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Lysis buffers differ in their ability to solubilize proteins, with those containing sodium dodecyl sulfate (SDS) and other ionic detergents considered to be the harshest and therefore most likely to give the highest yield.
The main consideration when choosing a lysis buffer is whether the chosen antibody will recognize denatured samples. When this is not the case, it will be noted on the antibody datasheet, and buffers without detergent or with relatively mild non-ionic detergents (NP-40, Triton X-100) should be used.
Protein location and lysis buffer choice
|Protein location||Buffer recommended|
|Cytoplasmic (cytoskeletal bound)||Tris-Triton|
|Membrane bound||NP-40 or RIPA|
|Nuclear||RIPA or use nuclear fraction protocol*|
|Mitochondria||RIPA or use mitochondrial fraction protocol*|
*Proteins that are found exclusively or predominantly in a sub-cellular location will be more enriched in a lysate of the sub-cellular fraction compared with whole cell or tissue lysates. This can be useful when trying to obtain a signal for a weakly-expressed protein. Please consult our separate protocols for sub-cellular fractionation.
Lysis buffer recipes:
This is a popular buffer for studying proteins that are cytoplasmic or membrane-bound, or for whole cell extracts. If there is concern that the protein of interest is not being completely extracted from insoluble material or aggregates, RIPA buffer may be more suitable as it contains ionic detergents that will more readily bring the proteins into solution.
RIPA buffer (radioimmunoprecipitation assay buffer)
*Can be prepared as a 10% stock solution, which must be protected from light.
RIPA buffer is useful for whole cell extracts and membrane-bound proteins, and may be preferable to NP-40 or Triton X-100-only buffers for extracting nuclear proteins. It will disrupt protein-protein interactions and may therefore be problematic for immunoprecipitations and pull-down assays.
In cases where it is important to preserve protein-protein interactions or to minimize denaturation, a buffer without ionic detergents (eg SDS) and ideally without non-ionic detergents (eg Triton X-100) should be used.
Cell lysis with detergent-free buffer is achieved by mechanical shearing, often with a Dounce homogenizer or by passing cells through a syringe tip. In these cases, a simple Tris buffer will suffice, but as noted above, buffers with detergents are required to release membrane- or cytoskeleton-bound proteins.
Tris-Triton buffer (cytoskeletal proteins)
All four of these buffers will keep at 4°C for several weeks or for up to a year if divided into aliquots and stored at -20°C.
As soon as lysis occurs, proteolysis, dephosphorylation and denaturation begin. These events can be slowed down significantly if samples are kept on ice or at 4°C at all times and appropriate inhibitors are added fresh to the lysis buffer.
Ready-to-use cocktails of inhibitors from various suppliers are available but you can make your own cocktail.
|Final concentration in lysis buffer||Stock (store at -20°C)|
|Aprotinin||Trypsin, chymotrypsin, plasmin||2 µg/mL||Dilute in water, 10 mg/mL. Do not re-use thawed aliquots.|
|Leupeptin||Lysosomal||5–10 µg/mL||Dilute in water. Do not re-use thawed aliquots.|
|Pepstatin A||Aspartic proteases||1 µg/mL||Dilute in methanol, 1 mM.|
|PMSF||Serine, cysteine proteases||1 mM||Dilute in ethanol. You can re-use the same aliquot.|
|EDTA||Metalloproteases that require Mg2+ and Mn2+||5 mM||Dilute in dH20, 0.5 M. Adjust pH to 8.0.|
|EGTA||Metalloproteases that require Ca2+||1 mM||Dilute in dH20, 0.5 M. Adjust pH to 8.0|
|Sodium fluoride||Serine/threonine phosphatases||5–10 mM||Dilute in water. Do not re-use once defrosted.|
|Tyrosine phosphatases||1 mM||Dilute in water. Do not re-use once defrosted.|
Sodium orthovanadate preparation
Perform all steps in a fume hood.
Denatured, reduced samples
Antibodies typically recognize a small portion of the protein of interest (referred to as the epitope) and this domain may reside within the 3D conformation of the protein. To enable access of the antibody to this portion it is necessary to unfold the protein, ie denature it.
To denature, use a loading buffer with the anionic detergent sodium dodecyl sulfate (SDS), and boil the mixture at 95–100°C for 5 min. Heating at 70°C for 5–10 min is also acceptable and may be preferable when studying multi-pass membrane proteins. These tend to aggregate when boiled and the aggregates may not enter the gel efficiently.
The standard loading buffer is called 2X Laemmli buffer (Laemmli UK, 1970. Cleavage of structural proteins during the assembly of the head of bateriophage T4. Nature, 227, 680–5). It can also be made at 4X and 6X strength to minimize dilution of the samples. The 2X is to be mixed in 1:1 ratio with the sample.
2x Laemmli buffer recipe
When SDS is used with proteins, all of the proteins become negatively charged by their attachment to the SDS anions. SDS binds to proteins fairly specifically in a mass ratio of 1.4:1. In doing so, SDS confers a negative charge to the polypeptide in proportion to its length. Denatured polypeptides become rods of negative charge with equal charge densities per unit length. Therefore, migration is determined by molecular weight, rather than by the intrinsic charge of the polypeptide.
SDS grade is important for high-quality protein separation: a protein stained background along individual gel tracts with indistinct or slightly distinct protein bands are indicative of old or poor quality SDS. Inclusion of 2-mercaptoethanol or dithiothreitol in the buffer reduces disulphide bridges, which is necessary for separation by size.
Glycerol is added to the loading buffer to increase the density of the sample to be loaded and hence maintain the sample at the bottom of the well, restricting overflow and uneven gel loading.
To visualize the migration of proteins it is common to include a small anionic dye molecule in the loading buffer (eg bromophenol blue). Since the dye is anionic and small, it will migrate the fastest of any component in the mixture to be separated and provide a migration front to monitor the separation progress.
During protein sample treatment the sample should be mixed by vortexing before and after the heating step for best resolution.
Native and non-reduced samples
Alternatively, an antibody may recognize an epitope made up of non-contiguous amino acids. Although the amino acids of the epitope are separated from one another in the primary sequence, they are close to each other in the folded three-dimensional structure of the protein, and the antibody will only recognize the epitope as it exists on the surface of the folded structure.
In these circumstances, it is important to run a western blot in non-denaturing conditions, and this will be noted on the datasheet in the applications section. In general, a non-denaturing condition simply means leaving SDS out of the sample and migration buffers and not heating the samples.
Certain antibodies only recognize protein in its non-reduced form (particularly on cysteine residues) and the reducing agents β-mercaptoethanol and DTT must be left out of the loading buffer and migration buffer.
|Protein state||Gel condition||Loading buffer||Migration buffer|
|Reduced, denatured||Reducing and denaturing||With 2-mercaptoethanol or DTT and SDS||With SDS|
|Reduced, native||Reducing and native||With 2-mercaptoethanol or DTT and SDS||No SDS|
|Oxidized, denatured||Non-reducing and denaturing||No 2-mercaptoethanol or DTT, with SDS||With SDS|
|Oxidized, native||Non-reducing and native||No 2-mercaptoethanol or DTT, with SDS||No SDS|
Rule of thumb: reduce and denature unless the datasheet specifies otherwise.
Protocols are provided by Abcam “AS-IS” based on experimentation in Abcam’s labs using Abcam’s reagents and products; your results from using protocols outside of these conditions may vary.