Solutions and equipment required
- Dialysis tubing
- Sucrose or PEG 20 kDa
- Clean Tupperware
Use a dialysis membrane with a 10 kDa molecular mass cutoff. Prepare the membrane by boiling in 100 mM EDTA for 10 mins. Rinse thoroughly with distilled water. Seal one end with clips or string and pour the protein solution into the tube. Seal the other end.
In a clean tupperware box, pour sucrose or polyethylene glycol 20 000 [PEG 20 kDa]. If you use sucrose, you will get an approximate 3-fold reduction in volume within an hour. However, the dissolved sucrose outside the membrane will diffuse into the protein solution, thereby contaminating with sucrose. Usually this is not a problem as sucrose is inert.
If you are using PEG 20 kDa, you must keep an eye on the solution at all times. Concentration may be completed in 30 min, depending on the starting volume of protein solution and the amount of PEG 20 kDa that you have poured around the membrane. Due to its size, PEG 20 kDa will not diffuse into the protein solution. Once the concentration is finished, gently wash the outside of the tubing with distilled water, cut open one end of the tubing and remove the concentrated solution of protein by aspiration with a micropipette.
Solutions and equipment required
- 100 mM NaOH (prepared in distilled water)
- 20% ethanol
- ultrafiltration membrane with 10 kDa cutoff
- nitrogen cylinder
- Get an ultrafiltration membrane from Blommie.
- Never touch the filter without gloves and try to avoid any manipulation that might scratch or tear the membrane.
- You will notice that the membrane is anisotropic. Always keep the shiny side up, i.e. this side is the one that will be in contact with the protein solution in the ultrafiltration device.
- Clean the ultrafiltration device with detergent and wipe down with 70% ethanol. Allow to air dry.
- Place membrane, shiny side up, at the bottom of the device. Fit rubber O-ring above the membrane and assemble the device completely.
- Add ~20 ml buffer into the device and screw on the top air-bleed valve.
- Place the ultrafiltration device on a stirrer and switch on the power. The ultrafiltration paddle should stir smoothly, and gently. Never stir protein solutions too vigorously. Always ensure that the ultrafiltration paddle is stirring or else there will be a build up of protein on the membrane surface and this will clog buffer flow and halt concentration.
- Place a clean beaker at the end of the waste tube.
- Attach the device to a nitrogen cylinder and admit nitrogen to a final pressure below the tolerances of the device.
- Look at the buffer eluting from the waste tube. If there are tiny, rapid bubbles coming out, this means that there is a tear in the membrane and you should replace it. In the beginning, as you pressurise the device, you will notice a few bubbles, but they will pass out of the tube quickly and the remainder of the buffer within the tube should be free of bubbles.
- Check the solution periodically – the rate of filtration is dependant on the protein concentration.
- If you want to remove the protein solution from the device or add more solution for concentration, first close off the nitrogen valve. Next, twist the upper portion of the air-bleed valve to equilibrate the pressure within the device. Twist off the main body of the bleed valve. You can remove the protein solution using a sterile plastic pipette.
- Usually, one would want to concentrate the protein solution to the smallest volume [assuming that it does not aggregate at high concentrations]. The ultrafiltration device can concentrate down to 500 μl.
- After concentration, disassemble the device and place the membrane in 100 mM NaOH for 30 min to remove residual/denatured protein and to sterilise the membrane. Wash the membrane well with distilled water to neutralise the pH. Store the membrane in 20% ethanol in the fridge until required. The membranes can be reused many times.
Ammonium sulfate precipitation (salting out)
Ammonium sulfate is a highly soluble kosmotrope that is useful for fractionally precipitating proteins. Purified proteins can be stored as ammonium sulfate precipitates in the fridge for extended periods, which can be advantageous if the protein is known to be sensitive to freezing. An added advantage of storing proteins as ammonium sulfate precipitates is that the high salt concentration is an effective bacteriostatic. Precipitation with ammonium sulfate may be performed by either solid salt or by dilution from a saturated solution [usually, 4.1M in distilled water]. Reproducible results only occur if you keep the temperature constant, i.e. if you perform salting out at room temperature and determined at which % saturation your protein of interest came out of solution, do not assume that it will do the same if you perform subsequent salting out on ice.
A downside of ammonium sulfate precipitation is that one usually has to get rid of the salt afterwards as it may interfere with ion exchange or enzyme assays or other chemical reactions [such as crystallisation]. Residual ammonium sulfate in resuspended pellets is not usually a concern nowadays before ion exchange as one can just ensure that the pellet is diluted in excess buffer to reduce the [ammonium sulfate] to negligible concentrations. Also, the strong ion exchangers [e.g. Q and SP] bind proteins far tighter than the conventional DEAE/CM columns and they therefore do not show reduced binding if there is a bit of salt left over from the precipitation.
- Crush ammonium sulfate with a mortar and pestle.
- Use the formula: G = (533 (S2 – S1)/ (100-0.3 S1)
S1 = % saturation in starting solution
S2 = % saturation in final solution
G = grams of solid ammonium sulfate to be added per litre.
- If you have not determined the ammonium sulfate solubility for your protein, it is useful to do a series of “cuts”. This involves increasing the %saturation of ammonium sulfate in 10% fractions.
- To do a “cut”, you first need to measure the volume of the starting protein solution and determine how much ammonium sulfate is required to go from a starting saturation of 0% to a final saturation of 10%. So, looking at the formula above, if I have 40 ml of protein solution and I wanted to increase ammonium sulfate saturation from 0-10%, I would need to add (533[10-0])/(100-0.3×0) = 2.132 g. Note that some ammonium sulfate tables in catalogues and on the web differ with the way they present the amounts. Some list the weight to be added per 100 ml while others are per litre. Also, some tables list amounts required if precipitating at room temp and others on ice. The formula above assumes you are doing it on ice.
