Dialysis tubing is a semi-permeable membrane that lets small molecules pass through while keeping larger ones trapped inside. To use it, you soak the tubing to hydrate it, clamp one end shut, fill it with your sample solution, clamp the other end, and submerge it in a buffer or water bath. The process is straightforward, but small mistakes in preparation or setup can ruin your results.
How the Tubing Actually Works
The tubing acts like a wall full of tiny pores. When you place a filled tube into a surrounding liquid, molecules smaller than those pores move freely back and forth until concentrations equalize on both sides. This is diffusion: particles naturally travel from where they’re concentrated to where they’re not. Larger molecules, like proteins, can’t fit through the pores and stay trapped inside the tubing.
Water itself also moves across the membrane in response to osmotic pressure. If the solution inside the tubing is much more concentrated than the liquid outside, water will flow in and cause the tubing to swell. If the outside solution is more concentrated, the tubing can shrink or collapse. This water movement is why headspace and clamping technique matter so much.
Choosing the Right MWCO
Dialysis tubing comes rated by its molecular weight cut-off (MWCO), which tells you the approximate size of the largest molecule that can pass through the pores. Common ratings range from 1,000 to 50,000 daltons. The general rule is to pick a MWCO that’s at least two to three times smaller than the molecule you want to keep inside the tubing. If you’re retaining a 30,000-dalton protein, for example, tubing with a 10,000-dalton MWCO gives you a comfortable margin. Using a cut-off too close to your target molecule’s size risks losing some of it through the membrane.
Conversely, if you’re trying to remove small contaminants like salts or buffer components (typically under 500 daltons), a higher MWCO lets those small molecules escape faster while still retaining your protein of interest.
Preparing the Tubing
Most dialysis tubing ships dry and coated with glycerol (a humectant that keeps it flexible during storage) and trace sulfur compounds from manufacturing. You need to remove these before use, or they can contaminate your sample.
Start by washing the tubing in running water for 3 to 4 hours to remove glycerol. For more thorough cleaning, treat the tubing with a 0.3% sodium sulfide solution at 80°C for one minute to strip sulfur-based preservatives, then wash it in hot water at 60°C for two minutes. Follow that with a brief rinse in dilute sulfuric acid (0.2%), and finish with another hot water rinse to clear the acid.
If your experiment doesn’t require that level of purity, a simpler approach works: soak the tubing in distilled water for 15 to 30 minutes until it becomes soft and pliable. Dry tubing is stiff and brittle, so never try to fill or clamp it before hydration. Once hydrated, handle the tubing with gloves to avoid introducing oils or contaminants from your skin.
Filling and Sealing the Tubing
Cut a piece of tubing longer than you think you need. A reliable formula for calculating length: divide your sample volume by the tubing’s volume-per-length (listed on the packaging), then add 10 to 20% for headspace, plus an extra 4 centimeters for the clamps on each end. That headspace is critical. If osmotic pressure draws water into the tubing during your experiment, the bag needs room to expand. Without it, the tubing can burst.
Fold over one end of the hydrated tubing and secure it with a dialysis clamp or a tight knot. Some people double-clamp for extra security. Before adding your sample, fill the sealed tubing with water and check for leaks by gently squeezing it over a sink. Any pinholes or a bad seal will be obvious.
Drain the test water, then use a pipette or syringe to fill the tubing with your sample solution. Pour slowly to minimize air bubbles trapped inside. A few small bubbles won’t derail your experiment, but large pockets of air reduce the effective membrane surface area and slow diffusion. Once filled, squeeze out excess air from the headspace gently (without forcing out your sample), fold the open end, and clamp it shut.
Setting Up the Dialysis
Place the sealed tubing into a container of your dialysis buffer or distilled water, depending on your experiment’s goal. The volume of the outside solution matters: use at least 200 to 500 times the volume of your sample for efficient exchange. A small beaker with barely enough liquid to cover the tubing will slow diffusion dramatically because the concentration gradient disappears quickly.
Stirring the outside solution with a magnetic stir bar speeds up the process significantly. Without stirring, a layer of already-equilibrated fluid builds up around the tubing surface and acts as a barrier. Gentle, continuous stirring keeps fresh buffer in contact with the membrane.
For most applications, you’ll want to change the outside solution at least two or three times during dialysis. A common schedule is to dialyze for 2 to 4 hours, replace the buffer, dialyze again overnight (8 to 12 hours), then replace the buffer one more time for a final 2-to-4-hour round. Each buffer change resets the concentration gradient and drives more of the unwanted small molecules out of your sample.
Temperature depends on your sample. Room temperature works for chemical experiments and classroom demonstrations. Protein samples and other biological materials typically need to be dialyzed at 4°C (in a cold room or refrigerator) to prevent degradation.
Common Problems and How to Avoid Them
Leaking is the most obvious failure. It usually comes from a bad clamp, a pinhole in the tubing, or tubing that was damaged during handling. Always do a water leak test before adding your real sample. If you’re reusing tubing, inspect it carefully since dried and re-wetted tubing is more prone to weak spots.
Osmotic bursting happens when you underestimate how much water will flow into the bag. Solutions with high salt or sugar concentrations inside the tubing will draw substantial water in from a low-concentration outside bath. Leave adequate headspace, and check the tubing periodically during the first hour to make sure it isn’t ballooning.
Loss of your target molecule is another concern. If your MWCO is too close to the size of the molecule you’re trying to retain, some of it will leak out over time. This is especially true for molecules that are near the cut-off threshold, since MWCO ratings aren’t perfectly sharp boundaries. Some molecules can also adsorb directly onto the cellulose membrane rather than staying in solution, which reduces your yield without any visible sign. Pre-rinsing the tubing with your buffer before adding the sample can help reduce this adsorption.
Contamination from the tubing itself is a subtler issue. Residual glycerol, sulfur compounds, or heavy metals from manufacturing can leach into your sample if you skip the preparation steps. For sensitive downstream applications, thorough cleaning is not optional.
Storing Prepared Tubing
If you’ve hydrated more tubing than you need, store it submerged in distilled water at 4°C. For longer storage (weeks or more), add an antimicrobial agent like 0.05% sodium azide to the water to prevent bacteria and fungi from breaking down the cellulose membrane. Rinse the tubing thoroughly with fresh water before using stored tubing, especially if sodium azide was in the storage solution. Never let hydrated tubing dry out completely, as it becomes brittle and unreliable once re-dried.