at 28°C checking for plaques each day.
7. Once individual plaques have formed, pipette the agar of the plague up with a serological pipette, and put it in 100mL of overnight before incubating it for 24 hours at 28°C, this is your now lysate.
8. Repeat this process with the lysate until the plaques all look the same.
Because many different phage may be contained in one sample, this procedure allows us to purify each individual phage based on host focus.
Materials per Sample
Efficiency of Plaquing Test
- 18mL of KB agar
- 200mL 500mM CaCl2
- 100mL overnight host
- Roughly 4 mL of Phage Buffer
- 10μL of lysate
- One large plate
Instructions
1. Section the large plate into 3 rows of 12.
2. While the agar is liquid, add the overnight culture alongside the calcium chloride and pour the plate.
3. Set up a row of 12 microtubes, and add 90μL of phage buffer to each.
4. Perform serial dilutions by first adding 10μL of filtrate to the first and then taking 10 from there for the next ect.
5. Once the plate has solidified, add 5μL of each dilution thrice, one for each row.
6. Incubate the plates at 28°C for the ammount of time previously determined for plaques to form.
7. Count the plaques at the dilution where you are first able to see individual plaques and use this to calculate your titer.
8. Repeat this process three times with the same lysate for accurate data.
This process is very similar to isolating phage, but takes place with purified high titer lysate to determine how many infecteous phage are in it per Liter.
Materials per Sample
Colony Forming Unit Test
- 108mL of KB agar
- Roughly 4 mL of King's Broth
- 2mL of overnight host
- 1 mL distilled water
- 6 large plates
- Two Cuevettes
- An OD reader
Instructions
1. Section the large plate into 3 rows of 12.
2. Pour the plates with 18 mL KB agar each
3. Set up 5 2.5mL microtubes each with 1mL of KB broth
4. Perform serial half dilutions of the overnight host into the 5 microtubes.
5. Calibrate the OD reader with a cuvette with 1 mL of DI water, before measuring first the overnight, then each half dilution all in the same cuvette.
6. Set up a row of 12 microtubes, and add 90μL of kings broth to each.
7. Perform serial dilutions with 10 mL of the overnight.
8. Repeat 6 and 7 with each of the half dilutions.
9. Add 5μL of each 10th dilution thrice, one for each row.
10. Repeat 9 using each half dilution's set of 12 each on a seperate labeled plate
6. Incubate the plates at 28°C for the amount of time previously determined for host colonies to form.
7. Count the colonies at the dilution where you are first able to see individual colonies on each plate and use this to calculate your titer at each half dilution.
8. Plot the colony forming units against the opical density to create a curve on which to predict CFU from OD.
Colony forming unit tests or CFUs can allow us to determine the amount of bacteria relative to the optical density of a sample.
Materials per Sample
High Titer Lysate Creation
- 30mL Kings Broth
- 100μL Lysate
- Plate of Host Culture
Instructions
1. Inoculate the broth with host, and place in a shaking incubator.
2. Monitor the broth until cloudy, after which the phage should be immediately added.
3. Continue incubating until the broth clears again.
4. Refrigerate the lysate and perform an EOP to determine titer.
To get lots of DNA for DNA Prep, you first need a lot of phage from which to get the DNA. This procedure is meant to create a solution containing many phage per mL, usually the target being around 10^9.
Materials per Liter
Crispy BRIP Method
- 1 µL Spe1_HF
- 1 µL CIP
- 5 µL 10x Cutsmart
- 1000ng pGRB
- 22.8µL Nuclease free H2O
5. Incubate at 37˚C for 2 hours then held at 4˚C
6. 2-way 10µL Gibson reaction is done using the following reaction mixture:
- 1.7µL pGRB Spe1 digest
- 3.3µL gRNA C-term oligo
- 5µL 2x Gibson MM
7. Incubate at 50˚C for 1 hour.
8. Dialyze and place into -20˚C for storage.
Creating Strains:
9. Electroporate the dialyzed pGRB_Cterm plasmid was into the electrocompetent E. coli DH10B, recovere in rich SOC then plate onto LB+Amp grown at 37˚C.
10. Picked and screen colonies using GoTaq colony PCR using primers 50nt up and downstream of the Spe1 cut site, pick reverify and create a frozen stock from a colony with the 161nt insertion.
11. Electroporate the mating strain E. coli BW25113 with the pRedCas9, recovere in rich SOC media for 1 hour before plating on LB+DAP+Spec at 30˚C.
12. Make a frozen stock using a single isolated colony.
13. Both mating strain BW25113+pRedCas9 and DH10B+pGRB_Cterm need to be spotted on top of one another on LB+DAP and grown overnight at 30˚C.
14. Next day, the mixture growth was streak plated on LB+Spec+Carb selecting for DH10B+pRedCas9+pGRB_Cterm unselecting for the mating strain BW25113 that required DAP to grow.
15. Pick a single colony of the E. coli DH10B pRedCas9+pGRB_Cterm streak plate and make into a frozen stock and an overnight liquid stock.
