This approach enables researchers to:
- identify the activation status of proteins
- determine post-translational protein modifications
- measure the molecular weight of a given protein
- capture protein-binding molecules in the study of protein-protein and protein-nucleic acid interactions
Target antigens are usually immunoprecipitated from complex solutions, such as cell lysates. The basic principle of an IP is diagrammed below. An antibody (monoclonal or polyclonal) against a specific protein is pre-immobilized onto an insoluble support, such as agarose or magnetic beads, and then incubated with a cell lysate containing the target protein. During the incubation period, gentle agitation of the lysate allows the target antigen to bind to the immobilized antibody. The immobilized immune complexes are then collected from the lysate, eluted from the support and analyzed based on the nature of the target antigen.
Alternatively, free, nonbound antibody is allowed to form immune complexes in the lysate and then the complexes are retrieved by the insoluble support. While the pre-immobilized antibody approach is more commonly used for IP, using free antibody to form immune complexes is beneficial if the target protein is present in low concentrations, the antibody has a weak binding affinity for the antigen or the binding kinetics of the antibody to the antigen are slow.
Traditionally, single proteins have been isolated from cell lysates by immunoprecipitation to investigate the identity, structure, expression, activation or modification of the individual proteins. IPs are also used to study the interaction of a target protein with other proteins or nucleic acids. Variations in the basic IP method provide the flexibility to perform a broad range of IP applications.
Co-immunoprecipitation (Co-IP) is a popular technique for protein interaction discovery. Co-IP is conducted in essentially the same manner as an IP, except that the target antigen precipitated by the antibody, also called the "bait", is used to co-precipitate a binding partner/protein complex, or "prey", from a lysate; i.e., the interacting protein is bound to the target antigen, which is bound by the antibody that is immobilized onto the support. Immunoprecipitated proteins and their binding partners are commonly detected by SDS-PAGE and Western blot analysis. The assumption that is usually made when associated proteins are co-precipitated is that these proteins are related to the function of the target antigen at the cellular level. This is only an assumption, however, that is subject to further verification.
Chromatin immunoprecipitation (ChIP) assays are performed to identify regions of the genome with which DNA-binding proteins, such as transcription factors and histones, associate. In ChIP assays, proteins bound to DNA are temporarily crosslinked and the DNA is sheared prior to cell lysis. The target proteins are immunoprecipitated along with the crosslinked nucleotide sequences, and the DNA is then removed and identified by PCR, sequenced, applied to microarrays or analyzed in some other way.
This approach is similar to ChIP, except that RNA-binding proteins are immunoprecipitated instead of DNA-binding proteins. Immunoprecipitated RNAs can then be identified by RT-PCR and cDNA sequencing.
A key limitation of the previously-described IP approaches is their dependence upon the availability of antibodies that specifically recognize the target protein with little or no cross-reactivity with other cellular targets. Due to this limitation, many proteins are unable to be immunoprecipitated because of the lack of an available antibody.
To circumvent this problem, proteins can be tagged with an epitope to which a high-affinity antibody is available and ectopically expressed in the cell of interest. Today, this approach is commonplace for all types of immunoprecipitations in molecular biology research. These tags can be either short peptide sequences or fluorescent proteins, including:
- Flag; peptide sequence DYKDDDDK
- c-Myc; peptide seqence EQKLISEEDL
- Hemagglutinin (HA); peptide sequence YPYDVPDYA
- Green fluorescent protein (GFP)
One downside of using tagged proteins is that the overexpressed tagged protein, not the endogenous protein, is immunoprecipitated, which limits the applicability of any findings using this approach to true biological relevance. Additionally, tagging the protein may interfere with protein function.
However, growth factors and conditions affecting a specific protein-protein interaction in cultured cells can be measured very precisely using a quantitative immunoprecipitation (qIP) method based on co-expression of epitope-tagged and luciferase-tagged proteins.
Although IP methods are logically and procedurally simple, the variables affecting the success of any specific experiment are as numerous and peculiar as the specific differences between individual proteins and different primary antibodies. Empirical testing is nearly always required to optimize IP conditions to obtain the desired yield and purity of target proteins. Nevertheless, consideration of the main factors involved can help to identify the components that are most likely to affect particular experiments.
