Team:Vilnius-Lithuania/Protocols

Protocols

Agarose Gel preparation (x%)

  1. To prepare agarose gels we used TopVision Agarose Tablets that are manufactured by Thermo Fisher Scientific.
  2. Add appropriate number of agarose tablets to the electrophoresis buffer based on the table below to prepare your desired gel percentage

    3. Note: Use a flask that is 2 to 4 times the volume of the solution being prepared.



  3. Before heating soak tablets in a buffer (~ 4 minutes) until tablets completely break into fine-particle slurry. Swirl the slurry to break up any remaining particles. Important: Ensure tablets break up entirely. Heating will render non-dispersed agarose particles insoluble.

    4. Note: Heating times are dependent on the volume of liquid and number of gel tablets to dissolve.

  4. Remove the flask from microwave, swirl gently to dissolve any remaining agarose particles.
  5. Reheat on high power for 1-2 minutes or until the solution is clear and all particles are dissolved.
  6. Remove the flask from the microwave oven, and gently swirl.
  7. Cool the solution to approximately 50-60 °C.
  8. 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). Mix well.
  9. Pour the gel into a tray of required size and place in the well comb, let the gel cool and solidify for 10-15 mins at room temperature.
  10. This gel can now be used to run electrophoresis gels.

    11. Note: To let the gel cool down to the required temperature of about 60 °C one can use a smaller volume container, which is not affected by heat to let a smaller volume of the melted agarose cooldown faster.

PCR

This protocol has been taken from Thermo Scientific and is as follows:
Pipette these items in order listed



*Optionally 5X Phusion GC Buffer can be used. See section 4.2 for details.
**The recommendation for final primer concentration is 0.5 μM, but I can be varied in a range of 0.2-1.0 μM, if needed.
*** Addition of DMSO is recommended for GC-rich amplicons. DMSO is not recommended for amplicons with very low GC % or amplicons that are > 20 kb.

Cycling instructions:



Note: The initial denaturation temperatures as well as annealing temperatures, and extension times are both primer and polymerase dependent, therefore, must be looked up before planning the cycles.

Annealing oligos for lengthier primers

This protocol was taken from Addgene

  1. Place the mixed oligos in a 1.5mL microfuge tube.
  2. Place tube in 90-95°C hot block and leave for 3-5 minutes.
  3. Then, gradually cool to 25°C over 45 minutes.

Colony PCR

  1. Prepare a PCR Master Mix as mentioned in the previous entry noted PCR.

    2. Note: For the possibility of some of the Master Mix volume being lost when dividing the huge stock into PCR tubes make sure to add 10% extra of all the components to the Master Mix as compensation.

  2. Make sure to use final primer concentrations of about 0.4 μM and run 35 cycles of the PCR.
  3. Transfer the Master Mix in 20 μL or 50 μL quantities to small PCR tubes.
  4. Take the dish with bacterial colonies intended for testing.
  5. Transfer a colony that has not yet merged with any other colonies to another plate where numbered and separated regions are marked with marker and afterwards use the same instrument used for transfer and shake the tip that touched the bacteria inside the aliquot of the Master Mix that you have previously pipetted into the PCR tubes.
  6. Repeat for the number of desired colonies.
  7. Place the second dish in the optimal temperature of the bacteria and incubate.
  8. Place the PCR tubes in the PCR thermocycler and run the program adequate to the polymerase used.

    9. Note: If the results have proven to be successful the already transferred colonies can be used for further experiments as they bacteria have usually grown enough for transfers into liquid media for overnight growth.

Gel electrophoresis

  1. Removed the comb from the already cast gel.
  2. Place the gel into the electrophoresis apparatus and make sure that the volume of buffer is sufficient for the electrophoresis, if not add the required amount (TBE or TAE).
  3. Add the DNA size marker or ladder (use about 3-5 μL of the ladder).
  4. After the gel’s wells have been submerged begin loading the samples.
  5. Load 8-50 μL of the sample depending on the purpose of the gel.
  6. Adjust the voltage to 120V.
  7. Set the required time for running the gel.