- After you have weighed out the correct amount of ammonium sulfate, add it all in one quick motion and stir rapidly but not too quickly, to prevent high local concentrations of ammonium sulfate. Once the salt has dissolved, leave on ice for 20 min to allow proteins to come out of solution. Centrifuge at 10 000 x g, 10 min, 4°C. Pour off the supernatant [keep aside] and resuspend the pellet in a minimal volume of salt-free buffer.
- Measure the supernatant that you have set aside and then calculate how much ammonium sulfate is required for you to increase the %saturation from 10 to 20 [as the supernatant still contains 10% ammonium sulfate and this is now the starting %]. Repeat as above, resuspending pellets in buffer and adding ammonium sulfate in 10% saturation cuts until protein no longer comes out of solution and/or ammonium sulfate saturation has been reached [i.e., no more salt will dissolve].
- Measure the protein concentrations in all the pellets.
- Dilute all the pellets to the same protein concentration and assay enzyme activity on each.
- Determine specific activity and assess protein profile by SDS-PAGE. Ideally, you would choose the %saturation that precipitates the majority of your protein. As this is the very first step in the protein purification, you cannot expect your enzyme-containing fraction to be free from contaminants. It is a very crude separation technique but can increase specific activity tremendously.
- A few proteins do not tolerate ammonium sulfate and, if this is the case, try other precipitation techniques such as heat treatment or solvent treatment to get rid of contaminating protein.
Ion exchange, followed by desalting chromatography
Ion exchange is a very fast method for concentrating proteins, although it will usually have to be followed up with a dialysis step or passage over a desalting column. If done in conjunction with a desalting column, protein concentration by this method can easily be completed in 30 min.
Before performing this method, you would have determined the salt concentration required to elute your protein from the ion exchanger you are going to be using. So, if you know that your protein is eluted with 300 mM salt, then use this concentration for the procedure below.
Solutions and materials required
- A 2 ml ion exchange column [e.g. Q-Sepharose] made up in a 5ml syringe on a layer of glass wool.
- Buffer with the appropriate concentration of salt to elute from the column.
- Sephadex G-25 column, pre-equilibrated with salt-free or reduced salt buffer.
- 0.22 μm filters for sterilisation.
- Peristaltic pump [make sure that the tubing has been cleaned with NaOH and 70% ethanol, and finally rinsed in distilled water]
- You will probably perform this step after ion-exchange on the big Q-Sepharose column as active fraction volumes tend to be quite large [>50 ml].
- Pool your active fractions and dilute with an equal volume of buffer containing zero salt.
- Open a sterile 5 ml syringe and remove the plunger.
- Fix a short piece of tubing on the end of the nozzle and clamp.
- Add a small amount of glass wool to the bottom of the syringe to acts as a gauze to prevent loss of the ion exchange matrix through the nozzle.
- Add the Q-Sepharose slurry and allow to settle. Once the volume has reached 2 ml, open the nozzle to allow excess buffer to drain. Wash the column with 10 ml salt-free buffer and clamp the end.
- Add protein solution into the column by using a peristaltic pump and release the clamp. Adjust the flow rate of the peristaltic pump so that it is slightly greater than the flow rate from the column. Once all the sample has been loaded onto the column, wash the column with 2 column volumes of salt-free buffer.
- Thereafter, wash the matrix with 2 column volumes of buffer with salt and collect the eluate into a clean tube.
- The eluate will contain your protein of interest and will probably need to be desalted before you move to the next step.
- I have attached a guide for Sephadex-25 desalting to this set of instructions. There should be powdered G-25 in the EMU or MCB. You can find the instructions for swelling and preparation on the web.
- Desalting is very useful in that it results in minimal dilution of the protein and it is rapid. The protein will elute in the void volume while the salt will elute at approximately total volume (Vt).
- Once you have collected the eluted protein, you can filter-sterilise and store until use in whichever state you have determined to be least denaturing.
These are useful in concentrating protein if your initial sample volume is less than 2 ml. The EMU has supplies of spin-filters. They are fairly expensive, but can be reused – check the conditions on the insert in the box. The downside to the spin columns is that concentration takes very long – usually around 2-3 hours centrifugation for a 2ml sample. However, the advantage is that the sample will not run dry due to the design of the tubes and that one could concentrate to less than 100 ¼l. The filters are resistant to most solvents and chaotropes, however, check the insert before you spin, if you have anything exotic in your protein solution.
Trichloroacetic acid/deoxycholate (TCA/DOC)
Only use this method if you need to concentrate proteins for SDS-PAGE. It is very good for precipitating low-concentration proteins. Be careful with TCA – it is highly caustic. Dispose TCA as for any chlorinated waste.
- 100% TCA solution. Weigh out 25 g TCA and add distilled water to a final volume of 25 ml.
- 2% [m/v] Na-deoxycholate (DOC) in distilled water.
- Laemmli sample treatment buffer.
- To one volume protein, add 1/100 volume of 2% DOC. Keep on ice for 15 min.
- Add 1/10 volume 100% TCA and keep on ice for 1 hour.
- Centrifuge at 10 000 x g, 10 min, 4°C.
- Wash the pellet in acetone to get rid of residual TCA.
- Resolubilise the pellet in appropriate buffer.
- When you add Laemmli sample treatment buffer to your protein solution, the bromophenol dye should remain blue. If it changes to yellow, it means that there is TCA still remaining and that your proteins will probably migrate cathodically rather than anodally. If the solution turns yellow, add a few microlitres of 1 M Tris, pH 8.8 until the colour changes back to blue.
- Load your samples and electrophorese.