Also known as agarose resin, this is the most prevalent form of support used in research-scale IP applications. Although the name implies solid, spherical structures, agarose beads are actually sponge-like structures of varying shapes and sizes (50 to 150μm) that can be modified for activation or coupling to an appropriate ligand. This highly porous material has the highest antibody binding capacity of all available IP support materials because all surfaces, both on the surface and within each structure, are available for binding.
The resin is durable and robust, able to withstand centrifugation up to 5000 x g, pressures of 100 psi (depending on the degree of crosslinking) and temperatures up to 120°C without significant loss of structure or flow rate. Agarose exhibits low nonspecific binding in complex samples and is not harmed by moderate levels of most detergents, salts, most organic solvents or extremes of pH (will tolerate pH range of 3 to 11 for extended periods without suffering measurable hydrolysis).
Magnetic particles, unlike agarose beads, are solid and spherical, and antibody binding is limited to the surface of each bead. While these beads do not have the advantage of a porous center to increase the binding capacity, magnetic beads are signficantly smaller than agarose beads (1 to 4μm), which collectively gives magnetic beads adequate surface area-to-volume ratios for optimum antibody binding.
High-power magnets are used to localize magnetic beads to the side of the incubation tube and out of the way to enable cell lysate aspiration without the risk of also aspirating immune complexes bound to the beads. Magnetic separation avoids centrifugation, which can break weak antibody-antigen binding and cause loss of target protein. However, an issue with the use of magnetic beads is that bead size variations may prevent all beads from localizing to the magnet. Additionally, while IP using agarose beads only requires standard laboratory equipment, the use of magnetic beads for IP applications requires high-power magnetic equipment that can be cost-prohibitive.
Protein A, Protein G, Protein A/G and Protein L are immunoglobulin (Ig)-binding proteins that, when attached to beaded support and used as affinity ligands, comprise the most popular antibody-binding platforms for IP applications. As shown in the diagram below, Proteins A and G both show specificity for the heavy chains on the Fc region of antibodies, which effectively orients the immobilized antibodies with antigen-binding sites facing outward; Protein G also shows some affinity for Fab fragments (1,3). Protein L binds to light chains, but because of specific binding characteristics, Protein L is only used for limited applications.
Most immunoprecipitations are performed with Protein A, G or Protein A/G, which is an engineered recombinant protein combining four Protein A and two Protein G antibody binding sites. Protein A and G both show high affinity for antibodies of multiple, but not necessarily identical, subclasses and Ig species, while Protein A/G binds all of the subtypes to which Protein A and G individually bind.
Immobilized Protein A, G and A/G (hereafter collectively called "Protein A/G") are effective tools for attaching antibodies to a beaded support for IP applications. Innovations in manufacturing of prepared Protein A/G resins have yielded commercially available supports that have very high binding capacities, enabling excellent immunoprecipitation results to be obtained with very small volumes of beads. Binding capacities of 30 to 50mg Ig per mL resin with very low nonspecific binding are possible with these affinity resins.
Protein A/G supports are not compatible with certain IP experimental systems, such as when an incompatible antibody species or subclass is used or when immunoprecipitating from serum, which contains nonspecific immunoglobulins that would compete with the IP antibody for binding to the support.
Covalent immobilization strategies chemically bind the antibody to the beaded support and remove the requirement for Protein A/G-dependent antibody immobilization. Commercial products are available that provide beaded supports that react with primary amines (-NH2) on the antibody to permanently bind the antibody to the support. Although this method couples antibodies in random orientation (based on whichever surface amines contact the reactive groups on the beads), this usually has only slight effects on the antigen-binding function and capacity of the IP antibody. Besides eliminating the dependence on Protein A/G, the direct immunoprecipitation method prevents the IP antibody from co-eluting with the antigen (when a non-reducing elution buffer is used) and interfering in SDS-PAGE analysis. Additionally, because the antibody theoretically remains intact and permanently bound to the support, it may be possible to reuse the antibody-coated support many times.