    8. Note: 20 minutes are sufficient for a gel with already incorporated Ethidium Bromide.

  8. Place the electrodes with the cover over the gel apparatus and initiate the electrophoresis.
  9. After the time has passed turn off the machine.
  10. Take out the gel and place it over a UV or Blue-light illuminator to visualize the EtBr intercalated DNA either for confirmation of experimental results or gel excision.

Gel extraction

  1. Take the gel out of the electrophoresis machine.
  2. To visualize the DNA prior to extraction one cannot use UV illuminators due to the possibility of mutation caused by UV irradiation, a compromise is a Blue-light illuminator in a dark room, which is sufficient enough for accurate excision of the gel.

    3. Note: If the possibility of avoiding UV irradiation is out of the question, please try to keep the exposure of the gel to UV under 10 seconds to avoid damaged DNA.

  3. Excise the bands with the DNA of interest keeping the excess of gel to a minimum as it decreases the yield of DNA after cleanup.
  4. Afterwards follow the Thermo Scientific GeneJET Gel Extraction and DNA Cleanup Micro Kit, which is listed below as follows:
  5. Excise up to 200 mg gel slice containing the DNA fragment using a clean scalpel or razor blade. Cut as close to the DNA as possible to minimize the gel volume. Place the gel slice into a 1.5 mL tube.
  6. Add 200 μL of Extraction Buffer. Mix thoroughly by pipetting.
  7. Incubate the gel mixture at 50-58°C for 10 minutes or until the gel slice is
    8. completely dissolved. Mix the tube by inversion every few minutes to facilitate the melting process. Ensure that the gel is completely dissolved.
  8. Add 200 μL of ethanol (96-100%) and mix by pipetting.
  9. Transfer the mixture to the DNA Purification Micro Column preassembled with a collection tube. Centrifuge the column for 30-60 seconds at 14,000 × g. Discard the flow-through. Place the DNA Purification Micro Column back into the collection tube.

    10. Note:
    1. If DNA fragment is ≥ 10 kb centrifuge the column for 2 minutes at 14,000 × g.
    2. Close the bag with DNA Purification Micro Columns tightly after each use!

  10. Add 200 μL of Prewash Buffer (supplemented with ethanol, see p. 3) to the DNA Purification Micro Column and centrifuge for 30-60 seconds at 14,000 × g. Discard the flow-through and place the purification column back into the collection tube.

    11. Note: If DNA fragment is ≥ 10 kb centrifuge the column for 1-2 minutes at 14,000 × g.

  11. Add 700 μL of Wash Buffer (supplemented with ethanol, see p. 3) to the DNA Purification Micro Column and centrifuge for 30-60 seconds at 14,000 × g. Discard the flow-through and place the purification column back into the collection tube.

    12. Note: If DNA fragment is ≥ 10 kb centrifuge the column for 1-2 minutes at 14,000 × g.

  12. Repeat step 7.
  13. Centrifuge the empty DNA Purification Micro Column for an additional
    14. 1 minute at 14,000 × g to completely remove residual Wash Buffer.

    Note. This step is essential to avoid residual ethanol in the purified DNA solution. The presence of ethanol in the DNA sample may inhibit downstream enzymatic reactions.

  14. Transfer the DNA Purification Micro Column into a clean 1.5 mL microcentrifuge tube (not included).
  15. Add 10 μL of Elution Buffer to the DNA Purification Micro Column. Centrifuge for 1 minute at 14,000 × g to elute DNA.

    16. Note.

    • If DNA fragment is ≥ 10 kb the elution volume should be increased between 15-20 μL to get optimal DNA yield. Elution volume less than 10 μL is not recommended.
    • Lower volume of Elution Buffer for DNA Micro Kit can be used (6-10 μL) in order to concentrate eluted DNA. Please notice that <10 μL elution volume slightly decreases DNA yield.
  16. Discard the purification column and store the purified DNA at -20°C.

Enzymatic reaction cleanup

This protocol was taken from Thermo Scientific and it is as follows:

  1. Adjust the volume of the reaction mixture to 200 μL with Water, nuclease-free or TE buffer (not included).
  2. Add 100 μL of Binding Buffer. Mix thoroughly by pipetting.
  3. Add 300 μL of ethanol (96-100%) and mix by pipetting.
  4. Transfer the mixture to the DNA Purification Micro Column preassembled with a collection tube. Centrifuge the column for 30-60 seconds at 14,000 × g. Discard the flow-through. Place the DNA Purification Micro Column back into the collection tube.

