Skip to content

5 Ingredients for the Perfect Protein Purification Buffer

Posted in: Protein Expression and Analysis
A woman surrounded by spheres to represent a purified molecule in the perfect protein purification buffer.

Listen to one of our scientific editorial team members read this article.
Click here to access more audio articles or subscribe.

Purified proteins must be kept soluble and active (happy) for your experiments. pH, the buffer system, salt concentration, reducing agents, and additives can all be adjusted to design the perfect protein purification buffer. Beware of buffer species interfering with purification steps and downstream experiments. Keep your buffer as simple as possible while keeping your protein active.

Unfortunately, purified proteins must be dissolved in a buffer solution when we experiment on them, and the buffer should keep them stable and active. This is the source of many headaches for protein scientists, as proteins tend to aggregate once removed from their native cells.

In this article, we explain the factors you need to consider to design the perfect protein purification buffer so that you can purify exquisite samples for your experiments.

Factors to Consider Before You Prepare Your Buffer

When purifying a protein, it’s important to keep it stable and active (happy, as we like to say). This is essential for most intended experiments, such as activity assays, surface plasmon resonance (SPR), and structural studies using cryo-EM or X-ray crystallography.

For success in these experiments, you must create a buffer that prevents protein unfolding and aggregation.

Creating the optimal buffer is more complex than it might first seem since the buffer needs to resemble the physiological environment of the protein (to keep it active) while being chemically simple because more ingredients increase the likelihood of one of them interfering with your experiment.

To design the perfect protein purification buffer, we must consider the following five factors:

  1. pH;
  2. buffering system;
  3. salt;
  4. reducing agents;
  5. stabilizing elements/additives.

Let’s look at each of these in closer detail.

How to Design the Perfect Protein Purification Buffer

1. pH

Many experiments are done at pH 7.4 to mimic biological conditions. If your protein is stable at this pH—great! If not, you must change the pH to find conditions that keep your protein in solution.

You should also consider whether your protein needs to carry a positive, negative, or neutral charge for your experiments.

To make your protein positively charged, make the pH of the buffer lower than the protein’s isoelectric point (more acidic).

To make your protein negatively charged, make the pH of the buffer higher than the protein’s isoelectric point (more basic).

To keep your protein neutral, make the pH of the buffer match the protein’s isoelectric point.

Remember, the isoelectric point is the pH at which a molecule has no net charge. Read our handy guide for a refresher on isoelectric points, pH, and pKa, A quick and easy way to calculate your protein’s isoelectric point is to paste its primary sequence into ExPASy’s ProtParam tool.

2. Buffering System

Once you’ve decided on a pH value, it’s time to decide which buffer you will use. The most important thing to keep in mind when choosing a buffer is to make sure that it has buffering capabilities at your pH of choice.

Choose a buffer with a pKa value within one pH unit of your desired pH. Fortunately, we have an article that explains how buffers work and tells you their pKa.

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.

Remember 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-adjust 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, consider that the pH you measured at room temperature may be different under your experimental conditions.

3. Salt

Breaking news! Water is actually quite a poor solvent.

Most proteins are not soluble in pure water and must be salted in. This is the process of increasing the ionic strength of a solution to increase the solubility of desired solutes—your protein—in it.

Therefore, a protein purification buffer usually contains NaCl to help keep proteins soluble and mimic physiological conditions.

Generally, you include NaCl at 150 mM. 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 salt concentration (5–25 mM). This will help screen ionic interactions and prevent the binding of unwanted proteins to the column while enabling your protein of interest to bind to 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 increased up to 500 mM NaCl to prevent nonspecific interactions between proteins and the column.

For less work in the lab, you can put your protein purification steps in order of decreasing salt concentration so that your final sample is your protein dissolved in a buffer with the fewest number of additional ingredients.

Expert tip: You can change the salt concentration by dialyzing your protein into a new buffer!

4. 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.

The typical concentration to use for these reducing agents is between 5–10 mM. You want to make sure that the concentration of the reducing agent is well above your protein concentration… and if you’ve got 10 mM protein, you’re doing very well!

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

Also, 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.

Also, consider the length of your experiment and how long you need to keep your protein reduced. If your experiment lasts several days, but your protein oxidizes in a few hours, use TCEP.

5. Stabilizing Elements and Additives

Finally, there is 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.

It’s a sort of never-ending list, but you can read about the purpose of common additives here.

Purification Buffers in Summary

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 perfect protein purification buffer that will keep your protein happy and active for any experiment you want.

Take your protein purification game to the next level with our two free eBooks: The Bitesize Bio Guide to Protein Expression and Five Methods for Assessing Protein Purity and Quality.

Originally published November 2011. Revised and updated July 2023.

Share this to your network:


  1. 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.

  2. 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.

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

    pretty good in concept, difficult in execution

  4. 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.

  5. 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).

Leave a Comment

You must be logged in to post a comment.

This site uses Akismet to reduce spam. Learn how your comment data is processed.

Scroll To Top