Team:UNebraska-Lincoln/Experiments

UNL 2017

Helping reduce methane emissions from livestock

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EXPERIMENTS



›Agar Plate Preparation

Materials

  • 24 petri dishes
  • 500 mL LB Agar
  • 500 µL cm34

Procedure

  1. Microwave 500 mL of LB Agar for 15 min on power level 3.
  2. Repeat step 1.
  3. Put LB Agar in the 55°C water bath until the temperature of the LB Agar equalizes to 55°C.
  4. Pipette 500 µL of antibiotic into the LB Agar and mix well.
  5. Label the bottom of the empty petri dishes with the name of the antibiotic.
  6. Pour LB Agar into petri dishes. Make sure Agar covers the entire bottom of the dish.
  7. Flame the petri dishes.
  8. Let them solidify at room temperature and then store in the 4°C.



›Cell Lysis and Protein Purification

Materials

  • 1M K2HPO4
  • 1M KH2PO4
  • 1M NaCl
  • Autoclaved H2O
  • 1M Imidazole
  • 1.5 mL Eppendorf Tubes
  • 20% ethanol
  • Laemmli Sample Buffer (Bio Rad)
  • Coomassie Brilliant Blue stain (Bio Rad)

Procedure

  1. Prepare a 5 mL starter culture of E. coli and let it incubate at 37°C for 16 hours
  2. Transfer 50 µL of the starter culture into 50 mL of LB broth and 50 µL antibiotic
  3. Let 50 mL culture incubate at 37°C for 24 hours
  4. While waiting for the cultures to grow prepare an SDS page and buffers under the specifications below. Adjust pH range between 7-8
    Lysis Buffer

    20mM Imidazole

    50mM imidazole

    Elution Buffer (500mM imidazole)
  5. Centrifuge culture for 10 min at 23°C at 5,000rpm.
  6. Weigh the cell wet mass. Then resuspend by pipetting lysis buffer into the solution in a 1:4 ratio of cell wet mass to volume of lysis buffer.
  7. Keep everything on ice for the rest of the procedure.
  8. Clean the rod of the sonicator with autoclaved H2O and then ethanol.
  9. Insert the rod as close as you can to the bottom of the tube without touching the bottom of the tube.
  10. Sonicate the cell suspension for 4 minutes using 10 seconds of sonication and 50 second pause intervals.
  11. Take the nanodrop protein concentration at 280 absorbance of the cells after they have been lysed.
  12. Dilute the Total Fraction(lysed cells) to make 20 µL of 10 mg/mL solution. Set this aside for the SDS page.
  13. Centrifuge the rest of the Total for 20 min at max speed at 4°C.
  14. While waiting for the centrifuge equilibrate the column by using a 5:1 ration of water to column in terms of volume.
  15. Slowly pipette this volume of water onto the edge so the column is not disturbed.
  16. Pipette into the column a 3:1 ratio of lysis buffer to column in terms of volume.
  17. Once the centrifuge is finished dilute the Total Soluble to make 20 µL of 10 mg/ml solution. Set this aside for the SDS page.
  18. Run all of the Total Soluble fraction through the column and collect the Flow Through in 1.5 mL eppendorf tubes.
  19. Run 2 mL of the 20mM imidazole wash buffer through the column and collect in 1.5 mL eppendorf tubes.
  20. Run 2 mL of the 50mM imidazole wash buffer through the column and collect in 1.5 mL eppendorf tubes.
  21. Run 2 mL of the Elution Buffer through the column and collect in 1.5 mL eppendorf tubes.
  22. Dilute the flow through tubes to make 20 µL of 10 mg/mL solution. Set this aside for the SDS page.
  23. Take 20 µL from each of the leftover from purification and set it aside for the SDS page.
  24. Store leftover samples in the -20°C freezer.
    To prepare solutions for the SDS page, make mixtures shown in the table below.
  25. Boil tubes in Thermal Cycler for 10 min.
  26. Centrifuge tubes to get rid of bubbles.
  27. Load wells of the SDS page with 15 µL of each solution.
  28. Fill well farthest to the left and farthest to the right with 5 µL of ladder.
  29. Run the gel for 30 min at 50 amps.
  30. Make sure that gel ran very close to the bottom (but not off the gel), and then wash the SDS page in water.
  31. Microwave the SDS page for 10 seconds.
  32. Rock until the blue line at the bottom disappears then stain the gel with coomassie blue, microwave for ten more seconds and cover with tinfoil.
  33. Let gel stain overnight and take an image in the morning.



