How to Design the Perfect Protein Purification Buffer

When purifying a protein, it’s important to keep your protein happy. If you are going to use the protein in binding and activity assays, such as the surface plasmon resonance (SPR) technique, then your protein needs to be soluble and active. For success in these experiments, it is crucial that you create a buffer that prevents unfolding and aggregation. To design the buffer, you should consider the following factors:

  • pH
  • buffering system
  • salt
  • reducing agents
  • stabilizing elements

As described below, each of these needs to be optimized for your protein of interest.


Many experiments are done at pH 7.4 to mimic biological conditions. If your protein is stable at this pH – great! If not, you need to change the pH to find conditions that keep your protein in solution. One rule of thumb is that proteins are generally less soluble at their pI value, which is the pH at which the protein has no net charge. A quick and easy way to calculate a protein’s pI from its sequence is to use ExPASy’s ProtParam tool.

Buffering System

Once you’ve decided on a pH value, you need to decide which buffer you are going to use. The most important thing to keep in mind when choosing a buffer is to make sure that your buffer of choice has buffering capabilities at your pH of choice. Choose a buffer that has a pKa value within one pH unit of your desired pH.

The second most important thing is to ensure that the concentration of buffer you are using is high enough to buffer the solution. Concentrations between 50-100 mM are common.

Keep in mind that the buffer you use should not interfere with the activity of your protein—this is especially important for SPR techniques. For example, phosphate inhibits kinases and should be thoroughly dialyzed out before performing reactions. Also, some buffers are sensitive to temperature. Tris is notorious for this. For example, if you pH your buffer to pH 8.0 at 25°C, the pH will increase to 8.58 at 5°C and decrease to 7.71 at 37°C. So, if you plan to store your protein at 4°C or do your experiment at 37°C, take into consideration that the pH you measured at room temperature may be different under your experimental conditions.


Many buffers contain NaCl to help keep proteins soluble and to mimic physiological conditions. Generally, 150 mM NaCl is used. However, during various protein purification steps, you may want to change the salt concentration. For example, if you are purifying your protein by ion exchange chromatography, you want to start with a low concentration of salt (5-25 mM). This will help screen ionic interactions and prevent nonspecific binding of proteins to the column while enabling your protein of interest to bind the column. In other types of chromatographic separations, like gel filtration and Ni2+ affinity columns, you may want to increase the salt concentration. I’ve gone up to 500 mM NaCl to prevent nonspecific interactions between proteins and the column.

Expert tip: Change the salt concentration by dialyzing your protein into a new buffer.

Reducing Agents

If your protein contains cysteine residues, oxidation could become a problem and cause protein aggregation. To prevent this, keep a reducing agent such as DTT, TCEP, or 2-mercaptoethanol in your buffer. In general, TCEP is the most stable of the three, but it can be rather expensive. I often use DTT in my buffer during purification and then add TCEP to the final buffer. A good concentration to use for these reducing agents is between 5-10 mM. Basically, you want to make sure that the concentration of the reducing agent is well above your protein concentration.

Because DTT and BME break down at room temperature, keep these buffers in the refrigerator. Alternatively, make the buffers without reducing agent and add the reducing agent when you’re ready to use the buffer.

Make sure any resins you use are compatible with reducing agents. For example, high concentrations of reducing agents reduce the nickel in nickel columns and turn the column brown. The column can be regenerated, but your protein is not likely to bind well. Many columns have suggested maximum concentrations of reducing agents that the column can tolerate; however, I’ve found that this is really trial and error.

Stabilizing Elements

Finally, there are a whole slew of additives you can add to your buffer to help increase protein solubility and stability. You can try adding an inert protein like BSA to your buffer. This can sometimes stabilize a protein, but you must ensure that the protein you’re adding does not interfere with your experiment. Sometimes it helps to increase the viscosity by adding agents like glycerol or PEG. These typically help prevent aggregation. Also, some detergents and other ionic compounds like sulfates, amino acids, and citrate can be used in small quantities to help shield ionic interactions and solubilize proteins.

So there you have it. By keeping these five things in mind: pH, buffering system, salt, reducing agents, and stabilizing agents, you are well on your way to creating a buffer that will keep your protein happy and active for use in various analytical techniques (including SPR applications).


  1. lianjin729 on May 15, 2018 at 2:15 pm

    I really like your introduction for protein purification. I have some question for you since it’s my first time to purity the polypeptide comprising thiols by using prep-HPLC. Can I use TCEP in basic buffer to purify protein? because my protein only soluble in basic buffer, which pH is near to 8.5 is there any suggestion to remover TCEP after purification?

  2. Syed Yusuf Mian on July 2, 2017 at 7:06 pm

    Is there a way to stop protein degradation during expression in E.coli? I am working on a number of proteins and almost all degrade. Can anyone suggest how to avoid this degradation? Another problem is that one of my protein is not even expressing in E. coli. Any suggestion on this, please?

  3. Jennifer Aniston on October 24, 2016 at 9:58 pm

    Say you had you protein buffered to pH 8, and you accidentally added water; what would this do to the protein and how could it be remedied?