    5. Note.
    1. If DNA fragment is ≥ 10 kb centrifuge the column for 2 minutes at 14,000 × g. 2. Close the bag with DNA Purification Micro Columns tightly after each use!

  5. Add 700 μL of Wash Buffer (supplemented with ethanol, see p. 3) to the DNA Purification Micro Column and centrifuge for 30-60 seconds at 14,000 × g. Discard the flow-through and place the purification column back into the collection tube. Note. If DNA fragment is ≥ 10 kb centrifuge the column for 2 minutes at 14,000 × g.
  6. Repeat step 5.
  7. Centrifuge the empty DNA Purification Micro Column for an additional 1 minute at 14,000 × g to completely remove residual Wash Buffer.

    8. Note. This step is essential to avoid residual ethanol in the purified DNA solution. The presence of ethanol in the DNA sample may inhibit downstream enzymatic reactions.

  8. Transfer the DNA Purification Micro Column into a clean 1.5 mL microcentrifuge tube (not included).
  9. Add 10 μL of Elution Buffer to the center of the DNA Purification Micro Column membrane. Centrifuge for 1 minute at 14,000 × g to elute DNA.

    10. Note. *If DNA fragment is ≥ 10 kb the elution volume should be increased to 15-20 μL to get optimal DNA yield.
    *Lower volume of Elution Buffer for DNA Micro Kit can be used (6-10 μL) in order to concentrate eluted DNA. Please notice that < 10 μL elution volume slightly decreases DNA yield.
    *Double the elution volume or perform two elution cycles when purifying larger amounts of DNA (for example > 5 μg).

  10. Discard the purification column and store the purified DNA at -20°C.

DNA digestion

This protocol is taken from Thermo Fisher Scientific and is as follows:

  1. Prepare the reaction mixture at room temperature in the order indicated.



  2. Mix gently and spin down.
  3. Incubate at 37°C in a heat block or water thermostat for 5 min. ***
  4. Inactive the enzyme (optional). ***

    5. Note:
    *The volume of water should be corrected to keep the indicated total reaction volume. The volume of DNA can be scaled up to 10 μl or down to 0.5 μl depending on the DNA concentration.
    **Only 2 μL of 10X FastDigest® buffer is required for unpurified PCR product in a 30 μL reaction volume.
    ***See the Certificate of Analysis for enzyme and substrate specific incubation time and enzyme inactivation conditions.


Ligation

This protocol is taken from Thermo Scientific and is as follows:

  1. Thoroughly mix the 5X Rapid Ligation buffer prior to use.
  2. Add the following to a microcentrifuge tube:
  3. Vortex and spin briefly to collect drops
  4. Incubate the mixture at 22°C for 5 min.
  5. Use 2-5 μl of the ligation mixture for transformation.


    6. Note:
    *The reaction mixture can be stored at 0-4°C until used for transformation. Prior to electroporation, chloroform extract the ligation mixture and use 1 μl for the electroporation reaction.
    *For the incubation of the mixture, it is recommended to keep the ligation mixture in room temperature for about 1 hour to ensure successful and high-yielding ligation results.


Chemical transformation

  1. Take out bacteria out of the -80°C freezer and immediately place under ice to let them fully thaw.
  2. Turn on the UV sterilization function in the working box for about 15 minutes to sterilize the working environment before opening the test tube with the bacteria to avoid any risk of contamination.
  3. Take out petri dishes with required antibiotic out of the refrigerator.
  4. Mix 50 μl with 10 μl of the ligation reaction mixture in the sterile box and then further incubate on ice for 5-10 minutes.
  5. Prepare as many of transformation controls as possible: transform the bacteria without DNA (contamination control), only the linearized digested vector (to check if the DNA is properly digested), positive control with purified non-digested DNA (to check whether everything is all right with competent cells and the transformation).
  6. Heat up 2 thermoblocks with 42°C and 37°C.
  7. Perform a heat shock for 1 min on 42°C.
  8. Incubate on ice for 5-10 minutes.
  9. Suspend the bacteria with 500-1000 μl of LB (Luria Broth).
  10. Incubate in the 37°C heat block for 20-30 minutes.
  11. Centrifuge the cells for 5 minutes at 3000rpm.
  12. Discard the supernatant in one quick movement, there should be some of the LB medium still left, resuspend the bacteria in the remaining LB medium and plate out on a LB-petri plate with the correct antibiotic.
  13. Incubate the plates overnight at the optimal temperature for selected bacteria.

