Laboratory 8B Dialysis

Transcription

1 Laboratory 8B Dialysis Concepts: Dialysis is used to effect buffer exchange. Dialysis is not a purification method. Dialysis takes advantage of the properties of a semipermeable membrane. Protein solubility is affected by ph and salt concentration and must often be determined empirically. Goals: You will be choosing an appropriate dialysis buffer, based on your understanding of your POI and dialysis and taking into account what was used previously for your POI, and then dialyzing your protein purified by affinity chromatography into this buffer. Following dialysis, your protein is ready for use in functional assays. I. Introduction In biochemistry, we use dialysis to separate molecules in solution based on the difference in their rates of diffusion through a semipermeable membrane. Only molecules that are small enough to fit through the pores in the membrane are able to equilibrate with the entire volume of the dialysis (sample and dialysis buffer). At equilibrium, molecules small enough to move through the pores do so in both directions equally. At this point, to further reduce the concentration of small molecules requires moving the sample to a fresh dialysis buffer. A new gradient is formed between the sample and the fresh dialysis buffer. Again, particles concentrated in the dialysis membrane that are small enough to move through the pores equilibrate by moving through the pores to the lower concentration buffer. Each change of dialysis buffer brings about a further dilution of small molecules in the sample. Molecules too big to pass through the pores in the membrane will not reach equilibrium with the dialysis buffer but will stay inside the dialysis tubing. Dialysis allows for the removal of salts and other small molecules from your protein sample. In simple terms, dialysis can be thought of as a method of buffer exchange. Dialysis makes use of semi-permeable membranes. In the simplest example, this membrane is manufactured in the form of tubing (looking much like a sausage casing). The main feature of this membrane is that it is porous. However, the pore size is such that while small salt ions can freely pass through the membrane, larger protein molecules cannot (i.e. they are retained). Thus, dialysis membranes are characterized by the molecular mass of the smallest typical globular protein which it will retain. The molecular weight cutoff (MWCO) of our snakeskin dialysis tubing is 3.5 kda (3,500). This is commonly referred to as the cutoff of the tubing (e.g., a 3,500 Dalton protein will be retained by the tubing but smaller molecular mass solutes will pass through the tubing). Dialysis proceeds by placing the high salt sample in dialysis tubing (i.e., the dialysis "bag") and putting it into the desired lower salt buffer.

2 More on the principle of dialysis: Over time the concentration of low molecular mass solutes within the bag, and in the low salt buffer, will come to equilibrium. In practical terms, salt molecules will diffuse out of the bag into the lower salt buffer. At equilibrium the salt concentration of the sample can be calculated. The buffer volume for the dialysis is a function of the required final concentration of salt in the sample and can be calculated. Choosing the right volume for Dialysis: a hypothetical example You have a 10 ml protein sample from an ion exchange column elution pool which contains 1.0 M NaCl. For your next step in the purification you can have no more than 1 mm NaCl in the sample. Thus, if you dialyzed 10 ml of sample (at 1.0 M NaCl) in 10 L of no salt buffer, after equilibrium the NaCl concentration in the sample would be 1.0 mm. Note that in the above example this would commonly be referred to as a "1:1,000" dialysis. Suppose that we don't want to make up 10 L of buffer. We can actually achieve the same results with two sequential "1:32" dialyses (i.e. the square root of the 1:1,000 dialysis; in other words, two sequential 1:32 dialyses is equivalent to a single 1:1,000 dialysis): After the first dialysis (against 310 ml of buffer) the sample NaCl concentration will be mm. This is determined by using the formula and solving for Concentration 2. (Vol 1 )(Concentration 1 ) = (Vol 2 )(Concentration 2 ) (10 ml) (1 M) = (320 ml) (C 2 ) Note that the Vol 2 is 320 ml, not 310 ml, because you must add the sample volume in. The NaCl will be equilibrated over the full volume in your dialysis beaker (= sample + dialysis buffer). After the second dialysis (against another 310 ml of buffer) the sample NaCl concentration will be mm (solving for C 2 in the equation below). (10 ml) (31.25 mm) = (320 ml) (C 2 ) Thus, instead of making 10 L of buffer, we could make 620 ml and achieve the same results with two dialysis steps. In this case, removing the salt would take twice as long, i.e. we need to perform two dialysis steps. Many factors can affect the rate and completeness of dialysis, including dialysis buffer volume, buffer composition, the number of buffer changes, time, temperature, and particle size vs. pore size (see Technical note from Pierce Scientific available on collab). As a general rule, smaller molecules will achieve equilibrium faster than larger substances. In practice it can be difficult to predict or determine the time required for dialysis. General guidelines given by the manufacturer of our snakeskin tubing suggest that thorough buffer exchange typically involves a dialysis volume of times that of the sample, dialysis at room temperature for a total of 6-8 hours, and 3 changes of dialysis buffer. The final dialysis can be left overnight at 4 o C.

