HIS tagged protein purification protocol – piss easy guide
Day 1:
1. Thaw sample, 37oC water bath x 10 mins
2. Set up column (in stand, remove original cap and add yellow cap), equibrilate with
Ni-NTA agarose (shaken) from radioactive fridge. Add using plastic pipette to the
line and leave to settle.
3. Pour sample into falcon tube. Wash sonification probe with ethanol then water.
Jam tube into plastic beaker full of ice. Full power (100 amp) x 20s. Off for 20s
minimum. Do x6 such bursts.
4. Back on ice; pour into centrifuge tube, 18,000 rpm JA-20 (about 35,000g) x 20 min.
Pour supernatant (full of protein) into new tube and repeat centrifuge.
5. During this, make up the lysis buffer (200 ml per sample). Remove yellow cap
from column and allow liquid to drip through agarose (can suck off top). Then gently
add lysis buffer on top of the agarose, then all the way to the top. Allow 15 mins
(25mins max, 22 perfect) (don’t leave it to dry out!) for buffer to reach line. Repeat
wash once more and leave till at the line.
6. Filter sterilise (0.45um) sample into fresh falcon tube. Keep on ice.
7. Once buffer has dripped through to line gently add sample with plastic pipette, then
pour. Insert yellow funnel and pour whole sample in. Leave to drip through till the
line (HIS tagged protein binds to column) – about 1h 15 mins for 25ml sample.
8. Wash with lysis buffer up to top of column tube – 30 mins. Repeat – 30 mins.
Final wash (large) with remaining lysis buffer, will need yellow funnel – 1 hour. Get
rid of excess after this time.
9. Make up elution buffer – 50ml per sample (only need about 10ml each) - (lysis
buffer + extra imidazole) anytime after sample is added to column. ADJUST to pH 8
10. Elute – add to top of column and COLLECT in 3 x 1.5ml labelled (1,2 & 3)
eppendorfs. Keep on ice.
11. Check to see if protein is present – dilute Bradford stock solution (5X, dark
brown, bottom of radioactive fridge). Add 2.5ml to 4 small falcon tubes. Add 50ul
from each eppendorf (the last of which probably contains no protein) and a negative
control of elution buffer. If protein present it turns blue.
12. Now must remove the imidazole (it fluoresces) by dialysis. Check that there are
enough dialysis clips (!). Make up 2L of dialysis buffer (lysis buffer minus
imidazole). Gloves on, cut dialysis (about 10 cm) into ¾ full 2L plastic beaker
dialysis buffer. Clamp one end (if doing multiple samples, label!) and do trial fill
with dialysis buffer using plastic pipette, then pour out. Now add the protein into
same membrane (prob only from first 2 eppendorfs) and clamp other end too. Put on
stirrer in cold room at low speed, and add remaing buffer, clingfilm over top and
leave o/n.
Day 2:
13. Next day, replace with fresh buffer and leave for another 24h. Can add multiple
samples to same 2L but do 2 solution replacements (another in the afternoon).
Day 3:
14. Do Bradford. Aliquot 50ul into small eppendorfs. Protein storage at -20oC.