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Protein Dialysis

Dialysis is the separation of small and large molecules in a solution by selective diffusion through a semi-permeable membrane. It is generally used for larger sample volumes, and can take hours to overnight for complete dialysis.

Please view our selection table to choose the right dialysis device for your experiment.

The first generation cassettes can require a syringe to insert the sample, and may require a float buoy. They can hold 0.5–30 mL solution. The second-generation cassettes can be loaded with a syringe or pipet, are self-floating, and can hold 0.5–70 mL.

Yes, we offer gamma-irradiated array Slide-A-Lyzer™ cassettes for both first-generation products (Cat. No. 66454, 66455, 66453, 66456) and second-generation products (Cat. No. 88250, 882551, 882552, 882553, 882554).

We do not test for or claim sterility.

The kit includes float buoys and syringes, and are only available for first-generation products. Please note that floats and syringes can also be purchased separately.

Please see the protocol summary below:

  • Insert syringe needle through the gasket via one of the corner ports. Inject the sample, withdraw the excess air and remove the syringe.
  • Attach a float buoy and dialyze. (Buoys also serve as convenient bench-top stands for the cassettes).
  • Insert empty syringe needle at a second corner port. Inject air to expand the cassette chamber, then withdraw the dialyzed sample.

Cassettes have easy handling and secure sample delivery, while traditional tubing is slippery when wet and can be difficult to handle, increasing risk of sample loss during sample addition/removal. Cassettes provide sample protection with a welded membrane and leak-proof cap, helping to minimize the risk of sample loss, whereas with traditional

tubing, samples can leak out if the clamp is loose/falls off. Lastly, cassettes are fast and efficient with high surface/volume ratio allowing for more rapid dialysis than conventional tubing.

There is an unseen internal gasket which reseals as the needle is withdrawn. The most important items to remember are:

  • Fill with air first to check that system is airtight
  • Puncture each porthole only once
  • Do not allow the membrane to come in contact with the sharp bevel of the needle.

No, the Slide-A-Lyzer™ Dialysis Cassette is designed as a single-use, disposable device. Once hydrated, the membrane immediately begins to dry after the sample is removed: and the whole membrane will be dry within a few minutes, changing the molecular weight cut-off (MWCO).

It is possible to add reagents to the Slide-A-Lyzer™ Dialysis Cassette at different times using different portholes, allow a reaction to occur, then dialyze in the same Slide-A-Lyzer™ Dialysis Cassette, reducing sample loss through reduced sample handling.

There are 3.5 K, 7K, and 10K MWCOs available in 0.1–0.5 mL, 0.5–3.0 mL and 3.0–12.0 mL. These units, especially at the smaller volumes, provide for greater sample recovery and shorter dialysis times than traditional tubing due to their greater surface area–to-volume ratio. For samples smaller than 100 µL, use the Slide-A-Lyzer™ MINI Dialysis Units and for larger volumes, refer to our SnakeSkin™ Dialysis Tubing.

The SnakeSkin™ Dialysis Tubing is ready-to-use, open, pleated form of regenerated cellulose dialysis tubing supplied in 8 inch sticks. You can cut off what you want, fold one end and close with a clip, add sample and close with a second clip. Different sizes of the tubing based on diameter (16, 22, 35 mm) of the tubing and MWCO (3.5K, 7K, 10K) are available. We also sell clips (Cat. No. 68011) and floats separately.

There are 3.5 K, 7K and 10K MWCOs available in 0.1–0.5 mL, 0.5–3.0 mL and 3.0–12.0 mL sizes. These units, especially at the smaller volumes, provide for greater sample recovery and shorter dialysis times than traditional tubing due to their greater surface area to volume ratio. For samples smaller than 100 µL, we recommend using the Slide-A-Lyzer™ MINI Dialysis Units or the Pierce™ 96-well microdialysis plates, and for larger volumes, we recommend the Slide-A-Lyzer™ Dialysis flasks (150–250 mL) or our SnakeSkin™ Dialysis Tubing.

There are 2K, 3.5K, 7K and 10K, and 20K MWCOs available in 0.1, 0.5 mL, and 2.0 mL size. Floats can also be purchased for this product (0.1 mL Cat. No. 69588).

The plastic casing is made of ABS plastic and the gasket is made from a non-leaching silicone-type material. The membrane is made from regenerated cellulose. We have successfully used 10% solutions of DMF, DMSO, acetonitrile, methanol, hexane, heptane, 100% acetic acid, and 70% ethanol with the device. The units are compatible with 75% acetonitrile and 1% TFA in the sample, however we recommend not dialyzing into a buffer of acetonitrile concentration greater than 10%.

No. The Slide-A-Lyzer™ Dialysis Cassette membrane is of a grade used for human kidney dialysis and requires no pretreatment. However, when using small volumes we do recommend pre-wetting the units in deionized water for 30 seconds. Trace metals and 12% glycerol are present in all Slide-A-Lyzer™ Dialysis Cassettes. During the dialysis process these will dialyze away to virtually no concentration remaining in the cassette.