3 One consequence of dialysis to watch out for is that while salt ions are moving out of the bag, water molecules are moving into the bag. Thus the volume of sample may actually increase (the bag will swell) and, therefore, the protein concentration will decrease In the extreme case, the bag may actually swell to the point of rupture. Therefore, it is a good idea not to fill the bag completely, but leave a void to allow for potential swelling. How to choose the appropriate buffer for dialysis: Protein solubility is a complex function of the physiochemical nature of the proteins: ph, temperature, and the concentration of the salt used. The specific composition of the protein (acidbase groups) determines its solubility in different buffers. The surface of a protein has a net charge that depends on the number and identities of the charged amino acids, and on ph. At a specific ph the positive and negative charges will have a net charge of zero. This ph is called the isoelectric point, and for most proteins it occurs in the ph range of 5.5 to 8. A protein has its lowest solubility at its isoelectric point. If there is a charge at the protein surface, the protein interacts with water, rather than with other protein molecules. This charge increases the protein solubility. Without a net charge, protein-protein interactions and precipitation are more likely. As a general rule, your protein should be in a buffer at least one full ph unit away from the isoelectric point. At low concentrations of salt, solubility of the proteins usually increases slightly (salting in). But at high concentrations of salt, the solubility of the proteins drop sharply (salting out). Initial salting in at low concentrations is explained by the Debye-Huckel theory: proteins are surrounded by the salt counter ions (ions of opposite net charge) and the charge screening results in decreasing the electrostatic free energy of the protein and increasing activity of the solvent, which in turn, leads to increasing solubility. The behavior of proteins in solutions at high salt concentrations was explained by Kirkwood: the abundance of the salt ions decreases the solvating power of the salt ions, the solubility of the proteins decreases and precipitation results. As a general rule, you will want to maintain some salt in your dialysis solution. Physiological conditions are generally mm salt and neutral ph. Despite taking care in choosing your dialysis buffer, you may find that your protein precipitates. While we can be guided by the information above, the appropriate buffer must often be determined empirically. Your survey of the literature may help you to determine conditions which were favorable for similar proteins. Some cofactors are absolutely essential for protein folding. You should determine from literature if your POI needs any small molecules, ions, or metals for stability and solubility. The information from literature and the pi of you protein may be the most useful in determining the buffer you choose for dialysis. II. Required reading The above introduction and lecture material Dialysis overview on collab The final manuscript from the CHEM4411/4421 group that previously studied your POI

