Sauer:"Quantitative" purification of His-tagged proteins from cell lysates (analytical scale)
This procedure works well for the small-scale purification of His6-tagged proteins from cell lysates. It can be carried out in a manner that allows for quantitation of purified proteins. It works well for proteins that are not easily detected with commercially available anti-His6 antibodies (e.g. proteins with internal His6 tags). The protocol is relatively easy to carry out and is highly amenable to analysis of samples from time course-type experiments. This protocol should also work well for purification of radiolabeled proteins in pulse-chase-type experiments.
- Take 1 mL of E. coli culture and spin for 1-2 min at full speed in a microfuge to pellet the cells. Remove the supernatent, being careful not to disturb the pellet. Resuspend in 40 µL UL buffer by vortexing and freeze at -80 ˚C until ready to purify.
- Thaw pellets at room temperature and add 200 µL of additional denaturing buffer (UL buffer if you want to run the input or flowthrough of your column on a gel, GuHCl buffer if you do not).
- Add a known amount (50-500 pmol) of a purified His6-tagged protein to each sample. Make sure that this protein will not run near your protein of interest on a gel. This will allow you to quantitatively compare your samples at the end of the experiment.
- Allow cells to lyse at room temperature for ~20 min, vortexing 2 or 3 times.
- Spin for 10 min at full speed in a microfuge to pellet the cell debris. Transfer the supernatent to a new tube.
- Wash enough 50% Ni-NTA (Qiagen) slurry with UL or GuHCl buffer such that you have 60 µL of 50% slurry per sample. To do this, spin the resin at low speed (~4000 rpm in most microfuges or in a "personal" minifuge/nanofuge that many people seem to have on their bench) and remove the supernatent. Add UL or GuHCl buffer and repeat the procedure 2-3 times. After the final wash, ensure that your total volume is the same as what you started with such that your resin is still at 50%.
- Add 60 µL of washed 50% Ni-NTA slurry to each sample.
- Allow samples to bind to the resin for 30-60 min at room temperature, shaking or rotating to facilitate binding.
- Spin the samples at low speed to collect resin and liquid at the bottom of the tube.
- Remove supernatent.
- Resuspend resin in 200 µL GuHCl or UL buffer.
- Transfer the resin and liquid to a small spin column. (I like Costar Spin-X tubes from Corning: #8170, 0.45 µm nylon filter in polypropylene tubes). Wash the tube (and your pipet tip) with an additional 200 µL of UL or GuHCl buffer and transfer to the column. This step helps to ensure that you are not losing too much sample.
- Spin out the liquid at low speed. Save the flowthrough, if desired.
- As an alternative procedure for flowthrough and washing of the column (if you do not want to collect these samples), a Qiagen vacuum manifold can be used. For each sample, attach a 5 mL luer-lock syringe to the vacuum manifold and set the spin filter in the top of the syringe. Apply a vacuum to pass liquid through the column. Transfer the spin filters back to microfuge tubes for elution.
- Wash the column with 500 µL GuHCl or UL buffer.
- Wash the column three times with 500 µL wash buffer 1.
- Wash the column two times with 500 µL wash buffer 2.
- Spin out any remaining liquid at low speed.
- Elute the sample with 100 µL elution buffer. Let the buffer incubate with the column for 10 min. Spin at full speed in a microcentrifuge to collect all of the eluate. Repeat with another 100 µL elution buffer and combine eluates.
- Speedvac the samples to dryness and resuspend in 30 µL UL buffer (use less volume if necessary to visualize your samples).
- Run 10-20 µL of each sample on an SDS-page gel.
- Stain the gel to visualize the proteins.
- If you have a lot of protein, coomassie staining may work well for this.
- For lower amounts of protein, silver staining, which is much more sensitive, may be necessary.
- Alternatively, I have had success using Sypro-ruby stain from Molecular Probes/Invitrogen. It is more sensitive than coomassie staining, and, in my hands, appears to be more sensitive than silver staining, as they claim.
- Quantitate your samples.
- The relative amount of each sample can be determined by comparing your band to that of the protein you spiked in at the beginning of the experiment (this should be the same in all lanes, so you can adjust for differential recovery).
- The absolute amount of protein can be determined by comparing your samples to purified samples of known concentration.
- All buffers can be stored at room temperature.
urea lysis buffer
10 mM Tris-HCl, pH 8
100 mM NaCl
6 M guanidine-HCl
100 mM NaH2PO4
10 mM Tris base
Adjust to pH 8
Wash buffer 1
100 mM NaH2PO4
150 mM NaCl
20 mM imidazole
Adjust to pH 8
Wash buffer 2
50 % Acetonitrile
50 % acetonitrile
0.1% trifluoroacetic acid (TFA)
- This procedure has been used successfully to isolate proteins that are synthesized from medium copy (10-20 per cell) plasmid templates in E. coli. It has worked well for proteins that are expressed at a low level (cannot discern the expressed protein among the other E. coli proteins on a coomassie-stained gel) and for those expressed at a higher level (obvious band on a coomassie-stained gel among cellular proteins). You may want to modify the protocol if you protein is expressed at a very low level. This is probably most easily done by increasing the volume of culture you use for each sample.
- This protocol has been used successfully for E. coli cultures with OD600 of 0.2-3 (according to my spectrometer). For densities above this range, the efficiency of lysis in the given volumes may become an issue, and you may want to consider increasing the lysis volume.
- Spinning the Ni-NTA resin too hard in a microfuge will crush the resin, compromising its integrity.
- For quantitative measurements you need to ensure that all of your protein is bound to the column. The volume of resin used in the procedure above is well above that necessary to bind all of the His6-tagged protein in most samples. However, you should do a "test run", monitoring input, flowthrough, and elution samples to ensure that all of your protein is being bound by the column, and adjust the column volume if necessary. Additionally, when doing the actual experiment, run a lane on the gel that contains the amount of tracer protein (the one you spiked in at the beginning) that you would to expect to see on the gel if you recovered all of the sample.