›Colony PCR

Materials

  • PCR tubes
  • Autoclaved Water
  • Cm34 Agar plate
  • Taq DNA polymerase
  • Primers
  • Thermocycler

Procedure

  1. Fill 8 PCR tubes with 10 µL of autoclaved.
  2. Get an unused cm34 agar plate and label the back with the numbers 1-8.
  3. Take the pipette and swipe up one of the colonies you want to analyze and then gently swipe the pipette tip onto the empty plate on top of the corresponding number. Be careful not to swipe all of the colony onto the plate. Repeat this for all eight colonies.
  4. Transfer the colony into the PCR tube. Pipette up and down to resuspend the cells.
  5. Boil the PCR tubes for 10 minutes in the thermocycler.
  6. Create a taq DNA polymerase mixture in a 1.5 mL eppendorf tube as such.
  7. Take 1 µL of the first boiled sample and put it into a fresh PCR tube.
  8. Pipette 25 µL of the Taq DNA polymerase mix into the fresh PCR tube. Do this for the next eight colonies.
  9. Start the Polymerase Chain Reaction in the thermocycler using the specifications defined below.
  10. Load the samples into a gel and perform gel electrophoresis as defined by the protocol to test if the PCR was successful.



›Enzymatic Assay of Bromoperoxidase

Materials

  • MOPS Buffer
  • NaOH
  • Ethanol
  • Monochlorodimedone
  • Potassium Bromide
  • Hydrogen Peroxide
  • Cell Lysate Solution

Procedure

    The entire procedure should be performed at 25°C with all reagents at pH 6.4.


  1. Prepare reagents A-E as Specified below.
    • 50 mM MOPS Buffer, pH 6.4 at 25°C (Prepare 250 ml in deionized water using MOPS, free acid, hydrate, and adjust to pH 6.4 at 25°C with 1 M NaOH.)
    • 100% (v/v) Ethanol (Use Ethyl Alcohol, Anhydrous, 200 Proof)
    • 0.1 mM Monochlorodimedone with 200 mM Potassium Bromide (MCD) (Prepare by dissolving 1.2 gram of Potassium Bromide, in 50 ml of Reagent A. Then dissolve 21.75 mg of Monochlorodimedone, in 0.5 ml of Reagent B. Add 20 µl of the MCD solution to the KBR solution.)
    • 100 mM Hydrogen Peroxide (H2O2) (Prepare by diluting 0.284 ml of Hydrogen Peroxide, 30% (w/w) solution, to 25 ml with Reagent C.)
    • Cell Lysate solution(Incubate at 25°C for 30 minutes prior to use)
  2. Pipette (in milliliters) the following reagents into the wells of a UV transparent microplate.
  3. With a microplate reader record the decrease in A290nm for several minutes until constant then add 20 µL of hydrogen peroxide to each well to catalyze the reaction.
  4. Record the decrease in A290nm for several more minutes. Obtain the A290nm/minute using the maximum linear rate for both the samples and controls.

NOTE= This procedure was adapted from a sigma aldrich protocol. (https://www.sigmaaldrich.com/content/dam/sigma-aldrich/docs/Sigma/General_Information/2/bromoperoxidase.pdf) We used a microplate reader instead of cuvettes and a spectrophotometer to measure change in absorbance. Because of this 400 µL total instead of 1 mL total. Ratios of reagents are the same just modified to be 400 µL total volume.




