co ip experiment protocol

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Co-immunoprecipitation (Co-IP) Overview

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Introduction

The intricacies of cellular processes can be unraveled through the study of protein-protein interactions in the discipline of proteomics. A crucial tool in this field is Co-immunoprecipitation (Co-IP), which has proven to be extremely valuable in investigating these interactions. This article provides a thorough explanation of Co-IP and delves into its underlying principles, advantages, applications, and comparisons with other similar techniques. Creative Proteomics, a leading company in the field, boasts a successful history of developing innovative Co-IP protocols and supporting researchers in their quest for a better understanding of protein interactions.

What is Co-immunoprecipitation?

Co-immunoprecipitation, often abbreviated as Co-IP, represents a ubiquitous and extensively employed technique in the realm of molecular biology. Its fundamental purpose revolves around the highly advantageous ability to selectively isolate and subsequently identify protein complexes. This technique ingeniously harnesses the power of specific antibodies, aptly designed to zero in on a protein of interest, thereby facilitating the capture of its intricate web of interacting partners. This captivating procedure finds its utility in a myriad of experimental settings, encompassing diverse biological specimens ranging from cellular lysates to tissue extracts. By judiciously employing Co-IP, researchers are endowed with an invaluable tool to unravel the intricacies of protein interactions within the complex milieu of biological systems.

The Principle of Co-IP

The mind-bogglingly complex yet awesomely useful principle of Co-IP is rooted in the dynamic interplay between antibodies and antigens. Through the power of selective binding, antibodies cling onto specific target proteins and such is the basis of this technique. Co-IP employs highly specific antibodies that possess the rare ability to recognize and zero in on the molecule of interest amidst a heap of proteins in a mixture, such as a tissue extract or cell lysate. This one-of-a-kind binding action enables the separation and identification of the protein complex that houses the much-coveted target protein and its associated partners.

Antibody Selection and Preparation

The success of a Co-IP experiment heavily relies on the careful selection of appropriate antibodies. Specific antibodies against the target protein are chosen based on their affinity and specificity. Monoclonal antibodies, generated against purified proteins or synthetic peptides, are commonly used due to their high specificity and reproducibility.

To prepare the antibodies for Co-IP, they are typically conjugated to a solid support, such as protein A/G beads or agarose resin. The conjugation facilitates the binding of the antibodies to the protein complex and the subsequent separation of the complex from the rest of the sample.

Protein Complex Stabilization and Precipitation

Once the antibody is bound to the solid support, the protein mixture containing the target protein and its interacting partners is added. The specific antibody binds to the target protein, stabilizing the protein complex. The nonspecific proteins are washed away to eliminate background noise and non-relevant interactions.

To precipitate the protein complex, various techniques can be employed. Protein A/G beads or other similar matrices have a high affinity for the antibody and can be used to efficiently separate the complex from the solution. The precipitated complex is then collected by centrifugation or magnetic separation.

Elution and Analysis of the Protein Complex

To analyze the protein complex isolated through Co-IP, it needs to be eluted from the solid support. This can be achieved by different methods, such as altering the pH or using specific elution buffers. The eluted protein complex is then subjected to further analysis to characterize its composition and functionality.

Advantages of Co-IP

1. specificity and selectivity.

One of the key advantages of Co-IP is its ability to selectively capture and identify specific protein complexes. By using antibodies that specifically recognize the target protein of interest, Co-IP allows researchers to isolate and study the interactions of that protein with its binding partners. This specificity ensures that the identified protein complexes are relevant and minimizes false-positive results.

The specificity of Co-IP can be further enhanced by optimizing the experimental conditions, such as antibody selection, washing steps, and elution methods. These optimizations help reduce nonspecific background interactions, allowing for a more accurate assessment of the protein-protein interactions under investigation.

2. Preservation of Native Protein Complexes

Co-IP enables the isolation of protein complexes in their native state, maintaining their structural integrity and functional relevance. Unlike in vitro methods that may disrupt protein interactions, Co-IP allows researchers to capture and study protein complexes as they exist in cells or tissues.

Preserving the native conformation of protein complexes is crucial for understanding their biological functions, regulatory mechanisms, and roles in disease. Co-IP enables the investigation of protein interactions in a physiologically relevant context, providing insights into the dynamics and organization of protein complexes within cellular pathways.

3. Identification of Novel Interacting Partners

Co-IP is a powerful tool for discovering novel interacting partners of a target protein. By immunoprecipitating the target protein, Co-IP enables the co-capture of its binding partners, which can be subsequently identified using techniques such as mass spectrometry.

Mass spectrometry-based proteomics allows for the identification and quantification of proteins within the Co-IP complex. This approach provides a comprehensive view of the protein interaction network, unveiling previously unknown interacting partners and facilitating the exploration of new biological pathways and functional relationships.

4. Study of Endogenous Protein Complexes

Co-IP is particularly valuable for studying endogenous protein complexes in their natural cellular environment. By using antibodies against the target protein, Co-IP enables the capture of protein complexes as they occur naturally, without the need for overexpression or artificial modifications.

Studying endogenous protein complexes is essential for understanding their physiological roles, regulation, and involvement in disease processes. Co-IP allows for the investigation of protein interactions in a more biologically relevant context, providing insights into the functional significance of these complexes and their contributions to cellular functions.

5. Complementary Techniques

Co-IP can be combined with other techniques to complement and validate the results. For example, Co-IP can be followed by Western blotting to confirm the presence of specific proteins in the immunoprecipitated complex. Western blotting allows for the detection and quantification of individual proteins within the complex, providing additional validation and supporting the Co-IP results.

Furthermore, Co-IP can be combined with functional assays, such as enzyme activity assays or cellular localization studies, to gain further insights into the functional relevance of the protein complexes and their interactions.

Co-IP for Measuring Protein Interaction

Co-IP is widely used to measure protein interactions in diverse research areas, including signal transduction pathways, protein function studies, and disease-related investigations. By selectively capturing protein complexes, Co-IP provides valuable insights into the composition and dynamics of these interactions. It helps in deciphering the roles of specific proteins in complex biological processes and contributes to the understanding of disease mechanisms.

Learn more Application of Co-IP in protein interaction research

Co-immunoprecipitation Protocol

A successful Co-IP experiment requires careful optimization and adherence to a robust protocol. The protocol involves several critical steps, such as sample preparation, antibody selection, immunoprecipitation, washing steps to remove nonspecific interactions, and elution of the protein complex for subsequent analysis. Creative Proteomics offers comprehensive Co-IP protocols and technical support to ensure reliable and reproducible results for researchers.

To learn about Co-IP Protocol and how to optimize the protocol

Co-immunoprecipitation Mass Spectrometry

Co-IP combined with mass spectrometry (Co-IP-MS) has emerged as a powerful tool for protein interaction studies. It enables the identification and characterization of protein complexes by coupling Co-IP with high-throughput mass spectrometry analysis. This approach allows for the simultaneous identification of multiple proteins within the complex and the determination of their stoichiometry.