It is also possible to covalently attach antibodies to Protein A/G-bound supports using a crosslinker. Examples of such crosslinkers include DSS and BS3, which are short carbon chains with reactive N-Hydroxysuccinimidyl (NHS) ester groups at each end. NHS esters react with primary amines (side chain of lysine residues in proteins) to form covalent amide bonds. If an antibody is first bound to a Protein A/G support and then mixed with a crosslinker solution, the crosslinker molecules can react to covalently link adjacent amines of the antibody and Protein A/G.
As with the direct immobilization method, the crosslink method eliminates co-elution of antibody fragments and potentially enables the antibody support to be reused several times. For obvious reasons, this method can only be used for antibodies that successfully bind to Protein A or G. Because antibodies contain multiple amine groups that are not exclusively limited to the Fc region, it is important to optimize the dosage of crosslinker. If too little or too much crosslinker is used, the antibody may not become successfully linked to the Protein A/G agarose or too many of the amine groups in the antibody binding site may become modified, rendering the antibody unable to bind antigen.
Immunoprecipitation as performed by the batch method simply involves mixing all the components of the reaction in a reaction vessel (usually a microcentrifuge tube) for a period of time to allow the immune complexes to form and bind to the beaded resin. At each step, the beads are pelleted by centrifugation and the solutions (nonbound sample and buffers) are carefully aspirated. Column methods involve incubating IP components with beaded resin that is packed in a plastic or glass column. The sample is either allowed to pass through the column by gravity or centrifugation (see next paragraph) or the column is capped and the sample incubated with the resin (with optional mixing) to allow the antibody and antigen more time to bind. In either case, the sample solutions are separated from the beads by gravity-flow or centrifugal collection from the column tip.
Large scale IPs (>10mL resin) are generally limited to gravity-flow because of the impracticality of centrifuging large columns, especially if they are not designed to fit into a collection tube. Conversely, very small-scale applications require centrifugation, as small volumes of just a few microliters of solution will not flow through a filter by gravity alone. Most medium-scale IPs can be performed by either gravity-flow or centrifugation so long as suitable columns and collection tubes are available and the beaded support is compatible with the increased pressures associated with centrifugation.
The use of spin columns has a distinct advantage over both gravity columns and batch methods because almost all of the residual solution can be spun through the filter allowing cleaner separation of the solid and aqueous phases. Gravity columns require constant monitoring to make sure the resin does not run dry and form air bubbles. In addition, the antigen is eluted in multiple fractions, each of which must be monitored for the presence of antigen. Fractions containing antigen are normally pooled, therefore the volume will end up being much greater than the original sample and the antigen may require concentration. A disadvantage of the batch method is the formation of the resin pellet, which contains a significant volume of solution that cannot be removed by pipetting; additional wash and elution steps are necessary to obtain good purity and yield.
The quality of the sample that is used for IP applications critically depends on the right lysis buffer, which stabilizes native protein conformation, inhibits enzymatic activity, minimizes antibody binding site denaturation and maximizes the release of proteins from the cells or tissue. The lysis buffer used for a particular application depends on the target proteins that will be immunoprecipitated, because the location of the protein in the cell (e.g., membrane, cytosol, nucleus) affects the ease of release during lysis.
Non-denaturing buffers are used when the IP antigen is detergent-soluble and when the antibody can recognize the native form of the protein. These buffers contain non-ionic detergents, such as NP-40 or Triton X-100. Denaturing buffers, such asRadioimmunoprecipitation Assay (RIPA) buffer, are more stringent than non-denaturing buffers because of the addition of ionic detergents like SDS or sodium deoxycholate. While these buffers do not maintain native protein conformation, proteins that are difficult to release with non-denaturing buffers, such as nuclear proteins, can be released with denaturing buffers. Both buffers contain NaCl and Tris-HCl and have a slightly basic pH (7.4 to 8). Detergent-free buffers can also be used if the target protein can be released from cells using only physical disruption, such as mechanical homogenization or heat. These simple buffers usually consist of just EDTA in phosphate buffered saline (PBS). See the table below for the ranges of each component to aid in protocol optimization.