The Slide-A-Lyzer™ Dialysis Cassette can be held at 56° C for 2 hours. At temperatures and times above these, the ABS plastic gets soft and can deform. We recommend maintaining the pH range of 5–9.

No, any 18 gauge needle at least one inch long, and syringes of sufficient volume will work.

The buoy plays two roles: it suspends the Slide-A-Lyzer™ Dialysis Cassette above the stirring bar during dialysis; it provides a stand to set the Slide-A-Lyzer™ Dialysis Cassette in on the bench-top

Typically, dialyzing three times for one hour each, at room temperature, in a 200-fold ratio of sample volume to buffer is sufficient for a simple buffer exchange (for a 1 mL starting sample, 200 µL x 200 µL x 200 µL = 8,000,000 µL or a dilution of 1/8,000,000). If required, this may be followed by an overnight dialysis at 4 degrees C to ensure complete dialysis.

However, each sample is different and the optimum times must be derived empirically.

The Slide-A-Lyzer™ Dialysis Cassette is the same regenerated cellulose as other dialysis tubing so you can expect about the same amount of protein loss as with tubing. As a guide, a 1 mg/mL solution will have a recovery rate of greater than 95%; at 100 mg/mL the recovery drops to 75–80%; and in solutions as dilute 10 μg/mL users may observe only a 50% (or less) recovery.

The MINI Dialysis Units are made of regenerated cellulose membrane, and the cup is made of polypropylene. The Units require 2 minutes’ hydration time before use.

Yes, the 0.1 mL devices can be placed into a foam float during dialysis, while the 0.5 mL and 2 mL sizes are capped by inserting them into the included 15 mL and 50 mL conical tubes, respectively.

In general, colder solutions take longer to dialyze. Dialysis completion will depend on the composition and volume of sample and dialysate. Simple pH exchange proceeds very rapidly; less than 10 minutes for 100 µL to change from pH 2.8 to pH 9.4. Dialysis of a 1 M salt solution against water proceeds very rapidly (less than 10 minutes for 10 µL). Dialysis of 100 µL of small compounds, 500–1,500 daltons, against a saline solution will be ~50% complete in 2–4 hours or 99%+ complete after overnight when dialyzing against ~1 liter of buffer. Dialysis will proceed faster with more frequent buffer changes.

The MINI Slide-A-Lyzer™ devices are manufactured in a clean environment. HEPA filters keep out contaminants. From molding to packaging, the MINI components have never been touched with ungloved (powder-free latex or nitrile) hands.

Yes, the MINI Slide-A-Lyzer™ device can be placed in a microfuge tube filled with concentrating solution. The average concentration rate of 35 µL/hour was observed in house when the sample was in PBS. We recommend using at least a 3:1 v/v ratio of concentrator to sample. In general, increased salt concentration in the sample will reduce the rate of sample  concentration.

Yes, but we’d suggest pre-treatment of the Slide-A-Lyzer™ device with EDTA or other DNase/RNase pre-treatment.

The plate comprises detachable strips so that you can pull out the exact number of wells needed for an experiment (scalable from 1–96 samples).

Typical protein recovery is greater than 90% after dialysis.

Most likely it’s because osmotic pressure from dialyzing from high osmolarity to low osmolarity solutions causes a sample volume increase, as each sample component—including water—will move towards equilibrium concentration on both sides of the membrane.

The Pierce™ concentrators are available with a polyethersulfone (PES) membrane, and are available in 0.5, 6, 20, and 100 mL volume capacities, with 3K or 5K, 10K, 30K, and 100K MWCOs.

Protein concentrators use ultrafiltration: hydrostatic pressure forcing a liquid against a semi-permeable membrane, with larger molecules retained within the unit. Dialysis works via diffusion across a semi-permeable membrane.

Please view our selection table to choose the right protein concentrator to fit your needs.

Equilibrium Dialysis

Equilibrium dialysis uses size-defined membranes to separate the free molecules (i.e., those molecules not bound to plasma proteins at equilibrium state). This type of dialysis mimics an in vivo environment. We offer our RED (rapid equilibrium dialysis) and competition RED systems to conduct plasma or tissue-to-plasma protein-binding studies.

Please use our selection table to choose the right plasma or tissue-to-plasma protein binding product for your experiment.

The membrane used is regenerated cellulose with a low glycerol content (acts as a humectant).

Yes. The Teflon™ Base Plate has the standard 96-well footprint. It can be used with any system compatible with standardized 96-well ELISA plates with 9 x 9 mm well spacing. The single-use format for the RED Device is especially convenient for labs using radioactive materials because the plate can easily be disposed of to minimize contamination and the need for cleaning.

No. The units do not undergo a sterilization procedure nor are they tested for endotoxin content.

No. They are manufactured in an ISO environment. The Teflon™ plate is also ISO certified.

In most cases, 4 hours is sufficient to reach equilibrium. This is due to the high surface area–to-volume ratio for the membrane (7.4:1), but can vary with different compounds.