Learn more about the application of Co-IP Mass Spectrometry

Difference between Immunoprecipitation and Co-immunoprecipitation

Immunoprecipitation (IP) and Co-IP are closely related techniques, but they differ in their targeted objectives. IP involves the isolation of a single protein of interest, whereas Co-IP aims to capture the target protein along with its interacting partners. Co-IP provides a more comprehensive understanding of protein interactions and enables the study of protein complexes within a biological context.

Explore the key differences between immunoprecipitation and co-immunoprecipitation .

Co-immunoprecipitation and Pull-Down Assays

While Co-IP is a widely used technique for studying protein-protein interactions, it is important to acknowledge that alternative methods, such as pull-down assays, also exist. Pull-down assays involve the immobilization of a bait protein, typically with an affinity tag, onto a solid support, followed by the incubation with a protein mixture to capture interacting proteins. The bound proteins are subsequently eluted and analyzed.

The choice between Co-IP and pull-down assays depends on the specific research objectives and the nature of the protein interactions being studied. Co-IP is particularly advantageous when studying endogenous protein complexes and their interactions in a physiological context. On the other hand, pull-down assays offer advantages in cases where a specific bait protein is overexpressed or fused with a tag, allowing for the isolation of interacting partners under controlled conditions.

Learn more how to choose Co-IP and Pull-down .

Difference between Co-immunoprecipitation and Western Blot

While both Co-IP and Western blotting are commonly used techniques in protein analysis, they serve different purposes. Co-IP focuses on the isolation and identification of protein complexes, providing information about the interacting partners of a target protein. In contrast, Western blotting is a technique used to detect and quantify specific proteins within a sample. It is commonly employed to validate and confirm the presence of target proteins, including those identified through Co-IP experiments.

Western blotting involves the separation of proteins by gel electrophoresis, followed by their transfer onto a membrane. The membrane is then probed with specific antibodies to detect the target proteins of interest. Western blotting complements Co-IP by confirming the presence of the target protein and validating the results obtained from the Co-IP experiments.

Learn more information about Co-immunoprecipitation and Western Blot

Co-immunoprecipitation (Co-IP) is a powerful technique for studying protein-protein interactions and characterizing protein complexes. With its ability to selectively capture target proteins and their interacting partners, Co-IP provides valuable insights into the composition, dynamics, and functional relevance of protein complexes in various biological processes. The combination of Co-IP with techniques such as mass spectrometry or Western blotting enhances the characterization and validation of protein interactions.

What Can Creative Proteomics Do for You

As a leading company in the field of proteomics, Creative Proteomics is dedicated to supporting researchers in their protein interaction studies . With a wide range of Co-IP protocols and technical expertise, Creative Proteomics enables researchers to efficiently and effectively investigate protein-protein interactions, advancing our understanding of cellular processes, disease mechanisms, and potential therapeutic targets.

• Anke Schiedel Prediction and Targeting of Interaction Interfaces in G-Protein Coupled Receptor Oligomers Current Topics in Medicinal Chemistry 2018

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Co-Immunoprecipitation (co-IP) Protocol

Co-immunoprecipitation protocol.

A detailed guide of how Co-IP works, a step by step Co-IP protocol and some hints & tips for your Co-IP workflow!

Introduction

Co-immunoprecipitation (Co-IP) is a widely used technique to isolate target proteins and their binding partners from cell lysates. It involves incubating lysates with specific antibodies, extracting antibody/antigen complexes using protein A/G agarose beads, and analyzing the isolated proteins through Western blot.

Key Takeaways

• Co-IP is essential for isolating target proteins and their binding partners from cell lysates.
• It relies on antibody/antigen complex extraction using protein A/G agarose beads.
• solated proteins are subsequently analyzed by Western blot for further study.

What is Co-immunoprecipitation?

Co-immunoprecipitation is a popular tool that is carried out in order to isolate a target protein and it’s binding partners from whole cell lysates.

Jump to a section:

• - Co-IP Lysis Buffers
• - Co-IP Protocol
• - Hints & Tips
• - IP Crosslinking
• - Bradford Assay

How does Co-immunoprecipitation work?

The lysate is incubated with a protein-specific antibody. The antibody/antigen complex is pulled out of the sample using protein A/G-coupled agarose beads, which isolate your protein of interest. The protein of interest is then separated from the agarose beads by centrifugation and analysed by Western blot .

Below you will find everything you need to carry out co-immunoprecipation (Co-IP) including buffers and solutions, an optimized protocol and hint and tips for the perfect IP.

C o-immunoprecipation (Co-IP) Principle

Protein A & G Agarose Beads

Protein G Agarose Beads are an affinity matrix for the small-scale isolation of immunocomplexes from immunoprecipitations (IP assays). Protein G is covalently coupled to agarose beads, which enables it to bind to various antibodies, including those of the IgG class. This makes protein G agarose beads an ideal tool for immunoprecipitation experiments involving protein A and G antibodies.

Protein G agarose beads can be used in a variety of immunoprecipitation assays, including those involving protein A and G antibodies.

Some of the advantages of using protein G agarose beads include:

• They are easy to use and can be stored at room temperature.
• Protein G agarose beads have a high binding capacity for antibodies, which makes them ideal for small-scale immunoprecipitation experiments.
• Protein G agarose beads can be used with a variety of antibodies, including those of the IgG class.
• Protein G agarose beads are a versatile tool for immunoprecipitation experiments involving protein A and G antibodies.

Co-immunoprecipitation Lysis Buffers

Ip lysis buffer, ripa buffer, related resources, co-immunoprecipitation protocol steps.

The below protocol is a recommended protocol for the isolation of proteins from whole cell extracts.

Helpful Tips for Co-Immunoprecipitation

Lysate preparation.

The quality of the lysate you use for co-immunoprecipitation will determine the success of your assay. Using the right lysis buffer can greatly improve the quality of your lysate. An ideal lysis buffer should stabilize the native protein conformation, inhibit enzymatic activity, prevent denaturation and above all ensure maximum release of proteins from cells or tissues. Non-ionic detergents like NP-40 and Triton X-100 are less harsh when compared to ionic detergents such as SDS and sodium deoxycholate. The use of denaturing buffers such as radio-immunoprecipatation (RIPA) are ideal for proteins that are difficult to release such as nuclear proteins.

Detergent-free buffers can also be used if the target protein can be released from cells by physical disruption, such as mechanical homogenization or heat. Always remember that proteolysis, de-phosphorylation and denaturation can start as soon as cell lysis occurs. This can be slowed down by keeping the samples on ice or at 4°C at all times and through the addition of protease and phosphatase inhibitors to the lysis buffer.

Pre-clearing the Lysates

Pre-clearing the lysates with the beaded support before beginning co-immunoprecipitation helps to remove any potentially reactive components which may bind non-specifically to the bead components. This pre-clearing step can also be performed using a non-specific antibody of the same species of origin and isotype as the capture antibody. This process will remove anything that might bind non-specifically to the capture antibody during co-immunoprecipitation. The end result will be a lowering of background and an improved signal-to-noise ratio.