›Gel DNA Recovery

Materials

  • Zymoclean™ Gel DNA Recovery Kit
  • ADB Buffer
  • Wash Buffer

Procedure

  1. Weigh the gel, then add 3 volumes of ADB buffer to each volume of gel. (max volume in tube = 750 µL)
  2. Incubate the 1.5 mL eppendorf tube in a 55°C water bath for 5-10 minutes.
  3. Use vortex to mix well.
  4. Add the melted agarose solution into a Zymo-spin column and place into a collection tube.
  5. Centrifuge for 30 seconds at 16,000 g. Empty the collection tube whenever necessary.
  6. Add 200 µL of wash buffer to the column and centrifuge for 30 seconds.
  7. Repeat step six.
  8. Centrifuge at max speed for 2 minutes.
  9. Place Zymo-Spin column into a new 1.5 mL tube.
  10. Add 6-10 µL of autoclaved H2O directly to the column matrix.
  11. Centrifuge at 16,000 g for 1.5 minutes to elute the DNA.



›Gel Electrophoresis

Materials

  • Agarose powder
  • TAE buffer
  • Erlenmeyer flask
  • Ethidium Bromide
  • Casting tray
  • Sub-Cell apparatus
  • 1kb ladder
  • 6x Loading Dye

Procedure

  1. Weigh 0.5 grams of agarose powder.
  2. Measure 50 mL of TAE Buffer into an erlenmeyer flask and add in the agarose powder. Mix well.
  3. Cover the erlenmeyer flask and microwave for two minutes at power level ten.
  4. Let the solution cool for ~10 minutes.
  5. Add 4 µL of ethidium bromide and mix well.
  6. Pour the solution into a casting tray and let it cool for 30 minutes.
  7. Set up the Sub-Cell apparatus and fill with enough TAE buffer to cover the gel.
  8. Remove the comb.
  9. Load 5 µL of the 1 kb ladder into the leftmost well.
  10. Add 5 µL of 10x loading dye into your sample and then load this into a well.
  11. Once your samples are loaded place the lid on the apparatus and run the gel at 100 volts for 1 hour.



›LB Agar Preparation

Materials

  • LB Broth powder (Lennox)
  • 500 mL media bottle
  • LB Agar powder
  • 500 mL nanopure water

Procedure

  1. Weigh 12.5 grams of LB Broth powder and transfer to the 500 mL media bottle.
  2. Weigh 7.5 grams of LB Agar powder and transfer to the 500 mL media bottle.
  3. Add 500 mL of nanopure water into the media bottle.
  4. Mix well.
  5. Autoclave media bottle on cycle 2.



›LB Broth Preparation

Materials

  • LB Broth powder (Lennox)
  • 250 mL media bottle
  • 250 mL nanopure water

Procedure

  1. Weigh 6.25 grams of LB Broth powder.
  2. Add LB Broth powder into a 250 mL media bottle.
  3. Pour 250 mL of nanopure water into the media bottle.
  4. Mix well.
  5. Autoclave media bottle on cycle 2.



›Nitrite Metabolism Testing Procedure

Note: In order to simplify the procedure, the symbol N will be used to designate the total number of samples being tested. For example, if five different nitrite concentrations are being tested, N for that experiment would be 12, because there would be five samples containing the various concentrations for both the control and experimental cultures, along with a 0 mmol samples for both cultures. For clarity see Table 1 below:


Materials

Day 1
  • Solid NaNO2 (Total required mass is calculated for experiment conditions)
  • 150 mL Erlenmeyer Flask
  • 50 mL Graduated Cylinder
  • LB Media
  • 20% Aqueous Glucose
  • Serum Vials, Rubber Stoppers, and Metal Caps
  • Serum Vial Crimper
  • Nitrogen Source and Tube
  • 1 mL Syringe Needles
  • Ring Stand, Clamps
  • Four Culture Tubes
  • Chloramphenicol (1 ug/mg)
Day 2
  • 96 Well Plate
  • Chloramphenicol (1 ug/mg)
  • Syringes and Needles
Day 3
  • Syringes
  • Nessler’s Reagent

