Purify, purify and…purify more: tips for improving your protein purification capabilities – part 5

Through a series of posts I have recently shared tips and insights about improving your protein purification preparation. It included bioinformatics analysis for protein purification, affinity chromatography, Ion-exchange chromatography and Size exclusion chromatography. Now that you have successfully established a pure sample, how should you store it? And what can you do to learn about the process and retain that crucial know-how about your protein?

Analysis of your protein
Before commencing with downstream experiments, you should first evaluate the purification level of your protein and its current concentration (and decide if you want to concentrate or dilute it).
The best and cheapest way to evaluate your protein purity is through the use of SDS-PAGE analysis combined with sensitive staining (coomassie staining or better, silver staining). You can also detect minute amounts of contaminants and degradation products can established using Matrix assisted laser desorption/ionization – time of flight analysis (MALDI-TOF). While this method requires only micrograms of protein sample, it bears two downsides: a) it requires buffer exchange into a salt-free buffered solution which can affect the stability of your protein sample and b) that it might not always detect all species within the sample (depends on the ionization capability of each peptide/protein specie).
The easiest and fastest test to assess your protein concentration is through UV absorbance measurement at 280nm. An exception for such method are cases in which your proteins doesn’t absorb at this wavelength, usually due to low number of Tryptophan and/or Tyrosine residues. In such case the use of the Bradford assay (as well as other colorimtetric assays) can be the only way to work around this limitation. It is crucial, however, to either determine which protein is suitable for the calibration curve or to keep using the same protein control so you will consistently calculate your protein sample’s concentration.

Monitor your protein batch collection with Labguru
An important aspect of protein purification and characterization is to maintain your knowledge, know-how and small intricate info about your protein’s behavior. While the process of protein purification has a goal on its own, you can learn a lot about your protein’s behavior to different buffers, pH range and adaptability to changing environments, all of which should be logged and kept safe. Labguru web-based tool for managing your research has a module which is explicitly deals with keeping track of all your samples, and in our case, your protein sample. The module contains all the aspects of your protein purification process such as name, gene ID and specie source, source of expression, purification methods and much more. Description of the exact box in which your samples are located makes this module highly comfortable when you’re ready to test your purified protein. No more wasting precious time and getting a cold burns searching through a deep freezer. The Labguru’s connectivity connects all aspects of your protein purification, whether it was the reserve of the FPLC, purification methodology and any modification you’ve performed. A text free description box is the place where you can describe key aspects of the protein’s behavior (“protein sediments at low salt”) or the specific aim in which this protein should be used for (see screenshot below). An excel sheet can be downloaded from the module to expedite the upload of the information into all the fields in a quick blaze. Try it now for free!

Labguru delivers an excellent way to monitor your protein samples status and storage

How to maintain long term storage of proteins?
Proteins are active moieties which rotate, tumble and vibrate at room temperature constantly and at nanoseconds rate. This rapid motion within the solution can have an adverse effect on protein’s long range structural and functional stability. Thus, once you are satisfied with the protein’s purity, protein concentration and buffer composition, you should place the protein at freezing temperature as soon as possible. The low temperature (the lower the better) will keep your protein at close to stationary state and will extend the proteins longevity within the freezer. The low temperature also inhibits any microbial growth, especially if you didn’t filtered your protein sample prior to freezing it.
The lowest temperature which can be achieved in a standard biochemical lab is close to -200 degree Celsius (liquid nitrogen) though for protein storage a -80 degree Celsius is sufficiently low enough and cost effective. But, wait! Don’t go and stash your protein samples into the deep freezer just like that!
The inherent limitation of freezing an aqueous solution is the transition of water from liquid to solid phase. At this transition temperature (a few degrees above zero Celsius) water molecules interaction shifts from disorder to ordered interaction just before they are transformed into ordered hexagonal crystalline structure, swiftly affecting the hydrophilic-hydrophobic environment in which proteins are held. From the hydrophilic hugging environment, less and less water molecules hydrate the polar residues leading eventually to a decrease of the hydrophobic effect which keeps folded proteins from unfolding an exposing their hydrophobic core.
This is the reason why it is recommended to flash freeze your proteins using liquid nitrogen; in such manner the passage through the denaturation-potential temperature is so short that most protein molecules are not expected to be affected before the temperature is freezing temperature
Since frequent freeze-thawing cycles pass each time through the freezing temperature, it can lead to the denaturation of more and more protein molecules in the sample. To cope with such a problem, you should divide your protein sample into aliquots taking into account the expected down-stream experiments and the amount of protein required. In such a case, a number of thawed vials should be used or thrown to the trash after several days, according to the stability of the specific protein.

How do you treat your proteins? Have you also monitored all your batch purifications?? Leave a comment and share!


10 comments on “Purify, purify and…purify more: tips for improving your protein purification capabilities – part 5

  1. i would like to add a bit of information into long term storage of proteins…proteins can be stored in powder form in deep freezer for long time. the process of removing water content is called lyophilization or freeze drying, initially protein solution is layered into lyophilization jar using Liquid Nitrogen or Methanol and connected to the lyophilizer and vacuum is applied. Some people use excipents to stabilize the protein, once the lyphilization is done, the lyophilized powder can be stored at -80 for long time.

  2. You’re right, Lypholization is another technique for long term storage of purified proteins which I should have mentioned. When working with a large mass of protein preps we found flash freezing using liquid Nitrogen as the best course of action, especially due to the speed and ease of use. Of course, if a certain proteins doesn’t recover well from the freezing process, it might be advised to see if it can recover well from the lypholization process.
    Thanks for your comment!

    • One more thing to mention is lyophilization / Freeze drying helps to reduce sample volume, since the protein sample will be in powder form at the end of lyophilization it is easy to store and also will occupy less space when you have higher volumes of protein sample….

  3. […] document.getElementById('singlemouse').style.display = ''; } Some Process of Protein PurificationPurify, purify and…purify more: tips for improving your protein purification capabilities – part… .recentcomments a{display:inline !important;padding:0 !important;margin:0 […]

  4. Sorry if this is a little off topic, but I read through your series on purification, and I wonder if you could comment on separating complex mixtures, i.e., blood plasma, etc. Recently, I’ve tried this without success, even after following some of the tips you mention. I’m running a Seph S-200 HiPrep 26/60 (320 mL) on a GE Akta purifier. My sample volume was 10 mL of plasma and the system was run at 0.3ml/min. I got 3 peaks early in the run and that was it. Any additional tips that could explain this?

    • Hi Joe,
      Thanks for stopping by.
      As you may well know, there are three major proteins in the plasma: Albumin (65 kDa), various Globulins (90-120 kDa) and Fibrinogens (340 kDa). I guess that the fibrinogen elutes as a major peak at void volume and the other peaks are of the various globulins and albumin.

      From your last statement, I guess you expected to see more proteins. Take into consideration that (a) other proteins might be bigger than the column and elute with the fibrinogen (or clog the filter) or that (b) their concentration is relatively low and you don’t see them due to the low absorbance @ 280nm. You also better increase your flow rate, as 0.3 ml/min is VERY slow and might lead to improper separation – try using 1-2ml/min.
      Good luck!

      • Hi Chen,

        Thanks for the tips–that makes sense. One thing, though–I thought, perhaps incorrectly, that a slower flow rate would lead to better resolution. Is this not true? Also, do you think I would get better resolution if I diluted the sample beforehand, e.g. 2-fold? GE recommended this, but then again, they also told me before to load as much protein as possible (pure plasma)… Thanks again!

      • Hi Joe,
        After making some calculation I am taking back my statement about your working flow rate – it IS at the optimal range (which is 2-5 cm/hr). If you go below 2 cm/hr you risk diffusion of the peak (refer to Van Deemter equation).
        In regard to diluting your sample – it really depends on the peaks height and width and since I can’t see the chromatogram its hard to comment on it. Generally speaking, injecting less material will inevitable lead to better resolution of peaks, but it will increase your work time by two fold on the one hand, and on the other you will dilute less prominent proteins which will make you identify and collecting them more difficult. You really need to see whether your TARGET protein is “overloaded”. Just look at the peak and observe if it is rounded at the top or flat. If it is flat than you’ve crossed the maximum reading of the inline spectrophotometer, which also means a LOT of protein on a 26/60 column.

        One more point: when separating a sample in which there is a highly abundant protein and a low abundant protein, there is a chance that you will miss the low abundant protein altogether due to the wide scaling of the absorbance, especially on a large volume column such as the 26/60 series. If you’re suspecting that additional proteins are present in the sample (and you haven’t seen them in the previous runs), I suggest you use a Y axis scale of 200-500 mAU, even go as low as sub 100. This way you will notice most proteins which has some abundance in the sample.

        And more importantly, if you’re looking to purify a low abundant protein in such a sample, you might want to consider salt/pH precipitation of the sample such you remove most of the abundant contaminants. Size exclusion chromatography excels when size differences are large and significant and makes a poor first step for highly contaminated samples.
        Hope this helps!

      • Thanks so much for the great information, Chen! I’m actually interested in proteins across the molecular weight range–my intention is to separate the proteins out roughly by size and use the fractions for further experiments. I suspect there are proteins in the fractions where the absorbance is extremely low, as you suggest. Changing the scale does allow me to see more tiny peaks. Perhaps pooling a few runs after concentration should give me enough material for the lower abundance fractions.

      • Yes, this can be a good strategy. Take into consideration that low abundant protein will be hard to observe under coomassie staining so you might want to concentrate the samples (irreversibly) through Trichloroacetic acid (TCA) – here’s a protocol if you wish to explore.
        Good luck!

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