Making Electrocompetent Cells:
16. Grow E. coli DH10B pRedCas9+pGRB_Cterm cells from an overnight culture until OD600 = 0.35-0.40.
17. Spin down 2ml of culture in microcentrifuge for 1 minute at full speed at room temperature.
18. Remove supernatant. Resuspend pellet in 2ml of ice cold sterile H2O.
19. Centrifuge for 1 minute at full speed at 4 degrees C. At this point, keep all cells on ice.
20. Remove supernatant. Resuspend pellet in 1ml of ice cold sterile H2O.
21. Centrifuge for 1 minute at full speed.
22. Remove supernatant. Resuspend pellet in 100ul of 10% glycerol.
23. Centrifuge for 1 minute at full speed.
24. Remove supernatant. Resuspend pellet in 40ul of 10% glycerol.
25. Make 20uL aliquots of cells in PCR strip tubes.
CRISPY-BRIP
26. Plate 8mL of MMB agar with 250uL of E. coli DH10B pRedCas9+pGRB_Cterm cells from an overnight and pour onto small petri dishes. Let cool for 10mins.
27. Electroporate 1uL of dsDNA substrate (Image 3 sequence) ] into electrocompetent E. coli DH10B pRedCas9+pGRB_Cterm cells.
28. Let cells recover and grow in 500uL SOC with the addition of phage for an MOI of 1-2.
29. Plate cells with 5mL-8mL of MMB agar and recovered cells as top agar onto previously made plates.
30. Grow plates overnight at 30˚C.
Next day, pick and use GoTaq colony PCR to screen phage with primers flanking homology arms to detect insert of sequence.
Gene modification technique utilizing gene editing tool CRISPR and recombination machinery lamda-red for bacteriophage. The following technique was created for model bacterium E. coli and coli phage but can be adapted for other organisms using respective recombination, CRISPR, and transformation of the new organisms. This is our most complex procedure and will be broken into pieces.
Instructions
1. To construct a working CRISPR system, a crRNA is needed to guide the CRISPR system. The crRNA will be 20nt long and must be on the coding strand of the DNA with a PAM sequence (5'-NGG-3' repeat) downstream of the gene of interest
2. The crRNA template is then placed in the pGRB plasmid using Gibson cloning or similar techniques creating the guide RNA (gRNA) system
3. Creating Plasmids:
4. Digest the pGRB vector using Spe1 restriction enzyme with the following reaction mixture:
dsDNA Phage Genome Extraction Protocol
- High Titer Phage Lysate
- 100 mg/mL Lysosome
- 10 mg/mL Rnase A
- DNase 1 (NEB, M0303S)
- 10x DNase 1 Buffer
- 10% SDS
- 10 mg/mL proteinase K
- Phenol:Chloroform:Isoamyl Alcohol (25:24:1)
- Chloroform
- 100% EtOH
- 3 M Sodium Acetate
- 70% EtOH
Instructions
1. In a 2 mL microcetrifuge tube, add the following:
- 738 μL of high titer phage lysate
- 9μL of 100 mg/mL Lysozyme (1 mg/mL final)
- 9μL 10 mg/mL RNase A without EDTA
- 9μL of NEB DNase 1
- 90 μLof 10x DNase 1 Buffer
2. Mix by inversion or pulse vortex, and incubate at 37°C for 90 minutes.
3. Inactivate the DNase by incubating at 75°C for 10 minutes.
4. Let the sample cool to room temp.
5. Add 10μL of 10mg/mL Proteinase K and 50 μL of 10% SDS, and mix by inversion.
6. Incubate at 55°C for 1 hour.
7. Let the sample cool.
8. Add 960μL of Phenol:Chloroform and mix by inverting several times until the sample is cloudy.
9. Centrifuge at 12,000 RPM for 10 minutes at room temperature.
10. Transfer the aqueous layer to a new 2 mL tube (be careful not to aspirate the organic layer).
11. Add an equal volume of chloroform (likely 750 mL) and mix by inverting the tube several times. Using chloroform rather than phenol:chloroform helps remove phenol which can shift the peaks in the spectrophotometer.
12. Centrifuge at 12,000 RPM for 10 minutes at room temperature.
13. Transfer the aqueous layer to a to a new 2 mL tube (roughly 550μL).
14. Add two volumes of 100% ethanol and then 1/10 volume of 3 M Sodium Acetate. Mix by inversion.
15. Incubate at -20°C for roughly 60 minutes. You can leave this overnight if you wish.
16. Centrifuge at 14,000 RPM for 5-10 minutes at room temperature.
17. Carefully decant the supernatant into a waste container.
18. Add 500μL of 70% Ethanol, being careful not to disturb the pellet. Carefully invert the tube once or twice.
19. Centrifuge at 14,000 RPM (or max speed) for 2-3 minutes at room temperature.
20. Carefully decant the supernatant into a waste container, and pipette off any ethanol that remains.
21. Air dry the pellet for for 15 minutes or until no droplets. Do not overdry the pellet or it will be difficult to dissolve.
22. Resuspend the pellet in 50-100μL of DI-H2O or a sufficient volume to dissolve the pellet.
23. Incubate at room temperature for 30 minutes or until it dissolves.
24. Check the DNA concentration in the spectrophotometer, and store the remaining DNA at -20°C.
This protocol has been adapted from a procedure from the lab of Dr. Kristen Parent.