Because cell lysates also contain proteases and phosphatases that can modify or degrade the target protein, most IP protocols are performed at 4°C. Proteasomal inhibitors, such as PMSF, aprotinin and leupeptin are commonly added to the lysis buffer just prior to use, along with sodium orthovanadate or sodium fluoride as aphosphatase inhibitor. While these components can be added individually, commercial inhibitor cocktails are available that are higher quality and easier to use.
|Non-ionic detergents (NP-40, Triton X-100)||0.1 to 2%|
|Ionic detergents (SDS, sodium deoxycholate)||0.01 to 0.5%|
|NaCl (sodium chloride, salt)||0 to 1M|
|Divalent cations||0 to 10mM|
|pH||6 to 9|
|EDTA||0 to 5mM|
Because lysates are complex mixtures of proteins, lipids, carbohydrates and nucleic acids, one must assume that some amount of nonspecific binding to the IP antibody, Protein A/G or the beaded support will occur and negatively affect the detection of the immunoprecipitated target(s).
Preclearing the sample is an optional step designed to remove potentially reactive components from the lysate prior to the immunoprecipitation steps. The basic approach to preclear a lysate is to incubate the sample with exactly the same components that will be used for the immunoprecipitation, except use a nonspecific antibody from the same species as the IP antibody. Any nonspecific immune complexes will form and be immobilized to the beaded support. Additionally, if Protein A/G or agarose beads are used, this approach will allow nonspecific binding to these IP components, which along with the nonspecific immune complexes are removed from the lysate. If successful, the nonspecific lysate products will be removed by this preclearing step so that they will not co-purify with the target antigen in the actual IP experiment.
These nonspecific immune complexes may be used as a negative control for an IP or co-IP experiment; any products obtained with these control conditions can be attributed to nonspecific (off-target) interactions. One advantage of the direct immobilization strategy described above is the lack of Protein A/G as a component, which is a potential source of nonspecific binding interactions in the assay system.
During the immunoprecipitation, assembling the immune complexes and maintaining complex stability depend on the compatibility of the binding buffer with all of the component binding interactions. In most cases, antibody-antigen interactions are fairly robust and will occur in any standard ionic strength buffer of near-neutral pH, such as PBS. By contrast, bait-prey interactions range in strength and time from irreversible and long-lived to labile and transient, which will be influenced by both the binding conditions and the temporal completion of protocol steps.
Even after lysate preclearing, IP components will still pull down nonspecific cellular components that must be removed by gentle washes prior to sample elution. Multiple washes with simple wash buffers, such as PBS either alone or with low detergent concentrations or by moderate adjustments to salt concentration, can be used to remove these contaminants.
Traditional IP for downstream analysis by reducing SDS-PAGE and Western blot detection typically involves elution directly in reducing SDS-PAGE sample buffer. This buffer, which is designed to denature and reduce proteins for electrophoresis, is very effective in dissociating the affinity interactions upon which IP is based. Other downstream applications for IP products are not compatible with this buffer system, nor is it possible to take advantage of certain IP methods, such as antigen elution, without antibody fragment contamination in the direct or crosslink IP methods, when this elution buffer condition is used.
The most generally effective, nondenaturing elution buffer for protein affinity purification methods is 0.1 M glycine at pH 2.5 to 3. The low pH condition dissociates most antibody-antigen interactions, as well as the antibody-Protein A/G interaction, assuming that it has not been crosslinked. Low-pH glycine is not universally effective; some antibody-antigen interactions do not dissociate with this buffer, and conversely, some antibodies and target antigens denature or become inactive in this buffer. Several alternative types of elution buffers are described in greater detail in Tech Tip #27.
- Bjorck L. and Kronvall G. (1984) Purification and some properties of streptococcal protein g, a novel IgG-binding reagent. J Immunol. 133, 969-74.
- Harlow, Ed, and Lane, David. (1999) Using Antibodies. Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press.
- Wikstrom M. et al. (1995) Mapping of the immunoglobulin light chain-binding site of protein l. J Mol Biol. 250, 128-33.
For Research Use Only. Not for use in diagnostic procedures.