This works when the proteins involved in the complex bind to each other tightly, making it possible to pull multiple members of the complex out of solution by latching onto one member with an antibody. This concept of pulling protein complexes out of solution is sometimes referred to as a "pull-down", although this term should be reserved for use in systems where no antibody is required. Co-IP is a powerful technique that is used regularly by molecular biologists to analyze protein-protein interactions.
Identifying the members of protein complexes may require several rounds of precipitation with different antibodies for a number of reasons:
♦ A particular antibody often selects for a subpopulation of its target protein that has the epitope exposed, thus failing to identify any proteins in complexes that hide the epitope. This can be seen in that it is rarely possible to precipitate even half of a given protein from a sample with a single antibody, even when a large excess of antibody is used.
♦ The first round of IP will often result in the identification of many new proteins that are putative members of ta complex being studied. The researcher will then obtain antibodies that specifically target one of the newly identified proteins and repeat the entire immunoprecipitation experiment. This second round of precipitation may result in the recovery of additional new members of a complex that were not identified in the previous experiment. As successive rounds of targeting and immunoprecipitations take place, the number of identified proteins may continue to grow. It should be noted that the identified proteins may not ever exist in a single complex at a given time, but may instead represent a network of proteins interacting with one another at different times for different purposes.
♦ Repeating the experiment by targeting different members of the protein complex allows the researcher to double-check the result. Each round of pull-downs should result in the recovery of both the original known protein as well as other previously identified members of the complex (and even new additional members). By repeating the immunoprecipitation in this way, the researcher verifies that the each identified member of the protein complex was a valid identification. If a particular protein can only be recovered by targeting one of the known members but not by targeting other of the known members then that protein's status as a member of the complex may be subject to question.
The principle underpinning this assay is that DNA-binding proteins (including transcription factors) in living cells can be cross-linked to the DNA that they are binding. By using an antibody that is specific to a putative DNA binding protein, one can immunoprecipitate the protein-DNA complex out of cellular lysates. The crosslinking is often accomplished by applying formaldehyde to the cells (or tissue), although it is sometimes advantageous to use a more defined and consistent crosslinker such as DTBP. Following crosslinking, the cells are lysed and the DNA is broken into pieces 0.2-1 kD in length by sonication. At this point the immunoprecipitation is performed resulting in the purification of protein-DNA complexes. The purified protein-DNA complexes are then heated to reverse the formaldehyde cross-linking of the protein and DNA complexes, allowing the DNA to be separated from the proteins. The identity and quantity of the DNA fragments isolated can then be determined by PCR. The limitation of performing PCR on the isolated fragments is that one must have an idea which genomic region is being targeted in order to generate the correct PCR primers. This limitation is very easily circumvented simply by cloning the isolated genomic DNA into a plasmid vector and then using primers that are specific to the cloning region of that vector. Alternatively, when one wants to find where the protein binds on a genome-wide scale, a DNA microarray can be used (ChIP-on-chip or ChIP-chip) allowing for the characterization of the cistrome. As well, ChIP-Sequencing has recently emerged as a new technology that can localize protein binding sites in a high-throughput, cost-effective fashion.
The two general methods for immunoprecipitation are the direct capture method and the indirect capture method.
From this point on, the direct and indirect protocols converge because the samples are now identical. Either scenario gives the same end-result with the protein or protein complexes bound to the antibodies which themselves are immobilized onto the beads.
Historically the solid-phase support for immunoprecipitation used by the majority of scientists has been highly-porous agarose beads (also known as agarose resins or slurries). The advantage with this technology is a very high potential binding capacity as virtually the entire sponge-like structure of the agarose particle is available for binding antibodies (which will in turn bind the target proteins). This advantage of extremely high binding capacity must be carefully balanced with the quantity of antibody that the researcher is prepared to use to coat the agarose beads. Because antibodies can be a cost-limiting factor, it is best to calculate backward from the amount of protein that needs to be captured (depending upon the analysis to be performed downstream), to amount of antibody that is required to bind that quantity of protein (with a small excess added in to acount for inefficiencies of the system), and back still further to the quantity of agarose that is needed to bind that particular quantity of antibody, and no more. In cases where antibody is not a cost-limiting factor, this technology is unmatched in its ability to capture extremely large quantities of captured target proteins. The caveat here is that the "high capacity advantage" can become a "high capacity disadvantage". This "high capacity disadvantage" is manifested when the enormous binding capacity of the sepharose/agarose beads is not completely saturated with antibodies. It often happens that the amount of antibody available to the researcher for the their immunoprecipitation experiment is less than sufficient to saturate the agarose beads to be used in the immunoprecipitation. In these cases the researcher will end up with a partially antibody-coated agarose particles. The portion of the binding capacity of the agarose beads that is not coated with antibody will then be free to bind anything that will stick. This results in an elevated level of random non-specifically bound proteins to the beads which results in an elevated background signal that can make it more difficult to interpret results. For these reasons it is prudent to match the quantity of agarose (in terms of binding capacity) to the quantity of antibody that one wishes to be bound for the immunoprecipitation.