    • Matthew on June 2, 2017 at 1:55 am

      Hi Jennifer,

      The nature of a buffer is that it maintains a given pH value even when confronted with small or moderate perturbations to the system. Say you have 1 L of your buffer at pH 8. If you pour extra water (a volume small compared to the starting volume of the buffer) into the buffer, the pH will remain essentially at 8. By diluting the buffer with the extra water, you would expect the hydrogen ion concentration to go down and thus the pH to go up. This, in fact, does happen, but the equilibrium between the acid and base forms of the buffering chemical then shifts in response. In the case of dilution, the equilibrium shift will cause more weak acid to dissociate, which restores the hydrogen ion concentration essentially to its starting point, meaning that the pH will still be 8 after everything has come to equilibrium. It is important to understand that buffers can only maintain the pH in the face of *small* or *modest* changes to the system. If you added 10 L of water to your original 1 L of typical-strength buffer, there would likely be an insufficient amount of weak acid in the system to compensate for the huge dilution, and the system would be unable to restore the hydrogen ion concentration to its original level. Thus, the final pH would go up.

  4. iqra qaddir on April 24, 2016 at 7:22 am

    in anion exchange cromatography cation resin used which is DEAE . my protein pH is 6.5. can i used tris HCl buffer with pH of 8

    • D on June 14, 2016 at 6:55 pm

      If the pH is above the pI, then the protein will have a negative charge. So in this case, yes, you can have a pH 8 buffer to perform anion exchange chromatography.

  5. Rodrigo on March 28, 2012 at 9:50 am

    When choosing pH do you generally go under or above the pI? For example if you have something with a theoretical pI of 5, does using a buffer of pH 4 give you some sort of an edge during purification? Also a lot of people tried to convince me that high salt conditions (eg. 500mM) help to solubilize proteins. That’s against my perception of hydrophobic interactions, which are promoted at high ionic strength. What’s your opinion.

    • Jennifer Cable on April 5, 2012 at 1:07 am

      Thanks for all the great comments everyone!

      Rodrigo: I wish there was a good answer for your questions, but I don’t think there is. I think the likely answer is going to be “it depends”. I can say that the protein that I worked with in grad school had a pI of about 7.0 and was most stable (thermal stability) at pH 5.0 (although we didn’t try to go lower). Also, I’ve never been able to figure out why proteins are not stable at their pI. I mean, they still have charges, they just all happen to cancel out. If anyone has a good explanation for this, I’d love to hear it.

      As for the salt, again, it depends. Back to my protein from grad school, I went up to 2 M NaCl, and that protein just kept getting more stable (again, thermally) the more salt I added. I think your reasoning about salt strengthening the hydrophobic effect is right on, but the hydrophobic effect is also the driving force for protein folding. So, I would think that adding salt might be advantageous if your protein has a lot of surface charge, especially if there are patches of highly negative or positive areas that may interact nonspecifically, but if your protein has some highly hydrophobic regions on the surface, increasing the salt may promote aggregation and “salt out” the protein. (I don’t have any evidence of this though, just kind of rationalizing it in my head).

      That’s just my two cents. Does anyone have any ideas?

      • Gabriel on December 3, 2015 at 5:55 pm

        Don’t quote me on this, but I believe that at a proteins normal pH those charges are part of what keeps the hydrophobic core folded away from the hydrophilic surroundings; when you remove those net charges by bringing a protein to it’s pI, the whole protein now acts hydrophobic (since there’s no charge), and there’s no advantage to those amino acids sticking to the outside

        • Martin on March 22, 2016 at 7:53 pm

          I don’t think PI correlates to no charge in the protein’s surface, but it means the protein as a whole doesn’t have net charge and won’t migrate in an electric field. There can be local charges, as the case of a zwitterionic at a pH that keeps both basic and acidic species ionized but the molecule neutral as a whole.

  6. MarkB on November 9, 2011 at 9:55 pm

    pretty good in concept, difficult in execution

  7. max-ch on November 7, 2011 at 2:57 pm

    Good introduction for buffer preparation. I also systematically supplement purification buffers with protease inhibitors like leupeptin or pepstatin A.
    Also metal chelators like EDTA is to consider depending on the protein.

  8. Jeff Hollins on November 4, 2011 at 3:24 pm

    I work with an incredibly unstable protein and have quite a lot of experience with enhancing protein stability. One of the best methods I’ve found so far is to use the thermofluor technique to determine which conditions best stabilise your protein of interest this technique uses a q-PCR machine and sypro orange to determine protein melting, using this technique you can run 96 different conditions in 15 minutes.

    Aside from thermofluor there are a few additional pointers I would like to add to the (very well written) above article: Potassium chloride is a much better salt than sodium chloride (most cells actively transport potassium into the cell and sodium out, and cytosolic concentrations tend to be between 100 – 300 mM); Glutathione and cysteine can also be used as reductants; TMAO is probably the best protein additive at concentrations greater than 1 M, in addition both me and a colleague studying completely different proteins have recently discovered that glutamate can considerably increase protein stability at concentrations of greater than or equal to 1 M, in addition glutamate is exceptionally cheap.

    For more information search for osmolytes (the terminology used for stabilising elements in journals) and the hofmeister series (a general series of stabilising and destabilising salts).

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