LIC cloning

This protocol was taken form Thermo Scientific and is as follows:

  1. To generate the necessary 5' and 3' overhangs on the purified PCR template, prepare the following reaction mixture at room temperature:



  2. Vortex briefly and centrifuge for 3-5s.
  3. Incubate the reaction mixture at room temperature (20-25°C) for 5 min.

    4. Note. Do not exceed 5 min.

  4. Stop the reaction by adding 0.6 μL of 0.5M EDTA, mix well.

    Note. Store the prepared PCR product in the reaction mixture at -20°C if the annealing step with the LIC vector cannot be performed immediately. Thaw and mix carefully prior to performing the annealing reaction.

  5. Set up the annealing reaction:
    Add 1 μL pLATE, LIC-ready vector (60 ng, 0.02 pmol DNA) to the T4 DNA polymerase treated PCR product prepared in steps 1–3 of this protocol. Vortex briefly and centrifuge for 3-5 s.
  6. Incubate the annealing mixture at room temperature (20-25°C) for 5 min.

    7. Note. Annealing is complete within 5 min of incubation. Reactions can be incubated up to 2 hours without affecting results. Longer incubation times do not improve efficiency.

  7. Use the annealed mixture directly for bacterial cell transformation.

Protein Electrophoresis (SDS Page)

Separating gel

  1. Prepare the gel casts in holders. Fill the holder with water to check for leakage after 5 min. After leakage check, pour all water from gel casts and dry residual with paper towel.
  2. To prepare separation-gel, mix the reagents listed in the table below with the proper volumes to receive a gel with desired composition.
  3. NOTE! Add TEMED and APS last, solidification of the gel will occur when these reagents are added.
  4. Pipette gel mixture into gel casts up to a height approximately 1 cm below the gels comb.
  5. Fill the remaining cast with isopropanol (if not available use distilled water).
  6. Allow to harden for 30-60 min.
  7. Pour all isopropanol (or water) from gel casts and dry residual with paper towel.

    8. *Ammonium Persulfate (APS) - 20 mg/mL
    *Acrylamide 30 % solution
    *TEMED
    *Separating Gel Buffer - 1.5 M Tris-HCl pH 8.8 with 0.4% SDS. Stored at 4°C.
    *Stacking Gel Buffer - 0.5 M Tris-HCl pH 6.8 with 0.4% SDS. Stored at 4°C



    Stacking gel

    1. To prepare stacking-gel, in a tube mix the reagents listed in table 4 below with the proper volumes to receive a gel with desired composition.
    2. NOTE! Add TEMED and APS last, solidification of the gel will occur when these reagents are added.
    3. Pipette gel mixture on top of separating gel to fill cast and remove all air bubbles.
    4. Fill the remaining cast with isopropanol (if not available use distilled water).
    5. Insert the comb whilst the gel is still in liquid form. The comb will form wells to load samples when the gel turns solid.
    6. Allow to harden for 45-60 min.

    Electrophoresis



    Running the gel

    1. Mount the gels into the tank, remove combs and fill the inner chamber of the tank with 1x running buffer to the top. Fill a third of the volume in the outer chamber of the tank with 1x running buffer.
    2. Pipette 2 µl ladder and 10 µl sample into each well.
    3. Connect power pack (red-red, black-black) and run at 100 V, 60 min or until the blue line almost reaches the bottom of the gel.
    4. Once electrophoresis has finished, remove gels from the glass plates.
    5. Carefully wash all equipment used for electrophoresis with water.

    10x Running buffer

    Dissolve 30.0 g of Tris base, 144.0 g of glycine, and 10.0 g of SDS in 1000 mL of H2O. The pH of the buffer should be 8.3 and no pH adjustment is required. Store the running buffer at room temperature and dilute to 1X before use.