Antibody Choice

Choosing the right antibody for purification is critical as it can alter your protein yield. Polyclonal antibodies are ideal for the binding of your target protein as they can bind multiple epitopes on the target protein, and form tighter binding immune-complexes with higher retention rates. A combination of a polyclonal capture antibody and a monoclonal antibody for detection will guarantee maximum capture efficacy with high detection specificity. It should also be noted that the use of secondary antibodies which recognize the heavy and light-chain of the primary antibody for western blot detection of IP samples will always result in two bands (the heavy-chain at 50kDa and the light-chain at 25kDa).

Wash Buffer Choice

The wash buffer used for co-immunoprecipitation assays should reduce non-specific protein binding and maintain desired protein interactions. PBS and TBS are commonly used as wash buffers as they have physiological concentrations of salt and pH. Moderate adjustments to the salt concentration of wash buffers can be used to reduce background in some instances.

If non-specific interactions are detected using the above wash buffers the stringency may be increased by increasing the sodium chloride concentration. A low level of reducing agents (such as 1-2 mM DTT or β-mercaptoethanol) can help disrupt non-specific interactions.

Elution Buffer Choice

The strength and pH of your elution buffer ensures the correct elution of your target protein. If the immunoprecipitated sample is going to be further analysed by SDS-PAGE or Western Blot elution into running buffer would be ideal. Elution in a milder buffer (0.1 M glycine, pH 2.5) and neutralizing before loading to SDS-PAGE gel is also an option.

Immunoprecipitation Cross Linking

UV cross-linking and immunoprecipitation (CLIP) are two techniques utilized in molecular biology to study protein interactions with RNA or identify RNA modifications. Cross-linking allows for the proteins and RNAs to be covalently bonded, while immunoprecipitation uses antibodies to pull down the protein of interest. This method can be used with different types of RNA, including mRNAs, lncRNAs, and circRNAs.

There are several variations of CLIP, including:

• Crosslinking and Immunoprecipitation of RNA-Binding Proteins ( CLIP-seq ): This method is used to study the RNA binding sites of a protein by immunoprecipitating the protein and then sequencing the co-immunoprecipitated RNAs.
• Crosslinking and Immunoprecipitation of Modified RNAs ( cLIP-seq ): This method is used to study RNA modifications, such as m6A, by immunoprecipitating the modified RNA and then sequencing the co-immunoprecipitated RNAs.
• Crosslinking and Immunoprecipitation of Native RNAs ( CLIP-NAT ): This method is used to study the native RNA-protein interactions by immunoprecipitating the protein and then sequencing the co-immunoprecipitated RNAs.

CLIP is a powerful tool that can be used to study RNA-protein interactions and RNA modifications. This method has been used to identify new RNA binding proteins, to map the binding sites of RNA binding proteins, and to study RNA modifications.

Bradford Assay for Protein Determination

Written by Sean Mac Fhearraigh

Seán Mac Fhearraigh PhD is a co-founder of Assay Genie. Seán carried out his undergraduate degree in Genetics at Trinity College Dublin, followed by a PhD at University College Dublin. He carried out a post-doc at the Department of Genetics, University of Cambridge. Seán is now Chief Technical Officer at Assay Genie.

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Co-immunoprecipitation (Co-IP)

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Page contents 

Immunoprecipitation (ip) vs. co-immunoprecipitation (co-ip).

• Co-IP optimization strategies
• Evaluating a co-immunoprecipitated protein–protein interaction
• Recommended reading

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• Co-Immunoprecipitation (Co-IP) and Pull-Down
• Cell Lysis (Total Protein Extraction)
• Immunoprecipitation
• Dynabeads Co-Immunoprecipitation Kit
• Antibody Selection Tool

The topic of co-immunoprecipitation (co-IP) is best preceded by an  overview of immunoprecipitation (IP)  to help frame an understanding of the principles involved. The description of IP methodology here is brief.

Immunoprecipitation is one of the most widely used methods for antigen detection and purification. The principle of an IP is very straightforward: an antibody (monoclonal or polyclonal) against a specific target protein forms an immune complex with that target in a sample, such as a cell lysate. The immune complex is then captured, or precipitated, on a beaded support to which an antibody-binding protein is immobilized (such as Protein A or G), and any proteins not precipitated on the beads are washed away. Finally, the antigen (and antibody, if it is not covalently attached to the beads and/or when using denaturing buffers) is eluted from the support and analyzed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), often followed by western blot detection to verify the identity of the antigen.

Schematic summary of a standard immunoprecipitation assay.

Watch this video to learn more about immunoprecipitation

Co-immunoprecipitation (co-ip).

Co-immunoprecipitation is an extension of IP that is based on the potential of IP reactions to capture and purify the primary target (i.e., the antigen) as well as other macromolecules that are bound to the target by native interactions in the sample solution. Therefore, whether or not an experiment is called an IP or co-IP depends on whether the focus of the experiment is the primary target (antigen) or secondary targets (interacting proteins).

Schematic summary of a standard co-immunoprecipitation assay.

Watch this video to learn more about co-immunoprecipitation

• Overview of Protein-Protein Interaction Analysis
• Overview of the Immunoprecipitation (IP) Technique
• Overview of Western Blotting
• Overview of Affinity Purification
• Overview of Cell Lysis and Protein Extraction
• Protein Isolation and Purification Information
• Protein Purification and Isolation Support Center
• Protein Immunoprecipitation (IP), Co-Immunoprecipitation (Co-IP), and Pull-down Support
• Protein & Cell Analysis eLearning Course Series

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While the co-IP methodology is straightforward, performing a co-IP reaction and identifying physiological protein–protein interactions can be difficult because of the nature of the interaction, nonspecific binding to IP components and antibody contamination that may mask detection. The following sections describe each aspect of the co-IP approach that can be optimized to improve detection.

Complex binding

Because co-immunoprecipitation depends so much on protein–protein interactions in order to detect the bound proteins, the ability to maintain stable physiological interactions throughout the mechanical and chemical stresses of the incubation and washing steps is a critical factor when performing a co-IP reaction. Therefore, low-affinity or transient protein–protein interactions may not be detected by co-IP unless the interaction can be stabilized.

A key factor in maintaining complex formation throughout the steps required for co-IP is the lysis and wash buffers. Many protein interactions will remain intact after lysis using standard non-denaturing lysis buffers, as described in the Immunoprecipitation method in the Pierce Protein Methods library. Buffers with low ionic strength (i.e., <120mM NaCl) that contain non-ionic detergents (NP-40 and Triton X-100) are less likely to disrupt protein–protein interactions; however, empirical testing may be required to determine the best buffer formulation for a specific protein complex of interest.