Procedure

Day 1
Nitrite Solution Preparation
  1. In separate eppendorf tubes, prepare 100 µL of sodium nitrite solution for each concentration that will be tested in the experiment. The final volume of each anaerobic culture, including the added nitrite solution, will be 10.6 mL.
Vial Preparation
  1. In a 150 mL Erlenmeyer flask, combine 146.25 mL of LB media with 3.75 mL of 20% aqueous glucose in sterile conditions. LB media can be measured using a 50 mL graduated cylinder.
  2. Obtain N of each of serum vials, rubber stoppers, and metal caps
  3. Transfer 10 mL of the previously prepared culture into each vial, along with 100µL of the corresponding nitrite solution.
  4. Seal each vial with a rubber stopper.
  5. Using a vial crimper, crimp a metal cap onto each vial in order to seal the vials shut. Make sure to label this cap to keep track of the nitrite concentration present.
  6. Dispose of the remaining media in the Erlenmeyer flask accordingly.
Sparging
  1. Obtain two 1 mL syringe needles, a ring stand, and two clamps. The syringes themselves are not necessary, only the needles.
  2. Fix one clamp onto the ring stand towards the base. Put a serum vial in the gripping end of the clamp and tighten. Make sure the base of the vial is resting on the base of the stand.
  3. Mount the other clamp higher on the ring stand. This will hold the nitrogen tube.
  4. Attach one of the syringe needles to a nitrogen line. This attachment will vary depending on the nitrogen source setup present in each lab.
  5. Use the other syringe needle to vent the serum vial. This needle is perforated through the rubber stopper. DO NOT PUT THE TIP OF THIS NEEDLE INTO THE MEDIA PRESENT IN THE VIAL. This will lead to expulsion of media from the vial onto the surroundings.
  6. The serum vial is now vented and securely mounted to the table, and a syringe needle is attached to the nitrogen tube. Slowly begin to turn on the nitrogen source until the desired flow rate is reached. Very little flow is required for sparging. After the desired flow is reached, push the nitrogen needle into the vial. Put this tip as far into the liquid as possible so it will bubble through the media. Adjust flow as necessary until a steady stream of bubbles is present.
  7. Each vial needs to be sparged for ten minutes. At the end of each sparging, remove the nitrogen source needle first, and then the vent needle. When starting a new vial, insert the vent needle before the nitrogen needle.
  8. After all vials have been sparged, shut off the nitrogen supply, dispose of the needles in the sharps container, and disassemble the rest of the setup.
  9. After sparging, autoclave the vials using the liquid cycle to ensure any potential contaminants are destroyed.
Creating Aerobic Cultures
  1. Obtain four culture tubes for this portion of the experiment.
  2. In sterile conditions, add 5 mL of LB media with 5 uL of chloramphenicol (1 µg/mg).
  3. The first two tubes are both for control cultures. Transfer a single colony from a culture plate into each tube, again in sterile conditions.
  4. Repeat the previous step for the experimental cultures.
  5. Incubate these cultures overnight in a 37℃ shaker incubator.
Day 2
Optical Density of Aerobic Cultures
  1. Obtain an OD reading for all four aerobic cultures. Blank with water. A 10:1 dilution is recommended, but other dilution factors are acceptable as long as performed consistently.
  2. Select the control and experimental cultures with the higher OD reading to perform the rest of the experiment.
Creating Anaerobic Cultures
  1. Obtain the control and experimental aerobic culture tubes with the higher OD reading. Add 5.6 µL of chloramphenicol antibiotic to each.
  2. Obtain N syringes and syringe needles.
  3. Syringe 0.5 mL of culture into each vial using a different syringe for each vial. Make sure to add the correct culture in the correct vial (control vs experimental).
  4. Incubate the vials overnight in the 37℃ shaker incubator.
Day 3
Optical Density of Anaerobic Cultures
  1. Obtain N Eppendorf tubes.
  2. Extract approximately 1.2 mL of each aerobic culture using a different syringe. Put each of these samples into a separate Eppendorf tube.
  3. From each Eppendorf tube, pipet 20 µL of culture into the desired well plate. Perform OD reading in the same fashion as the aerobic cultures.
Isolation of Supernatant
  1. Centrifuge the Eppendorf tubes made in the previous section at approximately 20000 g for three minutes.
  2. Pre-weigh N new large Eppendorf tubes.
  3. Transfer 1 mL of supernatant from each sample into the corresponding pre-weighed Eppendorf tube. Discard the remaining supernatant and pellet in a biohazard waste container.
Nessler’s Test
  1. 0.5 mL of Nessler’s Reagent is required for each supernatant sample. Obtain an adequate amount (0.5 mL * N) of reagent in a disposable culture tube.
  2. Add 0.5 mL of the reagent into each supernatant sample tube. Close the lid and invert 5x for each sample to ensure complete reaction of the reagent with the ammonia present in the sample.
  3. Centrifuge the samples at approximately 20000 g for 10 minutes.
  4. After centrifuge, remove as much supernatant as possible from each sample using a separate pipet tip and discard in a heavy-metal waste container. Be careful to not remove any precipitate.
  5. Centrifuge again highest rpm for 10 minutes.
  6. Again, remove any remaining supernatant using a separate pipet tip for each sample and dispose in the heavy metal waste container.
  7. Weigh each sample and subtract the initial mass of the pre-weighed Eppendorf tube. Leave in the fume hood for 24 hours, and re-weigh. Continue to re-weigh every 24 hours until the masses stabilize. Record the final stabilized masses and subtract the initial corresponding container masses.
  8. Using proper technique, clean up any waste in the appropriate manner. Ensure that all items that came into contact with Nessler’s reagent are disposed of in a heavy metals waste container.