4 III. Pre-lab Assignment ed to your TA by Sunday at 8 PM (Be sure to do pre-lab 8A on page 139 as well.) Determine what buffer you will dialyze your protein into. Provide justification for that buffer (e.g., cite a reference). If you choose a different buffer than used previously from the students, justify the change. Determine and report how you will make the buffer you have chosen (what amount of each chemical you will need for the volume of buffer you are making, etc.) Explain what each component of your buffer does/why it is necessary (i.e., ph, salt, cofactors). To be shown to your TA at 2PM on the day of lab None IV. Materials: Large beakers Aluminum foil Length of snakeskin dialysis tubing Protein sample Dialysis buffer (volume and concentrations to be determined by you) Dialysis clips Serological pipet and bulb or p1000 and tips Magnetic stir bar Stir plate V. Procedure: Part I. Test your dialysis buffer Before you commit to dialyzing your entire sample in the buffer you have chosen, it is sometimes wise to test your protein for solubility in that buffer on a small scale. To do this, take an aliquot of your protein in elution buffer (~200 L) and dilute it with ~ 800 L of the proposed dialysis buffer (for highly concentrated proteins (see your gel from Laboratory 9A and consult your TA), dilute ~100 L with 900 L). Let sit at room temperature after thorough but gentle mixing and check for precipitation. If no precipitation, proceed with dialysis. If you see precipitation, re-evaluate your buffer choice. 1. Prepare a dialysis buffer ( L). 2. Collect ~2 ml of this buffer to set aside for use in Lab 10. Part II. Dialysis 1. Fill a large labeled beaker with an appropriate volume of the appropriate dialysis buffer ( ml). The dialysis buffer you will use is dependent on the type of assay you will be performing. You will determine the volume of the buffer and how many changes you will do. Place a stir bar in the bottom and cover the top with a piece of aluminum foil. 2. Cut a suitable length of dialysis tubing. The length you will need per volume is listed on the tubing label (i.e., 3.7 ml sample / 1 cm of length). To this length, add 6-7 additional

5 cm to have room for the clips and air. For this class you should cut at least 10 cm of tubing. Handle the tubing very gently, and only with gloves. Do not let the tubing touch the bench or other surfaces. Do not poke or damage the tubing. Use a clean razor blade to cut the tubing. If necessary, one person should hold the tube with the tubing and the other cut. 3. Dip/soak your length of tubing into your dialysis buffer to wet it completely. 4. Fold the bottom edge of tubing (approximately 1-2 cm) over and gently clip. Squeeze buffer out of the tube gently using a gloved hand. 5. Turn aluminum foil (used to cover buffer beaker) buffer side up and work over this surface. This serves as a safety net. Any spilled protein will be caught by the foil and you will be able to recover it. It may work best to have one person hold the tubing and the second person pipet. Be extremely careful to avoid poking tubing. 6. Using a serological pipet, carefully pipet your protein solution (the elution fraction(s) from Laboratory 9A) into the tubing. Do not puncture the tubing. You may elect to use your P1000 repeatedly if you are more comfortable. Neither is more or less correct. 7. Fold top of tubing over and clip; leave 2-4 cm of air in tubing above your solution so that it will float. 8. Place tubing in buffer, stir for at least 2 hours. Keep the stirring to a low speed so that the dialysis bag is not pulled down by the stirring. 9. Replace buffer and let stir at least 2 hours. If necessary because of time, it may stir overnight. Arrange to return in the morning to change the dialysis solution again and/or to remove your protein from dialysis. Part III. Removing sample after dialyzing: Again, working over foil, carefully pour your protein solution from the tubing into a labeled conical tube. 1. Gently dry the exterior of the tubing with a kimwipe (just to remove the excess buffer; the tubing will remain wet). 2. Unclip one end of the tubing. Straighten the tubing and invert a clean, empty conical tube over the tubing. 3. Flip the tubing and tube together and squeeze the entire volume into the tube. Alternatively, you can put your conical tube into a rack to pour the solution in. Second alternative is to pipet the solution out of the dialysis tubing. This is best done with a partner 4. Place the labeled protein tube at 4 o C. Do not freeze your protein sample. You will observe this protein sample over the next couple of weeks so see how it behaves in the buffer you chose. You will also use this protein next week when you learn how to determine protein concentration.

6 VI. Data Analysis Needed for Lab Report What buffer was used? Why was this buffer chosen? Why was it necessary to perform dialysis? What overall dilution of the elution buffer was obtained?