Additionally, lysing cells by sonication or vortexing the lysate or bead-bound immune complexes during the wash steps should be avoided to prevent the disruption of the protein–protein interaction(s) of the target complex. And while centrifugation is a standard method to separate the precipitated complexes from the remaining lysate and during wash steps, the samples should be handled gently to prevent the loss of bound complex proteins.

An advanced technique to strengthen protein–protein interactions is by crosslinking the binding partners. Using this approach, all proteins within the active distance of the specific reagent in a cell lysate are covalently crosslinked, and the target protein can then be immunoprecipitated along with the other proteins in the complex without the risk of losing binding partners.

Agarose vs. magnetic beads for co-IP

Whereas agarose beads have long been a popular support for immunoprecipitation and other affinity-based purification procedures, magnetic beads are replacing them in IP/co-IP and other small-scale affinity procedures. Although agarose beads generally have a higher binding capacity due to their porous surface, magnetic beads offer advantages such as ease of use, lower nonspecific binding, and compatibility with automation.

High background from nonspecific interactions

With the myriad of proteins in cell lysates, it is inevitable that nonspecific binding to the IP antibody will occur, especially when using the batch method (a gentle, large-scale procedure) of immunoprecipitating the target protein. Additionally, because proteins that are normally separated into discrete cellular compartments are now mixed together, nonphysiological binding to the target complex is likely to occur, especially with abundant proteins such as actin. These nonspecific interactions are often broken by thoroughly washing the bead-bound immune complexes, but other strategies may be applied to optimize nonspecific binding, including:

• Changing the ionic strength of the IP buffer by titrating the salt concentration from 120 to 1000 mM.
• Decreasing the amount of primary antibody until the signal-to-noise ratio is maximized.
• Pre-clearing the lysates, as described in the Immunoprecipitation section .

Protein Preparation Handbook

Learn more about how to desalt, buffer exchange, concentrate, and/or remove contaminants from protein samples, immunoprecipitation and other protein purification and clean up methods using various Thermo Scientific protein biology tools in this 32-page handbook.

• Immunoprecipitation (IP), co-IP, and chromatin-IP
• Recombinant protein purification tags
• Dialyze protein samples securely using Slide-A-Lyzer dialysis cassettes and devices
• Rapidly desalt samples with high protein recovery using Zeba spin desalting columns and plates
• Efficiently extract specific contaminants using resins optimized for detergent or endotoxin removal
• Concentrate dilute protein samples quickly using Pierce protein concentrators

Antibody contamination

One of the most commonly encountered problems with both IP and co-IP approaches is interference from antibody bands during gel analysis. In those cases where several proteins may be co-precipitated with the target, the presence of the co-eluted antibody light and heavy chains (25- and 50-kDa bands in reducing SDS-PAGE gels, respectively) in the sample can obscure the results. The ideal situation would be to analyze the co-IP without contamination of the eluted antigen with antibody; with this potential interference eliminated, only the co-precipitated proteins would be present and detected on a gel.

Antibody contamination can be circumvented using methods described in the Overview of Immunoprecipitation Methods page , including crosslinking antibody to Protein A/G–coated beads or covalently binding antibody directly to treated beads. An added benefit of these approaches is the potential reuse of the antibody-coated beads. A key to preventing antibody contamination using these strategies is to elute the antigen under non-denaturing conditions; otherwise, the denatured antibody fragments will be eluted with the antigen.

Another direct coupling approach incorporates the binding association between streptavidin and biotin, in which the IP antibody is biotinylated and the beads are coated with streptavidin. The immune complexes are captured by the beads, and because biotin binds strongly to streptavidin, the antibody is not eluted from the beads when mild conditions are used to release the target antigen. A wide selection of affinity resins, magnetic beads and coated plates based on immobilized avidin, streptavidin or Thermo Scientific NeutrAvidin Protein facilitates this strategy.

By contrast, when popular fusion tags are incorporated into the primary target protein to be used in a co-IP experiment, pre-immobilized anti-fusion tag antibodies may be used for protein complex purification. For example, antibodies specific to the HA (YPYDVPDYA) or c-Myc tag (EQKLISEEDL) can be covalently immobilized to beaded agarose resin, enabling their use in IP or co-IP experiments involving HA- or c-Myc-tagged "bait" proteins.

Co-IP of active Rac1 with HA-tagged Pak1-PBD (p21 binding domain). Human 293 cells were transfected with HA-Pak1 protein binding domain (PBD) alone or co-transfected with constitutively activate Rac1 (Q61L). Anti-HA agarose slurry (6 µL) was incubated with 50 µL HA-tagged positive control lysate (Lane 1) or 500 µL cell lysate from Rac1 (Q61L) and HA-Pak-PBD co-transfected cells (Lane 2). HA-Pak1-PBD-transfected cells (Lane 3) or non-transfected cells (Lane 4). IP and co-IP reactions were performed at 4°C overnight. The western blot was first probed with anti-Rac1 antibody (A) and then reprobed with anti-HA antibody (B).

Description of the problems of traditional co-IP methods and solutions for optimization

• Protein Interaction Analysis
• Protein Purification Tools Poster
• Tech Tip #4: Batch and spin cup methods for affinity purification of proteins
• Pierce Classic Magnetic IP/Co-IP Kit
• Pierce Direct Magnetic IP/Co-IP Kit
• Pierce Crosslink Magnetic IP/Co-IP Kit

When a protein–protein interaction is detected, it is critical to confirm that the detection is a true physiological interaction as opposed to an artifactual interaction due to some aspect of the protocol. A summary of approaches to verify a protein–protein interaction follows.

Verify antibody specificity

The quality and specificity of antibodies range from those that weakly bind and are nonspecific to those that show high affinity and specificity for a single epitope. A critical part of confirming any detected protein–protein interaction is first confirming that the target protein can be immunoprecipitated from the sample, which is confirmed using well-characterized antibodies that are known to specifically bind to the target antigen. If data on the specificity of an antibody is not available, then cells that lack the target protein should be used with the IP antibody to show that nothing is precipitated using the antibody. Of course, when testing non-characterized antibodies, one should always include a control to show that the target protein can be precipitated from a stock of purified protein using the test antibody.

Whereas a number of antibody validation strategies can be used to verify the specificity of an antibody, antibody validation by immunoprecipitation followed by mass spectrometry analysis (IP-MS) can also identify previously known protein–protein interactions as well as suggest potential interacting partners that have not been previously described.

If a binding partner detected by co-IP truly interacts with a particular target protein, then multiple primary antibodies specific for the same epitope on that target protein should yield the same results. Antibodies that bind the same target protein but differ in epitope specificity may also co-IP the same proteins, although antibodies are known to prevent or disrupt the protein–protein interactions of protein complexes. Another indicator of a true protein–protein interaction, as opposed to an artifact, is that either protein can be co-immunoprecipitated when the IP antibody against the binding partner is used (i.e., protein A can be used to co-IP protein B, and protein B can be use to co-IP protein A).