    Coomassie Blue Staining

    1. Prepare the staining solution containing 0.1 % Coomassie Blue in 40 % ethanol, 10 % acetic acid.
    2. After electrophoresis, incubate 1 or 2 gels in a staining container containing 100 mL Coomassie Blue staining
      3. solution.
    3. Incubate at room temperature for 1 hour until bands are visible.
    4. Decant the stain and rinse the gel once with deionized water.
    5. Prepare 100 mL destaining solution containing 10 % ethanol and 7.5 % acetic acid.
    6. Gently shake the gel at room temperature on an orbital shaker until the desired background is achieved.

    Chemically competent cell preparation

    Required: Ice, sterile 1.5 mL tubes (label before starting), pre-cooled centrifuge, 100 mM CaCl2, glycerol, both sterile, 85 mM CaCl2 with 15% glycerol.

    1. Inoculate a 5 mL overnight-culture in LB-medium without antibiotics (for DH5a) or with appropriate antibiotics (for other strains).
    2. Grow over night at 37 ˚C at 250 rpm.
    3. Inoculate 100 mL LB-medium with 1/100 volume of the overnight culture and incubate to an OD600 = 0.5 - 0.6 at 37 °C and 230 rpm.
    4. Transfer cells to two sterile 50 mL falcon tubes and incubate on ice for 20 min.
    5. From this step on, always keep cells cold.
    6. Centrifuge for 5 min at 4500 rpm and 4 °C (pre-cool centrifuge) and discard the supernatant.
    7. Carefully suspend each pellet in 20 mL of ice-cold 100 mM CaCl2 and incubate on ice for 1 h.
    8. Centrifuge for 5 min at 4500 rpm and 4 °C (pre-cool centrifuge) and discard the supernatant.
    9. Carefully suspend each pellet in 2 mL ice cold 85 mM CaCl2 containing 15% Glycerol.
    10. Pipette 50 - 200 µL aliquots in prepared, pre-cooled 1.5 mL tubes (work on ice), shock-freeze in liquid nitrogen or on dry ice and store at -80 °C.

    His-tagged protein purification using magnetic beads

    Dynabeads™ His-Tag Isolation and Pulldown magnetics beads were purchased from Invitrogen (Thermo Fisher Scientific). The following purification steps were performed based on manufacturer recommendations.



    * Note that the 2X Binding/Wash Buffer needs to be diluted to 1X concentration prior to use. After centrifugation, cultivated cells, harboring the protein of interest were resuspended in 1X Binding/Wash Buffer.

    1. Thoroughly resuspend the Dynabeads™ magnetic beads in the vial (vortex >30 sec or tilt and rotate 5 min).
    2. Transfer 50 μL (2 mg) Dynabeads™ magnetic beads to a microcentrifuge tube. Place the tube on a magnet for 2 min. Aspirate and discard the supernatant. Add your sample (prepared in 1X Binding/Wash Buffer) to beads. Mix well.
    3. Incubate on a roller for 5 min at room temperature (or colder if the protein is unstable at room temperature). The incubation time may be increased up to 10 min.
    4. Place the tube on the magnet for 2 min, discard the supernatant.
    5. Wash the beads 4 times with 300 μL 1X Binding/Wash Buffer by placing the tube on a magnet for 2 min and discard the supernatant. Resuspend the beads thoroughly between each washing step.
    6. Add 100 μL His-Elution Buffer. Incubate the suspension on a roller for 5 min at room temperature (or colder if the protein is unstable at room temperature).
    7. Apply on the magnet for 2 min and transfer the supernatant containing the eluted histidine-tagged protein to a clean tube.

    Double stranded SELEX (using magnetic beads/antibodies)

    1. The initial single-stranded oligonucleotide library was chemically synthesized (IDT).
    2. To amplify the DNA pool, six 50 μL PCR reactions were set up using 0.1 μM of the synthetic pool oligonucleotide as template, 2 μM of each primer, nuclease-free water and Platinum SuperFi PCR Master Mix. Amplified DNAs were purified using GeneJET PCR purification kit (Thermo Fisher Scientific).

    Using magnetic beads

    1. Thoroughly resuspend the Dynabeads™ magnetic beads in the vial (vortex >30 sec or tilt and rotate 5 min).
    2. Transfer 50 μL (2 mg) Dynabeads™ magnetic beads to a microcentrifuge tube. Place the tube on a magnet for 2 min. Aspirate and discard the supernatant. Add your sample (prepared in 1X Binding/Wash Buffer) to beads. Mix well.
    3. Incubate on a roller for 10 min at room temperature.
    4. Place the tube on the magnet for 2 min, then discard the supernatant.
    5. Wash the beads 2 times with 200 μL 1X Binding/Wash Buffer, containing 2 mM MgCl2, 1 mg/mL BSA, and 10% glycerol. Resuspend the beads thoroughly between each washing step.
    6. Resuspend beads in 50 µL of Binding/Wash buffer, containing 2 mM MgCl2, 1 mg/mL BSA, 0.25 µg/µL Herring sperm DNA and 50 ng of double stranded oligonucleotide library. Incubate suspension at room temperature for 1.5h with gentle shaking.
    7. Wash beads four times with 200 µL of Binding/Wash buffer, containing 2 mM MgCl2, 0.1 mg/mL BSA (3-4 min each wash).
    8. Add 10 µL of DNA elution buffer containing 10 mM Tris-HCl (pH 8) and 0.5 mM EDTA. Expose the mixture to blue light for 15 minutes with gentle shaking.
    9. Collect the beads and transfer the suspension to a clean tube for PCR amplification.

    Using antibodies

    1. Coat the wells of a Nunc Maxisorp ELISA plate with an antibody (6x-His Tag Antibody (Invitrogen)) at 1–10 μg/mL concentration in carbonate/bicarbonate buffer (pH 9.6). Cover the plate and incubate overnight at 4°C.
    2. Remove the coating solution and wash the plate twice by filling the wells with 200 μL 50 mM Tris-HCl, 100 mM NaCl and 0.05% Tween 20. The solutions are removed by flicking the plate over a sink. The remaining drops are removed by patting the plate on a paper towel.
    3. Block the remaining protein-binding sites in the coated wells by adding 200 μL blocking buffer (Tris-HCl/BSA) per well. Cover the plate and incubate for at least 1–2 h at room temperature or overnight at 4°C.
    4. Wash the plate twice with 200 µL wash buffer (50 mM Tris-HCl, 100 mM NaCl, and 0.05% Tween 20).
    5. Add 100 µl of protein lysate (of various dilutions). Incubate for 2 h at room temperature.
    6. Remove samples and wash the plate twice with 200 μL washing buffer.
    7. Add 30 µl of double-stranded oligonucleotides (of various dilutions), incubate for 1 h at room temperature.
    8. Wash the plate twice with 200 μL washing buffer.
    9. Add 10 µl of Tris-HCl. Expose plate to blue light for 15 min.
    10. Transfer the suspension to clean tubes for PCR amplification.

    Fluorescent EMSA

    1. Prepare a binding reaction: EMSA binding reaction mixture consists of Tris binding buffer (20 mM Tris-HCl, 5 mM MgCl2, 0.1 mM EDTA, 6% sucrose, 50 mM KCl and 1 mM DTT, pH 7.5), 12 ng of double-stranded oligonucleotides and purified protein of interest (5–50 μg); final volume 30 µl. Incubate samples for 2 h at 4°C.
    2. Transfer samples to 6% native TBE gel; run the electrophoresis for 2 h at 100V.
    3. For visualization, stain polyacrylamide gel with GelRed fluorescent stain. Result analysis can be performed using transilluminator.

    Fluorescence measurement using plate reader

    1. Transform bacteria with plasmids containing your target genes (Day 1).
    2. Pick 2 colonies from each plate and inoculate in 5-10 mL LB medium + Chloramphenicol (Day 2).
    3. Grow cells overnight (16-18 hours) at 37 °C and 220 rpm (Day 2).
    4. Make a 1:10 dilution of each overnight culture in LB with Chloramphenicol (0.5 mL of culture into 4.5 mL of LB with Chloramphenicol (Day 3).
    5. Measure Abs600 of these 1:10 dilutions (Day 3).
    6. Record the data in your notebook (Day 3).
    7. Dilute culture further to your target Abs600 of 0.02 in a final volume of 12mL LB medium with Chloramphenicol in a 50 mL Falcon tube (Day 3).
    8. Take 500 μL samples of the diluted cultures at 0 hours into 1.5 mL eppendorf tubes, prior to incubation. At each time point between 0 hours and 6 hours, you will take a sample from each of the 8 devices, two colonies per device, for a total of 16 eppendorf tubes with 500 μL samples per time point, 32 samples total (Day 3).
    9. Place the samples on ice (Day 3).
    10. Incubate the remainder of the cultures at 37 °C and 220 rpm for 6 hours (Day 3).
    11. Take 500 μL samples of the cultures at 6 hours of incubation into 1.5 mL eppendorftubes. Place samples on ice (Day 3).
    12. At the end of the experiment, you should have two plates to read. You will have one plate for each time point: 0 and 6 hours. On each plate you will read both fluorescence and absorbance.
    13. Samples should be laid out according to the plate diagram below



    14. Pipette 100 μL of each sample into each well. From 500 μL samples in a 1.5 mL eppendorf tube:

      15. *4 replicate samples of colony #1 should be pipetted into wells in rows A, B, C and D *Replicate samples of colony #2 should be pipetted into wells in rows E, F, G and H

    15. Be sure to include 8 control wells containing 100 μL each of only LB with Chloramphenicol on each plate in column 9, as shown in the diagram.
    16. Set the instrument settings as those that gave the best results in your calibration curves (no measurements off scale). If necessary, you can test more than one of the previously calibrated settings to get the best data (no measurements off scale).
    17. Instrument temperature should be set to room temperature (approximately 20-25 C) if your instrument has variable temperature settings.

    Western Blot

    Solutions

    30 %acrylamide – bis solution
    30 g acrylamide and 0,8 g N, N’-methylenebisacrylamide are solvated in 100 mL deionized H2O. The solutions is filtered and stored in +4°C, no longer than 30 days.

    1,5 mol/l TRIS-HCl, pH=8,8
    18,15 g TRIS dissolved in 70 mL H2O; pH is corrected with HCl. Solution is diluted to 100 mL, filtrated and stored in +4°C, in the dark for no longer than 30 days.

    0,5 mol/l TRIS-HCl, pH=6,8
    3 g TRIS are dissolved in 35 mL H2O, pH is adjusted using HCl. The solution is diluted to 50 mL, filtrated and stored in +4°C, in the dark for no longer than 30 days.


    10 % SDS (NDS) solution
    5 g SDS is dissolved and diluted to 50 mL. The solution is stored in room temperature.

    10 % APS (ammonium peroxide sulphate)
    25 mg APS is dissolved in 250 μL H2O. Solution must be used during the day of preparation.

    5 x “Loading” buffer solution
    Each of these components: 350 μl 0,5 mol/L TRIS-HCl (pH=6,8), 150 mL glycerol, 50 mg SDS, 46 mg DTT (), 10 μL bromophenol blue are to be dissolved in H2O one after the other. The solution is frozen. Protein samples are heated up with this sample, and before loading in to the gel – cooled down.

    PBS (phosphate buffer solution) 10x
    1L: 80,06g NaCl; 2,01g KCl; 11,4g Na2HPO4 (Na2HPO4*7H2O – 21,6g; Na2HPO4*12H2O – 29,0g); 2,04g KH2PO4. Dissolve in 800 mL deionized H2O in the order that the components are mentioned. Dilute to 1L. Pipette in 100mL portions and store in +4°C.

    Protein transfer buffer solution
    100mL: 300mg TRIS, 1,12g glycine, 10mL methanol. Dilute with H2O to 100mL. (400mL is the amount to be made)

    Blocking solution
    PBS with 2% milk powder.

    Tween dilution
    5 mL concentrated Tween dilute to 50mL. 1L PBST 10mL diluted Tween (Tween 10%/10x)

    PBS-T
    PBS su 0,1 % Tween-20.

    Chloronapthol solution
    1 tablet chloronaphtol + 10 mL methanol. For membrane resolution use 10mL PBS 2ml chloronaphtol dissolved in methanol 30μL 30% H2O2.

    ELECTROPHORESIS - IMMUNOBLOTTING

    Electrophoresis

    1. Everything done in this step is covered in the SDS page step.
    2. The protein from the polyacrylamide gel are transferred onto the PVDF membrane (Polyvinylidene fluoride). The membrane is cut to be nearly the same size as the gel, as well as same sized filter papers are prepared.
    3. After electrophoresis the polyacrylamide gel is submerged in the protein transfer solution. The membrane is washed with methanol and as well as the gel is soaked in the transfer solution, same is done with the filter papers.
    4. Then perform the semi-dry fractionated protein transfer onto the membrane. On the protein transfer apparatus’ cathode base a piece of soaked filter paper is placed, afterwards the membrane, gel, and lastly, another layer of filter paper. Everything is pressed down using the anode base and for 40 minutes the protein transfer is performed, the current is adjusted based on the area of the gel. (Length cm x width cm = area cm2 mA usually ~40mA) (always check if there is an already set protein transfer current based on the lab you are working in).
    5. The membrane is washed with PBST and afterwards blocked with PBS + 2% milk powder solution for 60 minutes, room temperature (can be stored in +4oC overnight).
    6. The membrane is washed 4 times (for ~5min) with PBST. The antibody solution is prepared: PBST 2% milk powder. If the hybridoma medium is used, it can be diluted 1:1-1:20 overall solution volume is ~10mL [50μl hybridoma medium]. The membrane is incubated with the antibodies for 1h in room temperature.
    7. Membrane is washed 6 times with PBST. The secondary antibody solution is prepared with 2% milk powder PBST. The conjugate is diluted 1:1000 – 1:5000 (usually. 1:4000), based on the manufacturer’s recommendations. Incubate for 1h in room temperature.
    8. Membrane is washed for 4 times with PBST, incubating 2 – 3 times for 5 min. on a shaking platform. Afterwards wash 2 – 3 with distilled water.
    9. The membrane with chloronaphtol solution, and only if nothing is seen or is barely resolved, used TMB – blotting substrate for the peroxidase (ready-to-use). (before TMB wash the membrane with water).
    10. The membrane is washed with water, dried, and scanned. Due to the light the peroxidase substrate blurs out in 3-7 days.
      11.  

      Immunoenzymatic assay

      Required solutions:

      PBS (Phosphate buffer solution)
      1l: 80,06g NaCl; 2,01g KCl; 11,4g Na2HPO4 (if Na2HPO4*7H2O – 21,6g; Na2HPO4*12H2O – 29,0g); 2,04g KH2PO4. Solvate in 800 mL deionized H2O, solvate components in the order described one after the other. Dilute to 1L. Aliquot into 100 mL portions and store in +4°C.
      Only 1x PBS is used. 100mL 10x PBS are diluted to 1L distilled H2O.
      PBST
      1x PBS with 0,1% Tween (10%/10x)
      “Coating buffer” (0,05 M Potassium Carbonate buffer solution, pH 9,5).
      Dissolve 4,2g NaHCO3 in 90mL distilled H2O, regulate to pH=9,5 with 20% NaOH and dilute to 1l with distilled H2O. Store in +4°C.
      Blocking buffer solution
      PBS with 2% BSA.
      TMB ELISA
      TMB Stop solution
      3,6 % H2SO4

      Method:

      1. Antigen immobilization (Protein sorption). A 5μg/mL antigen solution is prepared in the immobilization buffer solution. 50μl is aliquoted into the wells. The plate is incubated for 1h. +37oC / incubated overnight in +4oC.
      2. Blocking. The free surface of the plate is blocked with albumin. 150 μL of buffer solution is pipetted into the wells, incubated for 1h in room temperature
      3. Sample antibody solution preparation. The media of interest samples are diluted with PBST. Before adding the medium, the plate should be washed with water. After addition of the medium, incubate for 1h in room temperature
      4. Incubation with marked antibodies. Plate is washed 4 times with 50:50 PBST: H2O solution. Conjugate is diluted in 1:5000 PBST solution. This solution is transferred in to the wells in 50 μL portions. Incubate for 1h in room temperature.
      5. Reaction resolution. The plate is washed 6 times (4 with 50:50 PBST: H2O and 2 with H2O and incubate afterwards). 50μL of TMB ELISA is added to each well and is kept for 5 min in the dark. The reaction is stopped with 3,6% H2SO4 25μL. Optic density is measured with a spectrophotometer, with the light wavelength of 450nm (alignment 620nm).
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