›Plasmid Mini-Prep

Materials

  • Thermo Scientific GeneJET Plasmid Miniprep Kit
  • 1.5 mL eppendorf tube x4
  • Column and Matrix x2
  • Wash buffer
  • Zyppy wash buffer
  • Lysis buffer
  • Neutralization buffer

Procedure

  1. Centrifuge cell culture for 10 min at 23°C at 5,000rpm.
  2. Pour out the LB Broth Supernatant into bleach.
  3. Resuspend cell pellet in 600 µL of autoclaved H2O.
  4. Transfer 600 µL of the cell pellet into a 1.5 mL eppendorf tube.
  5. Add 100 µL of lysis buffer to the eppendorf tube.
  6. Invert tube 2-4 times and lyse for 2 minutes.
  7. Put 350 µL of neutralization buffer into the tube and continue to invert tube until all of the blue color disappears and a yellow precipitate forms.
  8. Centrifuge tube at 21,000g for 5 minutes.
  9. Transfer the supernatant into the zymo-spin column.
  10. Place the column into a collection tube and centrifuge at 16,000 g for 15 seconds. Discard the flow-through and place the column back into the same collection tube.
  11. Add 200 µL of Endo-Wash buffer to the column. Centrifuge at 16,000 g for 15 seconds.
  12. Add 400 µL of Zyppy-wash buffer to the column. Centrifuge at 16,000 g for 30 seconds.
  13. Centrifuge for two minutes at 21,000 g.
  14. Transfer the column into a clean 1.5 mL microcentrifuge tube then add 30 µL of Autoclaved H2O directly to the column matrix.
  15. Let it stand for one minute at room temperature.
  16. Centrifuge at 16,000 g for 1.5 minutes to elute the DNA.



›Polymerase Chain Reaction

Materials

  • PCR Tubes
  • DNTP’s
  • 10x PCR buffer
  • MgSO4
  • Template DNA
  • Primers
  • KOD Polymerase
  • Autoclaved Water
  • Thermocycler

Procedure

  1. For one 25 mL reaction mix the materials stated below into a PCR tube.
  2. Place the reaction tube into the thermocycler and set it based on the specifications below.



›Restriction Enzyme Digest

Materials

  • Restriction enzymes
  • Cutsmart Buffer
  • DNA insert
  • Plasmid

Procedure

  1. Prepare 14 µL of uncut DNA insert or plasmid in PCR tube.
  2. Add 2 µL of Cutsmart buffer.
  3. Add 2 µL of each desired restriction enzyme (i.e. XbaI and SpeI).
  4. Gently mix together.
  5. Place in thermocycler for 1½ hours at 38°C.
  6. Run gel electrophoresis and perform gel DNA recovery on product.
  7. Store in 4°C chamber.



›Restriction Enzyme Digest Cloning

Materials

  • T4 Ligase
  • Ligase buffer
  • Restriction enzymes
  • PCR tubes
  • Autoclaved Water

Procedure

  1. Perform restriction enzyme digest on vector using EcoRI and PstI.
  2. Perform restriction enzyme digest on DNA insert using XbaI and SpeI.
  3. Calculate the amount of cut vector and insert needed to have a 1:3 molar ratio. The total volume of reaction should have a DNA concentration near 10 ng/µL.
  4. Load the cut vector and insert into PCR tube.
  5. Add enough 10x Ligase Buffer so that the buffer is one-tenth the total volume.
  6. Add 0.5 µL of T4 DNA Ligase into the PCR tube.
  7. Prepare a negative control by replacing the cut DNA insert portion with water.
  8. Place PCR tubes into thermocycler and run the ligation protocol.
  9. Transform product.



›SDS Page

Materials

  • 30% acrylamide mixture (29:1)
  • 50 mL Flask
  • 1.5 M Tris, pH 8.8
  • Nanopure Water
  • Degassing apparatus
  • 10% SDS
  • 10% APS
  • TEMED
  • Gel Casting Module (Bio-Rad)
  • 2-propanol
  • 1.5 M Tris, pH 6.8

Procedure

  1. In a 50 mL filter flask, mix the following components for preparing two resolving gel
  2. Degas the solution for at least 30 minutes then add
  3. While the gel mixture is degassing, assemble the gel-casting module. Examine the glass plates, the inside of the plate (the side that faces the gel) needs to be free of stain, fiber, or any types of visible particles. Pair a spacer plate with a short plate, and fix them in the casting frame. Make sure the bottoms of the two glass plates are leveled.
  4. Examine the rubber seal at the bottom of the casting stand. It should be free of residual gels. Fix the casting frame on the gel casting stand.
  5. Place the gel comb between the glass plate. Make a mark on the place about 0.5 cm below the tip of the comb.
  6. Take the comb out.
  7. Cast the gel by pipetting the resolving gel mixture between the glass plates up the marked line.
  8. Add 2-propanol on the top of the gel, let it stand for 1 hour. When waiting start to degass the stacking gel solution.
  9. In a 50 mL filter flask, mix the following components for preparing two stacking gels.
  10. Degas the solution for at least 30 min, then add
  11. When the polymerization of the resolving gel is finished, rinse off the alcohol with Nanopure water. Remove liquid between the portion of the plates that is above the resolving gel.
  12. Insert the comb between the plates, cast the stacking gel, and let it stand for 1 hour.
  13. Remove the comb, rinse it with Deionized Water to remove gel residues. Then rinse it with nanopure water, dry and store it.