Even high quality, monoclonal antibodies may bind to nonspecific proteins; therefore, performing a co-IP using a non-target antibody (often referred to as an 'irrelevant antibody') is critical to confirm that the immunoprecipitated protein complex is the specific complex that was sought. And because antibody specificity varies by subclass, it is recommended to use control antibodies that match the primary antibody as close as possible. 

Authenticate a functional interaction

Many protein–protein interactions are dependent upon the activation of one or more of the binding partners in a complex. Therefore, to test if a true interaction occurs, cells that express an inactive variant of one of the binding partners can be used for co-IP of the protein complex; if activation is required, then the complex will not be co-precipitated with the target antigen.

Confirm a physiological interaction

Cell lysis causes proteins that never interact to come into close association, and it is inevitable that some proteins will bind to each other. To test if a detected protein complex forms after cell lysis, Ohh et al. metabolically labeled all proteins in cells and then lysed the cells with a lysis buffer that included the purified, unlabeled form of the protein of interest. Because this unlabeled protein was unable to compete with the radiolabeled protein for complex formation recovered by co-IP, the researchers concluded that the complex represented a physiologically relevant interaction that formed prior to lysis.

• Invitrogen Antibody Validation
• Immunoprecipitation–Mass Spectrometry (IP-MS) Antibody Validation
• Application Note: Characterization of Antibody Specificity Using Immunoprecipitation and Mass Spectrometry
• Protein Mass Spectrometry Information
• Pierce MS-Compatible Magnetic IP Kit, protein A/G
• Pierce MS-Compatible Magnetic IP Kit, streptavidin
• Protein Mass Spectrometry Analysis
• Phizicky EM, Fields S (1995) Protein-protein interactions: Methods for detection and analysis. Microbiol Rev 59:94–123.
• Golemis E (2002) Protein-protein interactions: A molecular cloning manual. Cold Spring Harbor (NY): Cold Spring Harbor Laboratory Press. p ix, 682.
• Ohh M et al. (1998) The von hippel-lindau tumor suppressor protein is required for proper assembly of an extracellular fibronectin matrix. Mol Cell 1:959–68.

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Co-immunoprecipitation Protocol: Your Practical Guide To Co-IPs

last updated: February 2, 2022

Do you wonder if your favorite protein interacts with another protein? Do you wish that you could shine a spotlight on your protein to determine its binding partner? You can use co-immunoprecipitation (Co-IP) to find your protein’s partner.

This article will get you ready for your first Co-IP, provide a handy Co-IP protocol, and discuss Co-IP controls you should include. It will also touch on how you can use Co-IP with SPR applications to gain a detailed insight into how your protein interacts with others.

The Basics of Co-immunoprecipitation Experiments

In essence, Co-IP is just a more delicate version of a traditional immunoprecipitation, which can be thought of as a small-scale affinity purification. With both techniques, you use antigen/antibody interactions to isolate proteins. Your protein-of-interest is “pulled down” and out of solution by an antibody, which is, in turn, captured using beads.

However, a Co-IP requires greater care and more physiologically relevant conditions than traditional IP. When successful, Co-IP pulls down not only your protein-of-interest but also its interaction partners. Thus, Co-IPs are a great way to identify protein complexes. Furthermore, you can use Co-IPs to determine protein–protein interactions under varying conditions. You can then go on to study binding kinetics using  SPR techniques .

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A Simple Co-immunoprecipitation Protocol

Co-IP protocols are very similar to traditional IP protocols, with the difference that Co-IPs require more gentle assay conditions to maintain the interaction with binding partners. The general steps are as follows (Figure 1).

1. Lyse your Cells

Here you gently break open your cells to make your protein accessible to the antibody. The method of lysis is important in Co-IPs. Non-detergent, low-salt lysis buffers are a popular choice for Co-IP of soluble proteins. This kind of lysis is least likely to disturb any protein interactions.

For less soluble protein complexes, however, lysis buffers may need to contain non-ionic detergents such as  NP-40  or Triton X-100. Set time aside to test a number of lysis conditions to find the optimal lysis solution. Some things in science, like the best lysis buffer, just need to be determined empirically.

Don’t forget to use protease inhibitors when lysing your cells to avoid your protein getting eaten up by proteases, which also get released during cell lysis.

The amount matters as well: the more cell lysate you use, the more of your protein you can pull down. The exact amount you should use will vary depending on how abundant your protein is, whether or not you are pulling down overexpressed protein or endogenous protein, and how much interaction it has with binding partners. A good starting point is to use at least 1 mg of total protein, but you may need to optimize this.

Protein concentration matters too . Co-IPs are usually performed in a microcentrifuge tube so you need to ensure your cell lysate is concentrated enough to fit the desired amount of total protein plus other reagents with room to move around.

2. Add Your Antibody

Now it’s time to add an antibody specific to your protein-of-interest. It can be difficult to choose the best antibody for a Co-IP but many antibody manufacturers provide helpful information in their product selection guides.

Here are a few tips for selecting a good antibody for Co-IP.

• Use an antibody that has been previously shown to precipitate the protein (e.g., from the literature).
• Choose a polyclonal antibody rather than a monoclonal antibody, where possible. Polycloncal antibodies will bind your protein in various locations, meaning a greater chance of successful Co-IP, and minimizes potential problems such as if your antibody recognizes the site that binds with the partner you are trying to find.
• If polyclonal antibodies are not an option, choose an antibody that recognizes an epitope on your protein-of-interest that is not masked by any other protein–protein interactions.
• Ensure the antibody binds to the native protein; some antibodies recognize only denatured proteins.

3. Add the Protein A/G Beads

In general, beads are used to physically pull down and purify the antibody–protein complex from the rest of your mixture.

There are two main types of beads you can use: beads coated in protein A or beads coated in protein G. Protein A and G are specialized bacterial proteins that recognize and bind to antibodies. You can also get beads coated in both protein A and protein G. The species and type of your antibody will determine which of these two types of beads you should use. To determine which one is correct for you, consult your antibody’s data sheet . AbCam also has a useful table showing the binding profiles for protein A, protein G, protein A/G, and protein L beads . 

Traditionally, immunoprecipitation beads were agarose spheres (50–150 µM in diameter) that allowed purification of the antibody–protein complexes via centrifugation. Centrifugation methods include the use of bead slurries or spin columns.

Spin columns are generally fast and come as part of a kit with highly optimized protocols. However, they may be an expensive option.

Bead slurries, on the other hand, are great for larger complexes that require very gentle handling, without high centrifuge speeds.

More modern techniques use magnetic beads of 1–4 µM in diameter instead of agarose beads. Magnetic beads allow you to isolate your antibody–protein complex by magnets! Magnetic beads may be better for high-weight protein complexes and processing low-volume samples to avoid excessive sample loss during centrifugation. Overall though, agarose beads often have a superior yield since they have a large surface area on which multiple A/G proteins are linked.

4. Incubate

This is the time for the bead–antibody–protein interactions to occur. Incubation times vary from 30 minutes to overnight depending on which bead you choose. Often, the incubation step includes gentle agitation, such as on a rocker plate or tube rotator at low speed because agitation enhances the opportunity for interactions. 