In most cases, it is best to perform an additional step known as preclearing (see step 2 in the "protocol" section below) at the start of each immunoprecipitation experiment. Preclearing simply refers to the addition of uncoated agarose beads to the protein mixture in an effort to bind and remove proteins that will non-specifically bind to the uncoated agarose. The beads used for preclearing will be discarded after a suitable incubation period. The hope after the preclearing step is that the non-specific agarose binders have all been removed from the protein mixture. Then, when the antibody-coated agarose beads are introduced to the protein mixture, non-specific binding to the beads will be reduced.
Monodisperse superparamagnetic beads are also available as a support material which offers certain advantages over the agarose beads which are polydisperse. One such advantage is the ability to bind extremely large protein complexes and the complete lack of an upper size limit for such complexes. This is due to inherent differences in the technologies. Whereas agarose beads are sponge-like porous particles of variable size, magnetic beads small, solid and (in the case of monodisperse magnetic beads) spherical and uniform in size. Because all of the antibody binding capacity is on the outer solid surface of these beads, all of the binding of proteins during the immunoprecipitation will occur on the bead outer surface thereby eliminating an upper size limit. However, this same "feature" results in a limited surface area and thus a reduction in binding capacity. This can be compared to the agarose beads which have higher potential binding capacity, but also have a limited capacity for large complexes which are unable to fit into the pores of the beads. Other characteristics of surface-only binding are faster binding kinetics as protein complexes are not required to penetrate deeply into a porous particle in order to be captured. Several notable disadvantages of magnetic bead technology exist. Magnetic particles are much more expensive than their agarose counterparts, they cannot compete with agarose for total binding capacity (as noted above), some fraction of the beads (and attached proteins)is generally lost from the magnet during the procedure, and they at times suffer from clumping and a tendency to fall out of solution. It should be noted that the lower overall binding capacity of magnetic beads for immunoprecipitation may make it easier to match the quantity of antibody needed for diagnostic immunoprecipitations precisely with the total available binding capacity on the beads which results in decreased background and fewer false positives. The increased reaction speed of the immunoprecipitations when using magnetic bead technologies results in higher yields of labile (fragile) protein complexes due to the reduction in protocol times and sample handling requirements which reduces physical stresses on the samples and reduces the time that the sample is exposed to potentially damaging proteases all of which contribute greatly to increasing the yield (in terms of numbers of protein complex members identified) when performing immunoprecipitation on protein complexes. Agarose bead-based immunoprecipitations may also be performed more quickly using small spin columns to contain the agarose resin and quickly remove unbound sample or wash solution with a brief centrifugation.
With superparamagnetic beads, the sample is placed in a magnetic field so that the beads can collect on the side of the tube. This procedure is generally complete in approximately 30 seconds and the remaining (unwanted) liquid is pipetted away. Washes are accomplished by resuspending the beads (off the magnet) with the washing solution and then concentrating the beads back on the tube wall (by placing the tube back on the magnet). The washing is generally repeated several times to ensure adequate removal of contaminants. If the superparamagnetic beads are homogeneous in size and the magnet has been designed properly, the beads will concentrate uniformly on the side of the tube and the washing solution can be easily and completely removed.
When working with agarose beads the beads must be pelleted out of the sample by briefly spinning in a centrifuge with forces between 600-3,000 x g (times the standard gravitational force). This step may be performed in a standard microcentrifuge tube, but for faster separations, greater consitency and higher recoveries, the process is often peformed in small spin columns with a pore size that aloows liquid, but not agarose beads to pass through. After centrifugation, the agarose beads will form a very loose fluffy pellet at the bottom of the tube. The supernatant containing contaminants can be carefully removed so as not to disturb the beads. The wash buffer can then be added to the beads and after mixing, the beads are then pelleted out of the wash solution by re-centrifuging the sample.
Following the initial capture of a protein or protein complex with either bead type, the solid support is washed several times to remove any proteins not specifically and tightly bound to the support through the antibody. After washing, the precipitated protein(s) are eluted and analyzed using gel electrophoresis, mass spectrometry, western blotting, or any number of other methods for identifying constituents in the complex. Thus, co-immunoprecipitation is a standard method to assess protein-protein interactions.
Protocol times for immunoprecipitation vary greatly due to a variety of factors, with protocol times increasing with the number of washes necessary or with the slower reaction kinetics of porous agarose beads. The vast majority of immunoprecipitations are performed using agarose beads. The use of magnetic beads for immunoprecipitaion is a much newer approach that is only recently gaining in popularity.