Team:DTU-Denmark/Lab protocols


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  1. Make a plate design specifying which strains will be in each of the wells in your BioLector plate. It’s highly recommended to run all strains in duplicates. The following controls should be included: strain used for transformation, strains expressing the GFP and/or RFP that will be measured in the other wells.
  2. Make overnight cultures of the strains you want to run in the BioLector.


  • Growth media (LB)
  • Overnight cultures
  • Adhesive Gas Permeable Seal
  • Adhesive Seal for Evaporation Reduction for 48-well plates


  1. Measure OD of the overnight cultures.
  2. Dilute the overnight cultures down to an OD of 0.05.
  3. Add 1.5 mL of diluted culture to each of the 48 wells.
  4. Apply the Adhesive Gas Permeable Seal to the plate and make sure it is stuck on tightly. The seal must be put on precisely, as putting it on crookedly will result in some of the wells not being fully covered.
  5. Apply the Adhesive Seal for Evaporation Reduction on top of the Gas Permeable Seal.
  6. Place the BioLector plate carefully into the BioLector in the specified slot until it clicks in.
  7. Using the Start Assistant in the BioLector, set the temperature and humidity to be held during the experiment. The filters to be used for measurements of each well in the experiment should also be set, as well as the sampling time (3 mins per filter used).
  8. Run the BioLector as long as needed. The measurements of the biomass and amount of FPs in each well can checked while the BioLector is still running. It is recommended to run the experiment at least until the cells in each well reach stationary phase.
  9. Use the following settings:
    • 37 °C
    • Fluorescence gain of 80
    • Biomass excitation 620 nm (light scattering)
    • GFP filter was 486nm (ex) / 510nm (em)



  1. Calculate your ligase concentration by estimation of DNA concentration from your restriction gel.
  2. Which vector to use?


  1. Prepare ligation mix with a total volume of 20 μl
    • 2 μl 10x Buffer
    • Parts to be ligated: vector+insert 1:5, DNA conc. less than 50 ng
    • fill up with water
    • 1 μl ligase
    • control (no insert)
  2. Add ligase and keep mix at RT for 1h.
  3. To inactivate the ligase place your mix at 65ºC for 10 min.
  4. Store on ice.

Before starting on the ligation(s), always run your digestion products after you’ve performed a clean-up/purification on a gel to verify you actually have been successful in your digestions.
If you want to be very precise with your DNA-content calculations, when running the gel you can load the DNA-ladder sequentially with a difference of 2-fold dilution.
If your digestions were successful and you are happy with the way your gel looks, you are set to start your ligation calculations.
Firstly, make sure you have a DNA-ladder illustration indicating the mass of DNA at different positions on the gel, then compare the intensity of bands of your digested products with the intensity and position of the bands from the DNA-ladder(s).

  • The same intensity of bands indicates they must have the same mass of DNA.
After you have determined how much DNA (mass) you have of each band, you can calculate the DNA content on a mole basis by dividing the mass with the length of the fragment. In general a ligation-mix should look like this:
  • Usually a ligation mix should have a total volume of 20 μl, although if you require additional amounts of the same ligation, make a duplicate of the ligation you are trying to make.
  • The buffer needed is simply called “10x ligation buffer”, and being 10 times concentrated, you need to add 1/10th of the total volume of buffer. Hence if the total volume is 20 μl, you need to add 2 μl of your 10x ligation buffer.
  • The amount of DNA added should conform to the calculations you made from the gel you ran with your digested products. A rule of thumb should be that you take more insert than vector, this way increasing your chances of actually getting a ligation. The ratio of insert to vector should be 5:1 and the total amount of DNA should not exceed 50 ng.
  • The two remaining items to be added into your mix will be ddH2O and Ligase. The amount of Ligase enzyme to be added is only 1 μl, so you know how much DNA (insert+vector), Ligase and the buffer to add; the rest of the volume (→ 20 μl) should be accounted for with ddH2O.
  • Remember though that Ligase should only be added in the end, because once added into your mix its going to get to work and you don’t want it playing around with your dna before you have added everything else. Another important thing to remember is that the Ligase enzyme cannot be taken from the freezer to your lab-bench; you have to instead add your Ligase into your mix by working at the freezer.
  • Controls are always important and good indications if things are working the way they should be, so a good control with ligations is a negative control, that being just vector with no insert, the rest of the recipe being the same. This should hopefully give you no colonies as it would indicate your digested vector is in fact digested and didn’t re-ligate. If there are colonies on your negative control plate, it will give you a good indication of the background noise on your actual ligation plates.

PCR product purification


  • 1 vol PCR product
  • 2 vol NT Buffer
  • 600 µl NT3 Buffer
  • 300 µl Elution Buffer NE


  1. Mix 1 volume of sample with 2 volumes of NT buffer in an 1,5 ml Eppendorf tube.
  2. Place a column into a 2 ml collection tube and load the sample.
  3. Centrifuge at 11.000 g for 1 min.
  4. Discard flow through and place the column back into the collection tube.
  5. Add 600 µl NT3 buffer and centrifuge at 11.000 g for 1 min.
  6. Discard flow through and place the column back into the collection tube.
  7. Centrifuge at 11.000 g for 2 min to remove NT3 buffer. Discard flow through.
  8. Place the column into a clean 1,5 ml Eppendorf tube.
  9. Add 30 µl Elution Buffer NE and incubate at RT for 1 min to increase the yield of eluted DNA.
  10. Centrifuge at 11.000 g for 1 min.

Plasmid purification by miniprep (Zymo Research Group)


  • 2 ml cell culture
  • 600 ul TE Buffer
  • 100 µl 7X Lysis Buffer
  • 350 µl (cold) Neutralization Buffer
  • 200 µl Endo-Wash-Buffer
  • 400 µl Zyppy Wash Buffer
  • 50 µl Zyppy Elution Buffer


  1. Spin down 2 ml of cell culture in a 2 ml Eppendorf tube at 11.000 g for 5 min.
  2. Remove supernatant, spin again down for 10 seconds, remove supernatant (if the pellet is not sufficient, repeat step 1 in the same tube).
  3. Resuspend your pellet in 600 ul TE buffer.
  4. Add 100 µl 7X Lysis Buffer (blue) and mix by inverting the tube 4-6 times. Proceed to the next step within 2 minutes.
  5. Add 350 µl cold Neutralization Buffer (Yellow) and mix carefully. The sample will turn yellow and a yellowish precipitate will occur, then the reaction is finished. Invert the tube an additional 2-3 times to ensure complete neutralization.
  6. Centrifuge at 11,000 g for 5 min.
  7. Transfer the supernatant (ca. 875 µl) into a column. Avoid disturbing the cell debris pellet.
  8. Place the column into a collection tube and centrifuge for 30 seconds.
  9. Discard the flow-through and place the column back into the same collection tube.
  10. Add 200 µl Endo-Wash-Buffer to the column and centrifuge for 30 seconds.
  11. Add 400 µl of Zyppy Wash Buffer to the column and centrifuge for 30 seconds.
  12. Transfer the column into a clean 1,5 ml Eppendorf tube and then add 50 µl Zyppy Elution Buffer directly to the column. Let the product(s) stand for 1-15 minutes on the table at RT.
  13. Centrifuge for 30 seconds to elute the plasmid DNA.



  • 25 µl DNA (depending on your DNA conc)
  • 10 µl FD-buffer
  • 1 µl (2.5 µl can also be used) restriction enzyme (1 & 2) NOTE: REs should be kept on ice.
  • 63 ul (60 ul when 2.5 µL RE is used) ddH2O

Enzyme selection for BioBricks digest

A BioBricks part:


Vector with upstream part - S,P

Downstream Insert - X,P

Upstream Insert - E,S

Vector with downstream part- E,X

E= EcoRI X= XbaI S= SpeI P= PstI


  1. Turn the thermo block on 30 min before use.
  2. Mix everything - the enzymes should be added last, giving a total volume of 100 µl.
  3. Leave for 2 hours at 37°C in thermo block/incubator.
  4. Deactivate the enzyme by leaving it at 65°C in a thermo block for 10 min.
  5. Keep on ice until its cooled down.
IMPORTANT: If you only use one RE on your plasmid, SAP treatment is required:
  1. Mix 50µL of the restricted DNA with 2.5µL SAP and 5µL SAP-buffer.
  2. Put it in the incubator 37°C for 60 min.
  3. 15 minutes on heat block 65°C to deactivate enzyme.
  4. Ice.

Transformation protocol


NOTE: The materials should be kept on ice.

  • 5X 1 ml LB media
  • Cuvettes
  • 50 µl electrocompetent cells
  • 1 µl ligated plasmid with insert(s)


  1. Add 50 µl electrocompetent cells to the cuvette.
  2. Add 1 µl ligated plasmid to the cuvette and make sure that it is mixed thoroughly with the cells without creating bobbles.
  3. Have 1 ml LB media ready in a pipette.
  4. Wipe the cuvette with tissue to ensure that the metal are free of water.
  5. Insert the cuvette in the electroporator and press ‘ Pulse’.
  6. Quickly add the LB media to the cuvette and transfer as much as possible back to the 5 ml tube.
  7. Incubate for 1 hour at 37°C.
  8. Plate out in duplicates (20 and 200 µl) on LB-plates containing your favorite antibiotic.