After incubation, you will need to collect the antibody–protein complexes. Depending on the method you are using, you will either use magnets or centrifugation to separate the complexes from the rest of the cellular proteins.

6. Wash the Beads

Now that your proteins-of-interest are hopefully tethered to your bead via your antibody it is time to get rid of everything else in your lysate.

Do this by washing your antibody–bead complex a few times to clear away cellular matter not specifically bound by your antibody. Washing also helps reduce non-specific binding of “sticky” cellular components to the beads.

Generally, you wash using cold lysis buffer or cold PBS. You may need to optimize this step to find the right level of stringency that doesn’t disrupt the interacting proteins.

Expert Tip:  Save all of your used wash buffer during this step. The wash buffers can show you if your protein-of-interest and any partner proteins were depleted from the lysate. This is critical knowledge for troubleshooting your Co-IP!

7. Elute your Protein(s)

Most SDS-PAGE loading buffers reduce and denature proteins. Therefore, it is often an effective way to separate the proteins from the beads. If you plan on using SDS-PAGE as your detection method (which most of us do), then you can add loading directly to the beads to harvest your protein(s).

If you want to detect your proteins using native PAGE protocol, or you plan on doing enzyme assays with your isolated proteins, then you may need to use something other than SDS-PAGE loading buffer to elute.

A frequently used non-denaturing elution buffer is 0.1M glycine at a low pH (around 2.5–3). However, some proteins will still denature or lose enzymatic activity under these conditions, and some proteins may not dissociate with this treatment. So chalk this up to yet another step to experiment with.

8. Detect your Protein(s)

By now you have isolated your protein(s) and are ready to go. (Yay!) Now the exciting part…taking a peek at your isolated protein-of-interest and its dance partner(s). How you want to do this—SDS-PAGE, mass spectrometry, enzymatic assays, etc.—is up to you.

Expert tip: Remember that the antibody you used to purify your protein is also in the eluent along with your protein-of-interest. And this antibody contamination can be a problem. A western blot may detect the antibodies in your eluent, thus masking the detection of your proteins-of-interest if they are close in molecular weight. (The molecular weight of an antibody’s heavy and light chain are about 50 and 25 kD, respectively.)

You might be able to overcome this issue by 1) using beads covalently bound to the antibody, or 2) ensuring that the secondary antibody chosen for your western blot recognizes a different species than that of your Co-IP antibody.

9. Study the Interaction With Surface Plasmon Resonance Applications

A positive finding in a Co-IP is great news, but you should temper your excitement. Many scientists are led astray by sticky proteins. You should do additional studies to confirm any potential interactions and to ensure that they are specific. For example, you can use mutational analysis to map binding sites. Alternatively, for more specific information about an interaction, consider using an  SPR application .

SPR applications not only confirm interactions between binding partners, but also provide quantitative measurements of the affinity, thermodynamics, and kinetics of these interactions in real-time.

An SPR biosensor  that allows measurement of the interaction at different temperatures facilitates a thermodynamic analysis of the interaction of interest. With SPR, you can confidently compare affinities between your protein-of-interest and different binding partners.

A major practical advantage of SPR is that you don’t need to label your protein-of-interest (so no more tag-cloning or radioactivity), saving you time and potential hassle with tricky cloning strategies.

If you want to detect and unravel interactions between immune receptors and their ligands, you might want to consider trying a  biosensor assay . As explained in a  previous Bitesize Bio article , biosensor assays allow you to measure the rate of association and dissociation of ligands to their specific antigen receptors. These assays follow SPR principles. Owing to their advantages and vast capabilities, SPR and its related applications have become the gold standard for kinetic and affinity determination, and SPR principles underline most color-based  biosensor chip  applications.

Controls for Co-immunoprecipitation

Controls are critical to know that your experiment worked and to troubleshoot if things don’t quite work out. There are several different controls you can use for Co-IPs (Figure 2).

Negative Controls

These controls show that any interaction you see in your experiment is valid. Use beads coated with a non-specific antibody incubated in your lysate and treated the same as your samples. It’s best to choose one from the same species as the antibody in your experiment to better mimic any non-specific binding.

You can also use just the agarose beads without any antibody or vice-versa, to determine the exact cause of any non-specific binding.

If you see your protein-of-interest or potential binding partners in this control, you’ve got non-specific binding and you need to review your protocol.

Positive Controls 

Check for the presence of your protein/potential binding partner in the total lysate (without any beads/ Co-IP performed). This ensures that if you aren’t seeing your protein of interest it isn’t due to some issue with the lysate or detection method.

If your protein is in the total lysate, but not in the IP, you will need to troubleshoot each step to ensure that the IP is working.

Purified recombinant protein can be used to ensure that your detection method is working—this could be for either the protein being pulled down or the potential interacting protein.

More Expert Tips for Co-immunoprecipitation

Of course, within these steps, there are many variables that can make your Co-IP successful, or leave you with a sad face at the end. And there are myriad companies, products, and kits from which you can choose. So prepare yourself for some trial and error!

Before we finish up, here are a few more expert tips to delicately sneak a peek at your protein interactions without disturbing them too much.

Pre-clear Your Lysate

“Pre-clear” your lysate by adding protein A/G beads to the lysate before introducing your antibody. Next, spin down the beads and discard them to pull out any cellular components that stick to the beads unspecifically. Then add your antibody/beads and proceed. 

Pre-clearing can help to reduce non-specific binding and reduce potential background. However, if you plan to use western blotting to detect your proteins at the end of your Co-IP, pre-clearing is probably not necessary unless you know that a contaminating protein is likely to interfere with your protein/band of interest.

Plan the Order Carefully

Introduction of the bead to the antibody is an important variable that can affect yield. Typically, the antibody is immobilized to the bead by incubating them together before introducing the lysate. However, if the protein-of-interest is present in low concentrations, or if the antibody has a weak affinity for your protein-of-interest, you may need to incubate the protein and antibody first, then add the beads.

Go Gentle, Go Slow

Your goal is to maintain the protein–protein interactions. Your antibody–protein complexes will be physically pulled out of solution by protein A or G conjugated beads. And while the tether from the bead to the antibody is highly specific, it is tenuous.

Therefore, treat your cell lysate with velvet gloves. This includes choosing the right lysis method, keeping your samples chilled, and avoiding overhandling.

Vortexing, mixing with small pipette tips, or using high spins in the centrifuge can all break apart your delicate chain of protein–antibody–bead, and fling your proteins back into solution.

To keep the proteins in your lysate from getting degraded or altered, perform your Co-IPs at 4°C. A cold room is perfect, especially if you can have all your equipment (e.g., rocker/centrifuge) set up in easy reach.

Co-immunopreciptation Summarized

Co-IPs are a useful and relatively straightforward tool to identify protein–protein interactions. The Co-IP protocol and tips provided in this article can help ensure you get great and meaningful results.

