Dialysis Methods for Protein Research
Dialysis is a separation technique that facilitates the removal of small, unwanted compounds from macromolecules in solution by selective and passive diffusion through a semi-permeable membrane. A sample and a buffer solution (called the dialysate, usually 200 to 500 times the volume of the sample) are placed on opposite sides of the membrane. Sample molecules that are larger than the membrane-pores are retained on the sample side of the membrane, but small molecules and buffer salts pass freely through the membrane, reducing the concentration of those molecules in the sample. Changing the dialysate buffer removes the small molecules that are no longer in the sample and allows more contaminants to diffuse into the dialysate. In this way, the concentration of small contaminants within the sample can be decreased to acceptable or negligible levels.
How dialysis membranes work. A dialysis membrane is a semi-permeable film (usually a sheet of regenerated cellulose) containing various sized pores. Molecules larger than the pores cannot pass through the membrane but small molecules can do so freely. In this manner, dialysis may be used to perform purification or buffer exchange for samples containing macromolecules.
Protein Preparation Handbook
This 32-page handbook provides useful information on our broad portfolio of reagents and tools for protein extraction, clean-up, immunoprecipitation and purification. Practical information, selection guides, and relevant data are included to help you improve your protein yield and downstream analysis.
- Specific topics covered included the following:
- Cell lysis and fractionation
- Protein dialysis and other purification techniques
- Immunoprecipitation and pull-down assays
- Other methods for protein preparation
Principles of diffusion
Dialysis works by diffusion, a process that results from the thermal, random movement of molecules in solution and leads to the net movement from areas of higher to lower concentration (until an equilibrium is reached). In dialysis, unwanted molecules inside a sample-chamber diffuse through a semi-permeable membrane into a second chamber of liquid or dialysate. Because large molecules cannot pass through the pores of the membrane, they will remain in the sample chamber. By contrast, the small molecules will freely diffuse across the membrane and obtain equilibrium across the entire solution volume, effectively reducing the concentration of those small molecules within the sample.
If dialysis is allowed to proceed to equilibrium before each change of dialysate buffer, the substances retained by the membrane are purified by a factor equal to the ratio of buffer volume to sample volume. For example, when dialyzing 1 mL of sample against 200 mL of dialysis buffer, the concentration of unwanted dialyzable substances will be decreased 200-fold when equilibrium is attained. Following two additional buffer changes of 200 mL each, the contaminant level in the sample will be reduced by a factor of 8 x 10^6 (200 x 200 x 200). If the original sample contained 100 mM DTT, this would potentially be decreased in the sample to approximately 12.5 nM following three complete cycles. If a further decrease in concentration is desired, the dialysis process can be continued with additional volumes of dialysate.
Use of dialysis cassette for protein cleanup. 3 mL of 1 mg/mL IgG in 0.1 M Tris buffer, pH 7 inside a dialysis cassette is placed in 1,000 mL of 100 mM PBS, with a pH of 7.6. The old dialysate is discarded and replaced with 1,000 mL of 100 mM PBS, with a pH of 7.6. IgG is too large to enter the pores in the membrane; therefore, the amount of IgG inside the cassette remains constant. The Tris buffer concentration drops below 0.01 M inside the cassette as the Tris buffer diffuses out and the PBS buffer diffuses in. Again, the old dialysate is discarded and replaced with 1,000 mL of 100 mM PBS, with a pH of 7.6. The IgG inside of the cassette remains constant. The Tris buffer inside of the cassette drops to near undetectable levels. The buffer inside the cassette is 100 mM PBS, with a pH of 7.6.
The time required to accomplish dialysis is determined by factors that affect the rate of diffusion of a molecule. Because heat affects the thermodynamics of molecules, increasing temperature speeds diffusion. Therefore, dialysis will proceed faster at room temperature than at 4°C. In selecting the most appropriate temperature, it is important to take into account the thermal stability of the molecule of interest. The rate of diffusion is also directly proportional to the concentration of a molecule, while inversely proportional to its molecular weight. As the concentration of a molecule increases, so does the probability that one of those molecules will contact the dialysis membrane and then diffuse across to the other side. However, as a molecule's molecular weight increases, the rate of movement in solution decreases along with the chance of diffusion through the membrane - even if it the molecule is small enough to pass through the pores.
