Team:TU Munich/lab/notebook/methods

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Contents


Cloning

Preperative digests were performed using 1 µg of Mini-DNA with subsequent gel purification. Gel slices were cut using 365 nm illumination as short as possible and a picture was taken afterwards. Slices were purified using Pormega's Wizard purification Kit. Ligations were done with 50 ng vector, T4 Ligase, a total volume of 20 µl and different molar insert:vector ratios. In case the vector is smaller than the insert, more vector is used. If both parts are approximately equal in size, a 1:1 ratio is mostly the best choice. Incubation of ligations were conducted over night (o/n) at 16 °C. Vector and insert controls were added for purity-information of samples. On the next day, ligations were precipitated using ethanol/glycogen and resuspended in 10 µl nuclease free water. The whole sample was transformed by electroporation into competent DH5 alpha cells.

Restriction digest

  1. Add 1 µg of DNA to be digested, and adjust with dH20 to a total volume of 43 µl.
  2. Add 5 µl of NEBuffer 4 to the tube.
  3. Add 1 µl of your first enzyme.
  4. Add 1 µl of your second enzyme.
  5. There should be a total volume of 50 µl. Mix by pipetting carefully and eventually pull down drops on the surface of the tube.
  6. Incubate the restriction digest at 37 °C for 60 min, and then 80 °C for 20 min to heat kill the enzymes.

Gel purification of digested DNA

Use 0.7% - 3% agarose gels (depending on sample size) in 1x TAE buffer to seperate DNA. Stain gels using SYBR Gold solutions (dilute 1:10000 in 1x TAE buffer) for 20 min and cut out correct gel slices using 365 nm illumination with a new scalpel for every sample. Purify Gel slices using Promega's Wizard purification Kit according to manufacturer's instructions.

Ligation

For a final volume of 20 µl:

  1. Mix water, vector and insert in appropriate I:V ratios.
  2. Add 2 µl 10x ligase buffer and 1 µl T4 ligase. Mix by pipetting carefully and eventually pull down drops on the surface of the tube.
  3. Incubate samples o/n at 16 °C.

Glycogen/ethanol precipitation

  1. Add 2 µl 3 M sodium acetate (or 2 M sodium chloride, or 5 M ammonium acetate), 0.5 µl glycogen (to a final concentration of 0.05 to 1 µg/µl) and 22.5 µl isopropanol (1 volume of isopropanol or 2.5 volumes of ethanol) to 20 µl ligation product.
  2. Incubate the mixture at -20 °C for one hour.
  3. Centrifuge the mixture for 15 min at 18,000 rpm at 4 °C.
  4. Discard the supernatant.
  5. Add 50 µl ice cold 70 % ethanol.
  6. Centrifuge the mixture for another 10 min at 18,000 rpm at 4 °C.
  7. Remove supernatant with a pipet CAREFULLY. If the pellet moves, centrifuge again.
  8. Air-dry the pellet until the ethanol is completely evaporated. Do not over-dry pellet as this makes dissolving more difficult.
  9. Resuspend the pellet in 10 µl nuclease-free water by pipetting carefully.

Note: The pellet is very small and hardly visible, but it is present! Make sure you have enough light to see it. Orienting all tubes in exactly the same direction in the centrifuge also helps, because if you find a pellet in one tube, you know that it has to be in the same spot in all other tubes

Transformation of purified ligations

10 µl purified ligation was mixed with 40 µl electrocompetent DH5 alpha cells and electroporated according to the protocol of the manufacturer. 1 ml SOC medium (SOB medium + 20 mM glucose) was used and incubation was carried out for 1h at 37°C, shaking at 500rpm in 2 ml eppis. Culture was then centrifuged at 4000 rpm for 5 min, and resolved in 100 µl medium. Whole sample was plated onto LB/agar plates supplemented with appropriate antibiotica.


Testing

Quantification

GFP Assay

  1. Cells are transferred to M63 medium, because LB shows slight fluorescence itself and diluted down to a low OD600 of about 0.5.
  2. Different concentrations of IPTG are used for induction.
  3. The measurement using the plate reader occured over night, with heating the plate to 37 °C and shaking it between measurements.

Filters used for flourescence measurements: excitation filter: 485 nm; emission filter: 520 nm.

Measurement of OD 600 to monitor cell density.

Miller Assay

induction of lacZ expression with IPTG or light, depending on construct

Start of Assay:

  1. Measurement of Abs(600 nm) in plate reader. The volume of bacterial suspension should be equal in all wells, ideally 170 µl.
  2. Take a new eppi with 0.5 ml of Zbuffer + 20 μl of freshly prepared 0.1% SDS + 40 μl of Chloroform (under fume hood) + 500 µl sample solution (e.g. 430 µl medium + 20 µl bacterial suspension).
  3. Mix the solution by vortexing for 10 sec (all samples with equal vortexing time).
  4. Let the chloroform settle down. This takes about 5 min. If tube still contains blurred solution, centrifuge for 1 min at room temperature at 4000 rcf.
  5. Transfer 100 μl of supernatant to 96 well plate (for photometer).
  6. Initiation of assay with 20 μl of ONPG (4 mg/ml), mix well, NOTE START TIME.
  7. Incubation at 37 °C.
  8. Stop reaction with 50 μl of 1 M Na2CO3, mix well, NOTE STOP TIME.
  9. Measurement of OD (420 nm) and OD (550 nm).

Convert file to .txt files for analysis in Excel (sort data in this order: OD600-raw data, OD600-corrected data, OD420-raw data, OD420-corrected data, OD550-raw data, OD550-corrected data).

Calculate Miller values according to the following:

[[File:TU_Munich_2011_Equation_Miller_Assay.jpg]]

Miller Units = 1000 * (ABS420 - (1.75 * ABS550)) / (time [min] * volume [ml] * ABS600)

  1. Time t = time of incubation between START with ONPG solution and STOP with Na2CO3
  2. Volume V = volume of cell suspension

Qualitative tests

S-gal plates

For 500 ml:

  1. 0.5l Milli Q water
  2. 7.5 g AgarAgar
  3. 10 g LB-Pulvermix
  4. 460 mg ferric ammonium sulfate (do NOT dissolve s-gal prior to addition; just add the powder: 150 mg S-Gal )
  5. 100 µg/ml Antibiotics

Put everything together and autoclave.

Normally 250 mg of ferric ammonium citrate (M = 261,97 g/ml) are used, but here it is replaced by ferric ammounium sulfate (M = 482,25 g/mol).

You don't need to keep s-gal out of the light it is heat and light-INSENSITIVE. Do not store prepared s-gal plates longer than 14 days!


Additional protocols

Quick Ligation Protocol Quick T4 DNA Ligase

Quick Ligase M2200L Buffers: Quick Ligation Reaction Buffer 2x

Protocol

  1. Combine 50 ng of vector with a 3-fold molar excess of insert. Adjust volume to 10 μl with dH2O.
  2. Add 10 μl of 2X Quick Ligation Buffer and mix.
  3. Add 1μl of Quick T4 DNA Ligase and mix thoroughly.
  4. Centrifuge briefly and incubate at room temperature (25°C) for 5 minutes.
  5. Chill on ice, then transform or store at -20°C.

Do not heat inactivate. Heat inactivation dramatically reduces transformation efficiency.

note: Quick ligation didn't work well for us! Ligations worked when using approach above.

Transformation

  1. 1 µl of desired plasmid (better: 0.5 µl in order to avoid arcing) was added to a tube containing 40 µl electrocompetent ells.
  2. The solution was gently mixed while steering with pipette tip. The mixture was transferred into a chilled cuvette (1 mm gap) and immediately inserted in the electroporation device.
  3. Electroporation was performed using 1510V.
  4. Directly after electroporation, 1 ml of room temperature SOC-medium was added per cuvette.
  5. The solution was mixed and transferred into 1ml Eppendorf tube for incubation at 37 °C, 200-300 rpm and 1-2 h.
  6. Afterwards, 100 µl and 100 µl concentrated (spun down and resuspended in 400ul) of each transformation reaction was spread on LB plates containing the appropriate antibiotics and raised over night at 37 °C.
  7. On the next day, colonies were picked and inoculated into 5 ml LB medium supplied with the appropriate antibiotics. This is either done VERY early in the morning, or late in the evening (in order to continue the following day).
  8. Next, the plasmid can be prepped.

Miniprep (Metabion mi-plasmid mini prep kit)

ADD RNase TO "CELL RESUSPENSION SOLUTION" AND 100% EtOH TO "COLUMN WASH BUFFER"!!!

All centrifugation steps are performed at 13.000 rpm and RT!

  1. Take 1.5 to 2 ml (max. 5 ml) bacterial overnight LB culture
  2. Centrifuge for 1 min
  3. Discard the supernatant completely (Removing all liquid at this step is critical!)
  4. Add 250 μl of Cell Resuspension Solution and resuspend the pellet completely by vigorously vortexing
  5. Add 250 μl of Cell Lysis Buffer to the cell suspension (If the Lysis Buffer stock solution is precipitated, heat to 55°C for 5 min to dissolve -> RT)
  6. Invert 5 times to mix. Do not vortex, as this will cause chromosomal DNA contamination of the plasmids! Incubation < 5 min (usually 3 min)
  7. Add 350 μl of DNA Binding Buffer
  8. Invert 5 times to mix (Do not vortex, see step 6)
  9. Centrifuge for 10 min
  10. Transfer all of the clear liquid supernatant to a spin column which has been set into a 2 ml collection tube
  11. Centrifuge for 1 min
  12. Add 600 μl of Column Wash Buffer to the spin column
  13. Centrifuge for 1 min
  14. Repeat steps 12 and 13 to make sure, no Column Wash Buffer is left in the column
  15. Place the spin column in a new 1.5 ml Eppi
  16. Elute the plasmid DNA by adding 50 μl of autoclaved water directly onto the middle of the white membrane of the spin column. Incubate for 1 min.
  17. Centrifuge for 1 min
  18. Directly use or store at -20°C

Preparing electro competent cells

  1. 20 ml LB medium was inoculated with 1 µl electrocompetent DH5alpha
  2. 350 ml LB medium were inoculated with 15 ml of the overnight culture and incubated at 37°C until a OD(600) of 0,7 was reached
  3. the medium was distributed on falcons and the cells were centrifuged at 4500 rpm for 20 min at 4°C
  4. after discarding the supernatant 350 ml of sterile ice-cold 10% glycerol was added and the pellets were resuspended using a pipet
  5. the resuspension was centrifuged at 4500 rpm at 4°C for 11 minutes
  6. after discarding the supernatant the cells were resuspended again in 350 ml of ice-cold 10% glycerol (cells kept chilled all the time)
  7. and again centrifuged at 4500 rpm at 4°C for 11 minutes
  8. the supernatant was discarded and the cells were resuspended in the remaining supernatant
  9. the cells were dispensed in 40 µl aliquots and stored at -80°C for further use

Glycerin Stocks

A mixture of 50 % LB and 50 % Glycerin is prepared and autoclaved. 300 ul of this solution are added to 700 ul cells, in order to get a about 10 - 15 % glycerin solution supplied with the cells to be frozen. The solution is then frozen at -80 °C.

Squeeze N Freeze

  1. Electrophorese the DNA sample in an agarose gel (TAE or TBE), then stain with an appropriate reagent, e.g., ethidium bromide or SYBRTM Green I.
  2. Using a clean razor blade, carefully excise the band of interest. Trim excess agarose from all six sides of the DNA band to maximize recovery and purity.
  3. Chop the trimmed gel slice and place the pieces into the filter cup of the Quantum Prep Freeze ‘N Squeeze DNA Gel Extraction Spin Column. Place the filter cup into the dolphin tube. If the volume of your trimmed gel slice is too great to fit into one filter cup, then use two or more and pool the recovered samples at the end of the protocol.
  4. Place the Quantum Prep Freeze ‘N Squeeze DNA Gel Extraction Spin Column (filter cup nested within dolphin tube) in a -20° C freezer for 5 minutes.
  5. Spin the sample at 13,000 x g for 3 minutes at room temperature.
  6. Collect the purified DNA from the collection tube; the agarose debris will be retained within the filter cup of the Quantum Prep Freeze ‘N Squeeze DNA Gel Extraction Spin Column. The DNA is ready to use for PCR, ligations, labeling or other enzymatic reactions. Ethanol precipitation is recommended for applications requiring a more concentrated sample and will also have the effect of further purifying the sample.

note: Not the best gel extraction method. Better use spin columns!

Transforming Chemically Competent Cells

Based on http://openwetware.org/wiki/Transforming_chemically_competent_cells

  1. Thaw chemically competent cells on ice.
  2. Add DNA, pipette gently to mix (1μl of prepped plasmid is more than enough).
    • Note: If you are adding small volumes (~1μl), be careful to mix the culture well. Diluting the plasmid back into a larger volume can also help.
  3. Let sit for 30 minutes on ice.
  4. Incubate cells for 30 seconds at <math>42^o</math>C.
  5. Incubate cells on ice for 2 min.
  6. Add 1 mL SOC at room temp.
  7. Incubate for 1 hour at <math>37^o</math>C on shaker.
    • Note: Can save some time here by reducing incubation to ~45 min.
  8. Spread 100-300 μl onto a plate made with appropriate antibiotic.
  9. Grow overnight at 37 °C.
  10. Save the rest of the transformants in liquid culture at 4 °C. If nothing appears on your plate, you can spin this down, resuspend in enough medium to spread on one plate and plate it all. This way you will find even small numbers of transformants.

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