Do you have any extra tips for performing Co-IPs? Let us know in the comments.

Originally published in 2014. Updated and republished in January 2022.

Ashleigh gained a Bachelor of Science from UC San Diego .

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Co-immunoprecipitation(Co-IP) is a technique to identify protein-protein interactions.

Co-immunoprecipitation (Co-IP) is a technique to identify protein-protein interactions. It involves isolating a protein complex from a solution using an antibody specific for one of the proteins in the complex. The specific antibody binds to the protein of interest and pulls down other proteins that are part of the complex. This technique is commonly used to study the interactions between different proteins in cells and tissues. Co-IP is crucial for understanding the formation of protein complexes, identifying unknown members of protein complexes, and studying the physiological interactions between proteins. It is widely used in molecular biology, cell biology, and biochemistry research to investigate protein interactions.

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Co-immunoprecipitation is a technique widely utilized to isolate protein complexes and study protein-protein interactions. Ubiquitinated proteins could be identified by combining co-immunoprecipitation with SDS-PAGE followed by immunoblotting. In this chapter, we use Herpes Simplex Virus 1 immediate-early protein ICP0-mediated polyubiquitination of p50 as an example to describe the method to identify a ubiquitinated adaptor protein by a viral E3 ligase by co-immunoprecipitation.

Application of Proteomics Technology Based on LC-MS Combined with Western Blotting and Co-IP in Antiviral Innate Immunity

As an interferon-stimulating factor protein, STING plays a role in the response and downstream liaison in antiviral natural immunity. Upon viral invasion, the immediate response of STING protein leads to a series of changes in downstream proteins,

As an interferon-stimulating factor protein, STING plays a role in the response and downstream liaison in antiviral natural immunity. Upon viral invasion, the immediate response of STING protein leads to a series of changes in downstream proteins, which ultimately leads to an antiviral immune response in the form of proinflammatory cytokines and type I interferons, thus triggering an innate immune response, an adaptive immune response in vivo, and long-term protection of the host. In the field of antiviral natural immunity, it is particularly important to rigorously and sequentially probe the dynamic changes in the antiviral natural immunity connector protein STING caused by the entire anti-inflammatory and anti-pathway mechanism and the differences in upstream and downstream proteins. Traditionally, proteomics technology has been validated by detecting proteins in a 2D platform, for which it is difficult to sensitively identify changes in the nature and abundance of target proteins. With the development of mass spectrometry (MS) technology, MS-based proteomics has made important contributions to characterizing the dynamic changes in the natural immune proteome induced by viral infections. MS analytical techniques have several advantages, such as high throughput, rapidity, sensitivity, accuracy, and automation. The most common techniques for detecting complex proteomes are liquid chromatography (LC) and mass spectrometry (MS). LC-MS (Liquid Chromatography-Mass Spectrometry), which combines the physical separation capability of LC and the mass analysis capability of MS, is a powerful technique mainly used for analyzing the proteome of cells, tissues, and body fluids. To explore the combination of traditional proteomics techniques such as Western blotting, Co-IP (co-Immunoprecipitation), and the latest LC-MS methods to probe the anti-inflammatory pathway and the differential changes in upstream and downstream proteins induced by the antiviral natural immune junction protein STING.

A Guideline Strategy for Identifying Genes/Proteins Regulating Antiviral Innate Immunity

Antiviral innate immunity is a complicated system initiated by the induction of type I interferon (IFN-I) and downstream interferon-stimulated genes (ISGs) and is finely regulated by numerous positive and negative factors at different signaling

Antiviral innate immunity is a complicated system initiated by the induction of type I interferon (IFN-I) and downstream interferon-stimulated genes (ISGs) and is finely regulated by numerous positive and negative factors at different signaling adaptors. During this process, posttranslational modifications, especially ubiquitination, are the most common regulatory strategy used by the host to switch the antiviral innate signaling pathway and are mainly controlled by E3 ubiquitin ligases from different protein families. A comprehensive understanding of the regulatory mechanisms and a novel discovery of regulatory factors involved in the IFN-I signaling pathway are important for researchers to identify novel therapeutic targets against viral infectious diseases based on innate immunotherapy. In this section, we use the E3 ubiquitin ligase as an example to guide the identification of a protein belonging to the RING Finger (RNF) family that regulates the RIG-I-mediated IFN-I pathway through ubiquitination.

Protein–Protein Interactions: Co-immunoprecipitation

Proteins often do not function as single substances but rather as team players in a dynamic network. Growing evidences show that protein–protein interactions are crucial in many biological processes in living cells. Genetic (such as yeast two hybrid,

Proteins often do not function as single substances but rather as team players in a dynamic network. Growing evidences show that protein–protein interactions are crucial in many biological processes in living cells. Genetic (such as yeast two hybrid, Y2H) and biochemical (such as co-immunoprecipitation, co-IP) methods are the commonly used methods to identify the interacting proteins. Immunoprecipitation (IP), a method using a target protein-specific antibody in conjunction with Protein A/G affinity beads, is a powerful tool to identify the molecules interacting with specific proteins. Therefore, co-IP is considered to be one of the standard methods to identify and/or confirm the occurrence of the protein–protein interaction events in vivo. The co-IP experiments can identify proteins via direct or indirect interactions or in a protein complex. Here, we use two different co-Ip protocols as an example to describe the principle, procedure, and experimental problems of co-IP. First, we show the interaction of two Agrobacterium type VI secretion system (T6SS) sheath components TssB and TssC 41 , and secondly, we show the protocol we used for determining the interaction of an epitope-tagged T6SS effector, Tde1 expressed in Agrobacterium with endogenously expressing adaptor/chaperone protein Tap1.

Co-immunoprecipitation Assays to Detect Protein–Protein Interactions

Proteins usually do not function as monomers but rather perform their functions by interacting with themselves or other proteins. Co-immunoprecipitation is an essential assay for detecting protein interactions in vivo. In this chapter, we describe

Proteins usually do not function as monomers but rather perform their functions by interacting with themselves or other proteins. Co-immunoprecipitation is an essential assay for detecting protein interactions in vivo. In this chapter, we describe how to use co-immunoprecipitation to detect protein interactions in Arabidopsis protoplasts, seedlings, and Nicotiana benthamiana leaves. When using co-immunoprecipitation assays to detect protein interactions, it is necessary to pay attention to the design of the experimental and control groups.

