Team:IIT Kanpur/Protocols


1. Take 750 mL of milliq water.
2. Weigh and mix 10 grams of NaCl.
3. Weigh and add 10 grams of Tryptone.
4. Weigh 5gm of yeast extract and add to the solution.
5. Make the volume up to 1 Litre using dH20.
6. Put cap on and wrap foil without tightening the cap.
7. Autoclave for 60 minutes.

1. Thaw competent cells in ice (0 Degrees) for 10 minutes.
2. Add 2-5 µl of DNA into competent cells and then flick. (Do NOT vortex)
3. Incubate the competent cells in ice for 30 minutes.
4. Heat shock at 42°C for 45-60 seconds.
5. Incubate in ice for 5 minutes for cell recovery.
6. Add 1 ml of LB media to the tube.
7. Place tube at 37°C for one hour.
8. Plate the cells onto the plates of corresponding resistance.
9. Incubate the plates for 12-16 hours at 37°C.

Materials required: T4 DNA Ligase(NEB), Ligase Buffer, Insert and Vector backbone

1. Fill x volume of water in an 1.5ml eppendorf for 10µl reaction
2. Now add the reagents strictly in the following order.
3. Add 40ng of Insert and 60ng of vector.
4. Add 1uL of ligase buffer.
5. Add 1µl of T4 DNA buffer.
6. Incubate at 16°C for 2 hours or overnight.


  • Thaw ligase buffer on ice as it contains ATP which deactivates on exposure to heat.
  • We optimised our molar ratio for insert to vector using NEB calculator specific to our construct

Materials required: DNA template, Restriction Enzyme, Buffer, dH20 Add DNA template(200-500ng) followed by dH2O (x µl) and RE (10units) followed by Buffer(10x) for 20µl reaction. Incubate at 37°C for 4 hours or overnight.

1. To make 1% Agarose Gel add 1 gm of agarose powder in 100 mL of 1xTAE.
2. Boil the mixture in microwave.
3. Then let the agarose solution cool down.
4. Add 2-3 μl ethidium bromide (EtBr). Mix to dissolve EtBr uniformly.
5. Pour the agarose into a gel tray with the well comb in place, avoiding bubbles.
6. Allow the gel to solidify.
7. Place the gel into the electrophoresis unit, which is filled with 1xTAE.
8. Samples are loaded into the wells of the gel.
9. The gel is run in accordance with the size of the sample to get appropriate resolution.


  • Remember to load the ladder for comparison while making inference.

Materials Required: Transfer buffer, Nitrocellulose membrane, Fast Semi-Dry Machine

Preparing Buffers and reagents: lysis buffers

  • 20 mM Tris-HCl
  • Protease inhibitors
These buffers may be stored at 4°C for several weeks or aliquoted and stored at -20°C for up to a year.
Preparing solutions and reagents: running, transfer and blocking buffers
  • 5X loading dye
  • 5% β-Mercaptoethanol
  • 0.02% Bromophenol blue
  • 10% SDS (Sodium dodecyl sulfate)
  • 250mM Tris-Cl (250 mM, pH 6.8)
pH must be adjusted to 6.8 ​
Running buffer (Tris-Glycine/SDS)
  • 25 mM Tris base
  • 190 mM glycine
  • 0.1% SDS

Check the pH and adjust to 8.3
Transfer buffer (wet)
  • 25 mM Tris base
  • 190 mM glycine
  • 20% methanol
  • Check the pH and adjust to 8.3

For proteins larger than 80 kDa, we recommend that SDS is included at a final concentration of 0.1%.
Transfer buffer (semi-dry)
  • 48 mM Tris
  • 39 mM glycine
  • 20% methanol
  • 0.04% SDS
Blocking buffer
Take 5% skimmed milk to TBST buffer. Vortex it.
Note: Make sure you filter the blocking otherwise it can lead to spotting, where tiny dark grains will contaminate the blot during color development.
Sample preparation from supernatant
1. Remove a small volume of supernatant to perform a protein quantification assay.Take absorbance at 280nm.
2. Add 5X loading dye one-fifth of the volume of protein sample. ​
3. Boil each protein sample at 100°C for 5 min to reduce and denature samples.

Note: Protein samples can be aliquoted and stored at -20°C for future use.Loading and running the gel
1. Load equal amounts of protein, around 20-30μg of total protein into the wells of the SDS-PAGE gel, along with molecular weight marker.
2. Run the gel for 2 h at 100 V.

The gel percentage required is dependent on the size of your protein of interest:
Protein size &nbsp &nbsp &nbsp &nbspGel percentage
4–40 kDa &nbsp&nbsp&nbsp&nbsp&nbsp 20%
12–45 kDa &nbsp&nbsp&nbsp&nbsp&nbsp 15%
10–70 kDa &nbsp&nbsp&nbsp&nbsp&nbsp 12.5%
15–100 kDa &nbsp&nbsp&nbsp&nbsp&nbsp 10%
25–100 kDa &nbsp&nbsp&nbsp&nbsp&nbsp 8%
Transferring the protein from the gel to the nitrocellulose membrane
Activate the membrane with methanol for 1 min and rinse with transfer buffer before preparing the stack. Apply 10V for 1 hr. Transfer of proteins was checked by Ponceau S staining before the blocking step.
Prepare the stack as follows:
Antibody staining
1. Block the membrane for 1 h with skimmed milk(blocking buffer) at room temperature or overnight at 4°C.
2. *Incubate the membrane with appropriate dilutions of primary antibody in blocking buffer.
3. Wash the membrane in three washes of TBST, 5 min, 10 min and 5 min respectively.
4. Incubate the membrane with the recommended dilution of conjugated secondary antibody in blocking buffer at room temperature for 1 h.
5. Wash the membrane in three washes of TBST, 5 min each.
6. *For signal development, follow the kit manufacturer’s recommendations. Remove excess reagent and cover the membrane in transparent plastic wrap.
7.Acquire image using darkroom development techniques for chemiluminescence, or normal image scanning methods for colorimetric detection.


· 0.1% w/w Ponceau S dye – 0.5 g
· 1% v/v acetic acid – 5 mL
· Make it up to 500 mL with m-Q water

10x TBS

Tris (mw 121.14), final conc 0.5M
NaCl (mw 58.44), final conc 1.5M
· Disolve 60.55g Tris and 87.66g NaCl in 800ml m-Q water.
· Adjust pH to 7.5 with HCl then make up to final volume of 1L.


· 100 ml of 10x TBS
· 1ml Tween 20
· 899 ml m-Q water


All steps on the roller at room temperature:
1. Stain the membrane with 5 mL of the Ponceau solution for 5 min.
2. Destain the background with m-Q water – i.e. until you see the band & image.

Ø Rinse with m-Q water a few times to destain further.
Ø Two (or more) 10 min washes with 20 mL TBS-T for the complete removal of the dye.
Ø Proceed with blocking the membrane as normal.

· Store solutions at room temperature.
· Staining solution can be re-used between 10-40 times.
· Do not use with nylon-based filter media.