The rate of dialysis is also directly proportional to the surface area of the membrane and inversely proportional to its thickness. Membranes normally used for laboratory dialysis applications are 0.5 to 1.2 mil (12 to 30µm) thick, providing good diffusion rate as well as structural integrity. While membrane thickness is not a variable that is easily modified, the surface area usually is. The flatter a sample can be spread over a membrane surface, the faster will be its dialysis because all molecules in the sample will be closer to the membrane and a higher proportion of them will be in direct contact with the membrane at any instant. High-performance dialysis products, such as Thermo Scientific Slide-A-Lyzer Dialysis Cassettes, MINI Devices and Flasks, are designed to maximize surface area-to-volume ratios (within practical limits) for different volumes of sample.
Influence of surface area to volume ratio on dialysis rate. Graph displays rates of removal of 1 M NaCl from 2 mL, 70 mL, 70 mL, and 200 mL samples dialyzed in four respective sizes of Thermo Scientific dialysis devices, each equipped with 3.5K MWCO membrane. Dialysis was conducted at room temperature against very large volumes (e.g., 4 L) of water (dialysate). At the indicated times (triangles), the dialysis buffer was changed and the percentage of NaCl removal was determined by measuring the conductivity of the sample. The large device (Flask) has about half the surface-area-to-volume ratio of the other devices, accounting for the slower rate.
Stirring the buffer during the dialysis process also increases the diffusion rate. As low molecular weight compounds exit through the pores on the outer side of the membrane, they form a microenvironment termed a Nernst diffusion layer. In this layer, which is approximately 200-300 molecules thick, the low molecular weight compounds are at a higher concentration in relation to the rest of the dialysate. This high local concentration effectively slows the rate of dialysis because molecules can randomly re-enter the dialysis membrane pores and return to the sample. Stirring efficiently breaks up the macroenvironment outside the Nernst layer, helping to maintain the concentration differential needed to drive the diffusion process.
Molecular weight cut-off (MWCO) describes membrane pore size measured in angstrom (Å) units. A larger MWCO corresponds to a wider pore size. The MWCO describes the smallest average molecular mass of a molecule that fails to diffuse across the dialysis membrane. For example, a membrane with a 10K MWCO will retain more than 90% of proteins with a molecular mass of 10 kDa or greater. The membranes most commonly used for laboratory dialysis are made of regenerated cellulose, manufactured using either the cuprammonium or viscose process. For both of these methods, dissolved cellulose is extruded as tubing or sheets and then dried. Glycerol is frequently added as a humectant to prevent cracking during drying and to help maintain desired pore structure. Pores range from 15-50 Å for 3.5K, 7K, 10K and 20K MWCO membranes. The membranes have a symmetrical pore structure that allows small molecules to migrate across them in either direction. Regenerated cellulose is hydrophilic and easily saturated in buffer to provide a homogeneous medium for dialysis in aqueous solutes. Membrane diffusion capacity is directly related to hydrophilicity. Thermo Fisher Scientific offers a variety of dialysis devices ranging in size from 2K-20K MWCO.
Protein sample retention. Both 2K and 20K MWCO Thermo Scientific Slide-A-Lyzer cassette membranes were used to dialyze proteins or vitamin B12 (1 mg/mL) in either saline or 0.2 M carbonate bicarbonate buffer, pH 9.4 overnight (17 h) at 4°C. The amount of retentate was estimated using either the Pierce BCA Protein Assay Kit or by absorption at 360 nm (for vitamin B12). Slide-A-Lyzers cassette membranes are available in these pore sizes: 2K, 3.5K, 7K, 10K and 20K.
The manufacturing process for regenerated cellulose membranes results in the presence of certain low-level "contaminants" in the final product; for certain experiments, the possible effects of these components on samples should be considered. The main contaminants are sulfur compounds (0.01-0.3%), heavy metals (trace) and glycerol (0-21%). Most of these small compounds diffuse out of the membrane during the dialysis process. Therefore, if it is suspected that these substances might interfere with the function of the molecule of interest, pre-dialyze the membrane or device with ultrapure water or buffer for 30 minutes before adding the sample.
Regenerated cellulose exhibits minimal protein adsorption. While some protein adsorption may occur with any sample, more concentrated samples will result in a lower percentage of total protein being lost. Regenerated cellulose membranes also have better chemical compatibility and heat stability than membranes made from cellulose acetate (cellulose ester). They are more resistant to organic solvents and weak or dilute acids, and they are compatible with the pH range and buffer salts that are commonly used in protein and molecular biology applications.
All cellulose membranes are sensitive to cellulase enzyme activity. Standard concentrations of antimicrobial inhibitors (e.g., 0.05% sodium azide) can be added to prevent growth of cellulolytic microorganisms if the membranes are stored wet for long periods of time before use. Membranes should not be allowed to dry out after wetting unless they are reglycerinated because drying can alter (decrease) the pore size. Avoid closing the ends of dialysis tubing by tying (knotting) because the tension can increase the pore size and MWCO near the tie. Avoid touching membrane with bare hands to prevent possible enzymatic and microbial contamination. Pre-assembled devices, such as the Slide-A-Lyzer dialysis cassettes, MINI devices and flasks, eliminate the need to manipulate the membrane during setup and sample handling.
Watch this video to learn more about protein dialysis
It is the difference in the composition of sample and dialysis buffer solutions that creates the concentration-differential across the membrane that drives the dialysis process. Using a high buffer-to-sample volume-ratio helps to maintain the concentration gradient. The number of dialysate buffer changes and the dialysis time also affect the outcome achieved in dialysis. Because of the variables associated with each sample, a universal dialysis procedure for all applications cannot be provided, only general guidelines. Similarly, the process completion-point is somewhat subjective. The goal is to reduce the concentration of low molecular weight compounds to a level that will not interfere with subsequent steps in the experiment.
A typical dialysis procedure for protein samples is as follows:
- Pre-wet or prepare the membrane according to instructions.
- Load sample into dialysis tubing or device.
- Dialyze for 1 to 2 h at room temperature.
- Change the dialysis buffer and dialyze for another 1 to 2 h.
- Change the dialysis buffer and dialyze overnight at 4°C.
Note: For best results, use a volume of dialysis buffer (dialysate) that is at least 200-fold greater than the sample volume. To conserve dialysate with large-volume devices, such as the Slide-A-Lyzer dialysis flasks, a 5-fold excess of dialysate is sufficient, especially if it is changed several times.
Small volume protein sample dialysis: The 0.1 mL Thermo Scientific Slide-A-Lyzer dialysis devices are designed to hold 10 to 100 µL samples. Dialysis is performed against a beaker of solution as shown, or by placing an individual device in a microfuge tube filled with distilled water.
Many samples will take on water or buffer during the dialysis process due to osmotic pressure. This occurs frequently with samples that have a high starting salt concentration or if a component of the sample is hygroscopic. In the case of high starting salt concentration, osmosis causes water to enter the sample faster than buffer salts within the sample are able to diffuse out, resulting in the swelling of the sample within the dialysis sample compartment. When this occurs, it may be desirable to return the sample to its original concentration, or to decrease the sample volume even further.
To concentrate the sample, dialysis membrane containing the sample is placed in a small plastic bag containing a solution of hygroscopic compound instead of ordinary dialysate. To avoid contamination of the sample, the hygroscopic compound must be composed of molecules that are larger than the pore size of the dialysis tubing (e.g., high-molecular weight polyethylene glycol). With this set-up, concentration occurs upon diffusion of the water (osmosis) and other small molecules out of the sample and into the hygroscopic solution.
Another method to concentrate samples is through forced dialysis. Vacuum is applied to a sample contained within a dialysis membrane; this effectively "pulls" water, buffer salts and other low-MW compounds out of the dialysis sample-chamber. Another form of diafiltration involves "pushing" samples through a dialysis membrane by centrifugal force; this is the basis for protein concentrators, which have become popular in recent years.
Watch this video to learn more about protein concentrators:
Walker JM. 2009. The Protein Protocols Handbook. Third Edition. Springer-Verlag New York, LLC
For Research Use Only. Not for use in diagnostic procedures.