Co-immunoprecipitation-Based Isolation of Double-Stranded RNA-Associated Protein Complexes in Nicotiana benthamiana

Double-stranded RNA (dsRNA) is associated with most viral infections, and is generated in host cells during viral replication. Viral RNA replication occurs within the viral factories called the viral replication complexes (VRCs). In addition to viral

Double-stranded RNA (dsRNA) is associated with most viral infections, and is generated in host cells during viral replication. Viral RNA replication occurs within the viral factories called the viral replication complexes (VRCs). In addition to viral genome, viral-derived dsRNA and replicase, the VRCs composition remains largely unexplored. The dsRNA binding domain of the B2 protein from Flock house virus has been reported to be used for detecting viral-derived long dsRNA in plants efficiently. Nicotiana benthamiana is widely used as a model plant for plant-microbe interactions owing to its susceptibility to diverse plant diseases, especially viral diseases. Here, we describe the use of Nicotiana benthamiana stably expressing GFP-tagged dsRNA binding protein (B2: GFP) to pull down dsRNA and associated host and viral proteins from turnip mosaic virus-infected plants. The obtained protein complexes are compatible with functional assays, Western blotting, and mass spectrometry. This system provides a valuable and robust tool to study VRC proteome in N. benthamiana upon plant viral infections.

In-vivo Cross-linking of Biotinylated Peptide Ligands to Cell Surface Receptors

In-vivo cross-linking of biotinylated peptides is a technique to analyze the interaction of small proteins or peptide ligands with their corresponding receptors. Here, we describe an in-vivo method in which leaves of living plants, transiently

In-vivo cross-linking of biotinylated peptides is a technique to analyze the interaction of small proteins or peptide ligands with their corresponding receptors. Here, we describe an in-vivo method in which leaves of living plants, transiently expressing receptor proteins, are infiltrated with biotinylated peptides. The interaction between ligand and receptor is irreversibly fixed by the infiltration of a cross-linking agent. Subsequently, co-immunoprecipitation is used to pull down the receptor-ligand pair. After western blotting, the biotin tag of the ligand peptide cross-linked to the receptor can be detected by streptavidin-AP conjugate on the membrane.

Rapid Identification of Peptide-Receptor-Coreceptor Complexes in Protoplasts

Secreted signaling peptides, also called peptide hormones, play crucial roles in regulating plant growth, development, and immunity. Plant peptide hormones are perceived by plasma membrane-localized receptor-like kinases (RLKs) or receptor-like

Secreted signaling peptides, also called peptide hormones, play crucial roles in regulating plant growth, development, and immunity. Plant peptide hormones are perceived by plasma membrane-localized receptor-like kinases (RLKs) or receptor-like proteins (RLPs) that harbor specific extracellular domains to bind and recognize the corresponding peptide ligands. Binding of a peptide ligand to its receptor usually induces the hetero-dimerization of the cognate receptor and a coreceptor, followed by the phosphorylation and activation of the receptor complex to transduce downstream signaling. Therefore, matching peptide ligands with their respective receptors/coreceptors is crucial for elucidating peptide hormone signaling pathways. In this chapter, using the RGF7 peptide-RGI4/RGI5 receptor-BAK1 coreceptor complex as an example, we describe a rapid method to identify the peptide ligand-receptor-coreceptor complexes via co-immunoprecipitation assays using recombinant proteins transiently expressed in Arabidopsis protoplasts.

Understanding Pseudophosphatase Function Through Biochemical Interactions

Pseudophosphatases have been solidified as important signaling molecules that regulate signal transduction cascades. However, their mechanisms of action remain enigmatic. Reflecting this mystery, the prototypical pseudophosphatase STYX

Pseudophosphatases have been solidified as important signaling molecules that regulate signal transduction cascades. However, their mechanisms of action remain enigmatic. Reflecting this mystery, the prototypical pseudophosphatase STYX (phospho-serine-threonine/tyrosine-binding protein) was named with allusion to the river of the dead in Greek mythology to emphasize that these molecules are “dead” phosphatases. Although proteins with STYX domains do not catalyze dephosphorylation, this does not preclude their having other functions, including as integral elements of signaling networks. Thus, understanding their roles may mark them as potential novel drug targets. This chapter outlines common strategies used to characterize the functions of pseudophosphatases, using as an example MK-STYX [MAPK (mitogen-activated protein kinase) phospho-serine-threonine/tyrosine-binding], which has been linked to tumorigenesis, hepatocellular carcinoma, glioblastoma, apoptosis, and neuronal differentiation. We start with the importance of “restoring” (when possible) phosphatase activity in a pseudophosphatase, so the active mutant may be used as a comparison control throughout immunoprecipitation and mass spectrometry analyses. To this end, we provide protocols for site-directed mutagenesis, mammalian cell transfection, co-immunoprecipitation, phosphatase activity assays, and immunoblotting that we have used to investigate MK-STYX and the active mutant MK-STYX active . We also highlight the importance of utilizing RNA interference (RNAi) “knockdown” technology to determine a cellular phenotype in various cell lines. Therefore, we outline our protocols for introducing short hairpin RNA (shRNA) expression plasmids into mammalian cells and quantifying knockdown of gene expression with real-time quantitative PCR (qPCR). We also provide a bioinformatic approach to investigating MK-STYX and MK-STYX (active mutant) . These bioinformatic approaches can stand alone experimentally but also complement and enhance “wet” bench approaches such as binding assays and/or activity assays. A combination of cellular, molecular, biochemical, proteomic, and bioinformatic techniques has been a powerful tool in identifying novel functions of MK-STYX. Likewise, the information provided here should be a helpful guide to elucidating the functions of other pseudophosphatases.

Analysis of Lipid GPCR Molecular Interactions by Proximity Labeling

G-protein-coupled receptors (GPCRs) are hepta-helical transmembrane proteins that mediate various intracellular signaling events in response to their specific ligands including many lipid mediators. Although analyses of GPCR molecular interactions

G-protein-coupled receptors (GPCRs) are hepta-helical transmembrane proteins that mediate various intracellular signaling events in response to their specific ligands including many lipid mediators. Although analyses of GPCR molecular interactions are pivotal to understanding diverse intracellular signaling events, affinity purification of interacting proteins by a conventional co-immunoprecipitation method is challenging due to the hydrophobic nature of GPCRs and their dynamic molecular interactions. Proximity labeling catalyzed by a TurboID system is a powerful technique for defining the molecular interactions of target proteins in living cells. TurboID and miniTurbo (a modified version of TurboID) are engineered biotin ligases that biotinylate neighboring proteins in a promiscuous manner. When fused with a target protein and expressed in living cells, TurboID or miniTurbo mediates the biotin labeling of the proteins with close proximity to the target protein, allowing efficient purification of the biotinylated proteins followed by a shot-gun proteomic analysis. In this chapter, we describe a step-by-step protocol for the labeling of GPCR neighboring proteins by TurboID or miniTurbo, purification of the biotin-labeled proteins, and subsequent sample preparation for proteomic analysis. We utilized S1PR1 as a model GPCR, a receptor for a bioactive lipid molecule sphingosine 1-phosphate (S1P) that plays various roles in physiological and pathological conditions. This analysis pipeline enables the mapping of interacting proteins of lipid GPCRs in living cells.

Coimmunoprecipitation and Proteomic Analyses

Defining protein-protein interaction networks is a major goal of proteomics. Here, we present a protocol for coimmunoprecipitation, a technique suitable for the isolation of whole protein complexes in vivo and their subsequent identification by

Defining protein-protein interaction networks is a major goal of proteomics. