Team:Utrecht/Protocols

BRET Measurement

This protocol can be used to measure the intensity of BRET signal in bacteria expressing a BRET donor-acceptor pair, and the effect different treatments have on the BRET signal. During our experiments, we used bacteria that co-express eYFP::CheY and CheZ::Rluc (biobrick BBa_K608003).
  1. Inoculate bacteria that express the BRET pair in 5 ml LB containing correct antibiotics overnight at 37 degrees.
  2. Add 1.5 ml of the overnight culture to an eppendorf tube and pellet the bacteria by centrifuging them for a minute at max speed.
  3. Remove the supernatant
  4. Add 900 μl of PBS and resuspend the pellet.
  5. Add 100 μl 75 mM coelenterazine (CAS# 77559-48-1)
  6. Measure the emission intensity of both the BRET donor and acceptor proteins in a photospectrometer by performing an emission scan that spans the excitation spectrum of both proteins.*

Excitation Emission
CheZ::RLuc Peak (nm) / 535
Range (nm) 400 600
eYFP::CheY Peak (nm) 512 528
Range (nm) 400-540 500-700
BRET signal Peak / 528
Slit used 5 10
To measure the activity of the Tar receptor:
  1. Measure the emission intensity of both the BRET donor and acceptor proteins in a photospectrometer by performing an emission scan that spans the excitation spectrum of both proteins.* Measure for 30 seconds with an interval of 50 ms.
  2. Pause the measurement and add 10 μl of 50 mM LD-aspartate to the bacterial suspension
  3. Continue the measurements. For every subsequent measurement wash the bacteria in 900 μl PBS, add coelenterazine, and add aspartate.

*alternatively, it is possible to measure the emission of only one of the proteins, since the relative bioluminescence of each protein changes.

Capillary Assay

  1. Inoculate a overnight culture of Dh5-alpha in 5 μl of LB medium.*
  2. Centrifuge the cells at 1667 rfc for 5 minutes, remove supernatant and add 4 ml of HEPES buffer (pH=7.0).
  3. Repeat step 2. Be gentle with the cells since harsh treatment can result in loss of flagella.
  4. Prepare a 96 wells plate by adding 230 μl of bacterial solution to aliquots and sealing the top using parafilm.
  5. Seal the end of one capillary by folding it in the flame of a burner.
  6. Dip the open end of the capillary in the substance to test (or in HEPES buffer in case of the negative control).
  7. Add the capillary into a sealed aliquot of the 96 wells plate.
  8. Incubate for 30 minutes at RT.
  9. Remove the capillary and wash the end open end with miliQ.
  10. Break the sealed end of the capillary.
  11. Add the bulb dispenser to the end of the capillary you just broke and empty the content in 1 ml of 0.9% (w/v) NaCl solution.
  12. Centrifuge for 10 seconds and plate out on agar plates containing 25μg/mL of chloramphenicol.
  13. Incubate overnight.
  14. Count the colonies containing your marker (colonies that are red due to the RFP expression).
*preferably a RFP producing strain as this simplifies the selection procedure at the end.

FRET Measurement

Figure 1: Excitation/emission spectra of CFP and YFP. Adapted from: https://www.thermofisher.com/nl/en/home/life-science/cell-analysis/labeling-chemistry/fluorescence-spectraviewer.html

This protocol can be used to measure the amount of FRET signal in bacteria expressing a FRET donor-acceptor pair, and the effect different treatments have on the FRET signal. Depending on the nature of the FRET pair, conclusions can be drawn with respect to the effect of treatments. During our experiments, we used bacteria with a CheY::YFP and CheZ::CFP expressing plasmid. For measurements we used peak excitation/emission for CFP and YFP (Figure 1.)

  1. Inoculate bacteria that express the FRET pair in 5 ml LB containing correct antibiotics overnight at 37 degrees.
  2. Add 1.5 ml of the overnight culture to an eppendorf tube and pellet the bacteria by centrifuging them for a minute at max speed.
  3. Remove the supernatant
  4. Add 900 μl of PBS and resuspend the pellet.
  5. Measure the light intensity of both the FRET donor and acceptor proteins in a photospectrometer by performing an emission scan for both proteins when they are excited at their peak excitation.*
  6. Measure the light intensity of both the FRET donor and acceptor proteins in a photospectrometer by performing an emission scan for both proteins when they are excited at their peak excitation.*

To just measure FRET signals:

  1. Measure the amount of FRET signal by performing an emission scan for both fluorophores while exciting only the donor fluorophore.
Excitation Emission
CheZ::RLuc Peak (nm) / 535
Range (nm) 400 600
eYFP::CheY Peak (nm) 512 528
Range (nm) 400-540 500-700
BRET signal Peak / 528
Slit used 5 10

To measure FRET changes over time:

  1. Measure the ratio of donor/acceptor emission over time while exciting the Donor fluorophore. Measure for 20 seconds and pause the measurement.
  2. Add 100 μl of aspartate for a final concentration of 500 μM to the sample and continue the measurement. **
  3. Wash your sample between every time you want to add new aspartate measurements.

*We used a Carry Eclipse Fluorescence Photospectrometer from Agilent Technologies. For details on measurements see Table 1
** Work as quick as possible when measuring chemotaxis FRET pairs since this pathway returns to its basal state within 100 seconds .

Agarose Gel (x%)

  1. Measure x g of Agarose
  2. Mix agarose powder with 100 mL 1xTAE in a microwave flask.
  3. Microwave for 1-3 min until the agarose is completely dissolved (but do not overboil the solution, as some of the buffer will evaporate and thus alter the final percentage of agarose in the gel. Many people prefer to microwave in pulses, swirling the flask occasionally as the solution heats up.).
  4. Let agarose solution cool down to about 50 °C (about when you can comfortably keep your hand on the flask), about 5 mins.
  5. Add ethidium bromide (EtBr) to a final concentration of approximately 0.2-0.5 μg/mL (usually about 2-3 μl of lab stock solution per 100 mL gel). EtBr binds to the DNA and allows you to visualize the DNA under ultraviolet (UV) light.
  6. Pour the agarose into a gel tray with the well comb in place.
  7. Place newly poured gel at 4 °C for 10-15 mins or at room temperature for 20-30 mins, until it has completely solidified.
This gel can now be used for Gel Electrophoresis

Annealing Oligonucleotides

  1. Mix the following:
    • 45 μl of each linker strand dissolved in H2O at 10 pmol/μl
    • 10 μl of 10x Annealing buffer
      • 250 mM Tris-HCl, pH 8.0
      • 100 mM MgCl2
  2. Boil a beaker of water. Turn off the flame, and put the tube in the water.
  3. Allow to cool to room temperature.

Use about 0.5 μl for a 10 μl ligation

Colony PCR

  1. Add 50 µl of LB without antibiotic to PCR tubes (one for each Colony PCR).
  2. Pick single colonies from plates.
  3. Incubate at 37°C for 1 hour.
  4. Perform a PCR using 2 µl of the LB containing bacteria (from step 3) as template.

Gel Electrophoresis

  1. Add 10 µL loading buffer to a set amount of your DNA samples
  2. Prepare a casting tray with the right amount and width of combs.
  3. Poor the agarose gel and let it solidify. This takes around 30 minutes for a small and 45 minutes for a larger gel.
  4. In case of DNA isolation, refresh the TAE buffer in the gel box.
  5. Put the casting tray in the gel box. Load 10 µL of GeneRuler 1 Kb DNA ladder in the first lane and load 20 µL of the samples on the gel.
  6. Run the gel at 80-150 V until the dye line is approximately 75-80% of the way down the gel. A typical run time is about 0.5-1.5 hours, depending on the gel concentration and voltage.
  7. Turn OFF power, disconnect the electrodes from the power source, and then carefully remove the gel from the gel box.
  8. Using any device that has UV light, visualize your DNA fragments. The fragments of DNA are usually referred to as ‘bands’ due to their appearance on the gel. Take care that no unprotected body parts are exposed to the UV light.

Gel Extraction

  1. Take gel to the extraction bench.
  2. Cut out bands of correct size.
  3. Add them to eppendorf tubes.
  4. Measure the weight of the extracted bands and add NTI (200μl + (200μl for every 100 mg of gel)).
  5. Incubate at 65°C until the gel completely dissolved (should take 20 minutes).
  6. Add the disolved DNA to a spin column and spin down for 1 minute at max speed (repeat if you cannot use all DNA at once).
  7. Add 700 μl of NT3 and spin down for 1 minute at max speed. Discard the flow through.
  8. Repeat step 7.
  9. Add 15-30 μl of elution buffer and incubate at RT for 2 minutes.
  10. Spin down for 1 minute at max speed.
  11. Measure concentration with Nanodrop.

Gibson Cloning

  1. Use the DNA calculator to calculate the amount of DNA to be used from each fragment. Preferably use a 3:1 insert to vector ratio and make the DNA mix. (mix should have a final volume of 10 μl).
  2. Add 10 μl of Gibson Assembly master mix to the DNA mix.
  3. Place in PCR machine and choose the Gibson assembly program. Run for 1 hrs maximum to obtain the optimal reaction conditions.
  4. Transform the Gibson product if necessary*.
*use 9μl of reaction product in 100μl of competent cells to increase the chance of successful transformation.

Plasmid Purification

  1. Inoculate single colonies from plates in 5 ml LB containing antibiotic overnight in a 37°C shaker (220 rpm).
  2. Transfer 1.5 ml of inoculate to an eppendorf tube and centrifuge for 1 minute at maximum speed.
  3. Discard the supernatant without disturbing the pellet.
  4. Add 150 µl of buffer A1 and vortex to resuspend the pellet.
  5. Add 250 µl of buffer A2 and gently invert 5 times.
  6. Incubate at room temperature for 2 minutes.
  7. Add 350 µl of Buffer A3 and invert until the lysate turns colorless.
  8. Centrifuge for 3 minutes at maximum speed.
  9. Load 700 µl of clear supernatant on spin columns and centrifuge for 1 minute at maximum speed.
  10. Discard the flow-through and add 450 µl of Buffer AQ.
  11. Centrifuge for 30 seconds at maximum speed and discard the flow through.
  12. Centrifuge dry for 30 seconds at maximum speed.
  13. Place the tubes in clean eppendorf tubes and add 50 µl of elution buffer to the membrane.
  14. Incubate at room temperature for 2 minutes.
  15. Centrifuge for 3 minutes at max speed and measure concentration.

PCR

  • Prepare the reaction mix in PCR tubes according to table 1
  • Enzymes should be added last
  • Enzymes should always be kept on the ice block
  • Place the eppendorf tubes in PCR machine and select the correct program
Component Reaction
KOD One-Taq Phusion
H2O Up to 50 µl Up to 20 Up to 50
10x Buffer 5 µl 2 µl 10 µl
10mM dNTPs 5 µl 0.4 µl 5 µl
25 mM MgSO4 3 µl / /
Forward primer 1.5 µl 0.4 µl 2.5 µl
Reverse primer 1.5 µl 0.4 µl 2.5 µl
DNA polymerase1,2 1 µl 0.1 µl 0.5 µl
DNA template 250 ng 250 ng 250 ng

Site Directed Mutagenesis

  1. PCR (see table)
  2. Run 16 cycles of PCR:
    • 95°C - 2 min
      • 95°C - 20 sec
      • 55°C - 10 sec
      • 70°C - 5 min
    • 70°C - 5 min
    • 4°C - 5 min
  3. Add 1 µl DpnI (to digest methylated DNA)
  4. Incubate for 1.5 hrs at 37°C
  5. Transform into bacteria
DNA template 250 ng
10mM reverse primer 2.5µl
10mM forward primer 2.5 µl
5x Phusion buffer 10µl
10mM dNTPs 5µl
Phusion polymerase 0.5 µl
MQ To 50µl

Transformation of Chemically Competent Cells

  1. Get competent cells from the -80°C freezer and thaw them on ice for 10-15 minutes.
  2. Add 50 µL of competent cells to pre-cooled eppendorf tubes. Then, add 2 µL of DNA to the tubes, or as much to get 200 ng in the total volume.
  3. Incubate on ice for 30 minutes.
  4. Heat shock tubes at 42°C for 45 seconds.
  5. Incubate on ice for 5 minutes.
  6. Pipette 500 µL LB medium to each transformation.
  7. Incubate at 37°C for 1 hour, shaking at 200-300 rpm.
  8. Pipette 100 µL of each transformation onto LB-petri plates containing the right antibioticum.
  9. Incubate the plates upside down at 37°C overnight.
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