The container for the device is Teflon™ (PTFE), the insert is made of high-density polyethylene (HDPE), and they are highly hydrophobic. The regenerated cellulose membrane is a standard material for commercial dialysis devices. A recovery study consistently shows 85% recovery of high and low protein binding compounds. This result is indicative of minimal nonspecific binding, since the recovery between the membrane side and the PTFE/HDPE housing side showed very small difference.

Plasma samples from human, mouse, and rat are typically tested for binding of ligands with the rapid equilibrium dialysis (RED) device. For toxicology studies, a monkey sample is also tested. Typically, pooled plasma samples purchased from commercial vendors are used, although researchers could test the differences in plasma from various physiological states using the RED devices.

We have not tested the units with cells or viral particles. Customers would have to design and optimize their experiments to use the RED device with this type of sample.

Each sample requires two chambers for equilibrium dialysis to take place (96/2 = 48); one chamber for the plasma sample, the other for buffer.

Protein Desalting

Desalting is the process where porous particles separate molecules of different sizes. Desalting products use gel filtration or size exclusion. Small molecules enter into the pores in the resin, resulting in a longer path length through the desalting column when compared to large molecule. This increased path length for molecules below the MWCO allows the large molecules to pass through the resin more quickly.

Our desalting resins are rated by what MWCO molecules can pass through. They also have a lower MWCO (1K or 2K) that can be effectively separated from larger molecules. Molecules in between these values cannot be effectively separated from larger or smaller molecules, given our protocols.

A centrifuge is used to first remove the resin’s void volume of liquid, followed by sample addition and centrifugation. After centrifugation, the macromolecules in the sample have moved through the column in approximately the same initial volume, but the small molecules have been forced into the pores of the resin and replaced by the buffer that was used to pre-equilibrate the gel-filtration matrix. Spin formats eliminate the need to wait for samples to emerge by gravity flow and require no chromatography system, allowing for multiple-sample processing simultaneously.

Note: The addition of larger volumes of buffer or longer centrifugation times than listed in the protocol will result in smaller molecules eventually emerging from the desalting column.

Yes, Zeba™ columns are compatible with most salts. The resin is stable to some organics. As organics may affect performance, we suggest using <10% organics.

Yes, the 7K Zeba™ columns have good recovery of dsDNA ladder down to 10 bp.

No, excess amounts of DyLight™ dyes (and any other planar molecule with multiple rings in their structure) “act” lager than their stated molecular weights in desalting columns. We do not recommend desalting, but refer researchers to our Pierce™ Dye Removal Column (Cat. No. 22585) for this purpose.

The desalting columns are designed to be used once and discarded after use.

Centrifuged-based desalting results in minimal dilution. A slight dilution of the protein solution results if the optional stacker buffer is added that ensures maximum protein recovery.

When used properly, the volume recovered from the column equals the volume of sample (and buffer) added to the column.

Glycerol and sugars add viscosity to a sample, while detergents create micelles, making the removal of each difficult. Although dialysis is a better choice for removing these components, Zeba™ Columns can be used by modifying the desalting protocol. Modifications include diluting the samples, processing smaller samples, and adding the sample slowly to the column allowing it to enter the gel and equilibrate into the pores before centrifugation.

With Pierce Peptide Desalting Spin Columns, the reproducibility coefficient of variation (CV) is ± 20%.

With Pierce Peptide Desalting Spin Columns, more than 90% of salt or labeling tags such as TMT tags can be removed.

Protein Concentration

The products are available in 4 sizes (0.5 mL, 6 mL, 20 mL, and 100 mL) with a range of molecular weight cutoffs (3K, 5K, 10K, 30K, and 100K). See our selection chart.

Typically, samples can be concentrated up to 10- to 30-fold in 5–30mins. Times may vary for different MWCOs and the buffer composition of the sample.

The concentrators are compatible with certain solvents. Please see the table below:

Chemical compatibility of Pierce PES concentrators

Acids and bases






Acetic acid (25%)




Ammonium sulphate (saturated)


Formic acid (5.0%)




Glycerine (70%)


Hydrochloric acid (1M)


Benzene (100%)


Guanidine HCl (6M)


Lactic acid (5.0%)


Chloroform (1%)


Imidazole (300mM)


Nitric acid (10%)


Dimethyl sulfoxide (5.0%)


Phosphate buffer (1.0M)


Sodium hydroxide (2.5M)


Ethanol (70%)


Polyethylene glycol (10%)


Sulfamic acid (5.0%)


Ethyl Acetate (100%)


Sodium carbonate (20%)


Trifuoroacetic acid (10%)


Formaldehyde (30%)


Sodium deoxycholate (5.0%)




Hydrocarbons (aromatic)


Sodium dodecylsulfate (0.1M)




Hydrocarbons (chlorinated)


Sodium hypochlorite (200ppm)




Isopropanol (70%)


Sodium nitrate (1.0%)




Mercaptoethanol (1.0M)


Tween™ 20 (0.1%)




Pyridine (100%)


Triton™ X-100 (0.1%)




Tetrahydrofuran (5.0%)


Urea (8M)




Toluene (1.0%)




A = Acceptable. NR = Not Recommended. Concentrations listed are provided as guidelines and do not necessarily represent maximum tolerances.