›SLIC

Materials

  • Vector Backbone
  • Insert
  • 10x Buffer 2.1 NEB
  • T4 DNA Polymerase
  • Autoclaved water
  • Thermocycler
  • PCR tubes
  • Pipets

Procedure

  1. 150 ng of vector backbone required. Calculate the volume of vector needed for reaction.
  2. Calculate the amount of insert needed for reaction based on the vector (1:1 molar ratio).
  3. Calculate the amount of autoclaved water needed to bring the final volume to 20 µL and load into a PCR tube.
  4. Load 2 µL of 10x Buffer 2.1 into the PCR tube.
  5. Load the vector and insert into the PCR tube.
  6. Dilute 2 µL of T4 DNA Polymerase into 10 µL of 10x Buffer 2.1.
  7. Load 1 µL of the diluted T4 DNA Polymerase into the PCR tube.
  8. Place PCR tube into the thermocycler and run SLIC Protocol.
  9. Transform product.



›Transformation

Materials

  • Genehog Chemically Competent Cells
  • Water Bath
  • Eppendorf Tubes
  • Chloramphenicol Agar Plates
  • Micropipet
  • Incubator (shaker)
  • Sterilized Beads

Procedure

  1. Thaw 40 µL of competent cells on ice for 5-10 minutes.
  2. Add about 100 ng of DNA then continue icing for 20-30 minutes. The added volume should be less than 10% of total volume.
  3. Place cells into 42°C water-bath for 35 seconds.
  4. Put cells back on ice for 2 minutes. Sterilize work area and pipet in this time.
  5. Add 200µL of LB broth to the mixture.
  6. Leave the cells at 37°C for 45-60 minutes in shaker. Meantime, prewarm agar plates.
  7. Plate cells using beads (volume varies slightly).
  8. Incubate the plate overnight at 37°C.



›Western Blot

Materials

  • Bio-Rad Blotting-Grade Blocker
  • Methanol
  • Nanopure Water
  • Tris
  • 1 x PBST Buffer
  • Bovine Serum Albumin (BSA)
  • Nitrocellulose membrane (Bio-Rad)
  • Western Blotting Filter Paper (Bio-Rad)
  • Ice packs
  • Stir Bar
  • Mini Trans Blot Cell (Bio-Rad)
  • Primary Antibody
  • Secondary Antibody
  • Opti-4CN diluent concentration
  • Opti-4CN Substrate

Procedure




  1. Soak the gel in the Transfer Buffer for 10-15 minutes.
  2. Assemble the transfer sandwich and make sure no air bubbles are trapped in the sandwich. The blot should be on the cathode and the gel on the anode.
  3. Order of sandwich.
    1. Matrix
    2. Filter Paper
    3. Membrane
    4. Gel
    5. Filter Paper
    6. Matrix
  4. To set up the apparatus, place the cassette in the transfer tank. Fill up to the four gel line with Transfer Buffer. Add a stir bar and stir the solution. Add ice packs to keep it cool.
  5. Run for one hour at 100 volts.
  6. Disassemble the apparatus, inspect the membrane and the PAGE. Except the largest size band, all other marker bands should be invisible. Air-dry the membrane for ~ 5 min.
  7. Wash for 5 min in 1 x PBST buffer. (40 cm2 blot, 10 mL/wash; 60 cm2 blot, 15 mL/wash) and put on a rocker.
  8. Block for one hour with Blocking Solution.
  9. Wash for 5 min in 1 x PBST buffer. (40 cm2 blot, 10 mL/wash; 60 cm2 blot, 15 mL/wash) and put on a rocker.
  10. Blot with primary antibody in 6 mL 1 x PBST with 1% BSA for 1 h.
  11. Wash for 5 min in 1 x PBST buffer. (40 cm2 blot, 10 mL/wash; 60 cm2 blot, 15 mL/wash) and put on a rocker.
  12. Blot with secondary antibody in 6 mL 1 x PBST with 1% BSA for 1 h.
  13. Wash for 5 min in 1 x PBST buffer. (40 cm2 blot, 10 mL/wash; 60 cm2 blot, 15 mL/wash) and put on a rocker.
  14. Prepare the detection solution. Make sure you use it as quickly as possible.
  15. Add the solution to blot and incubate for 1 minutes or more to obtain desired result.
  16. Pour off detection solution, wash membrane in nanopure H2O for 15 min.
  17. Document results.





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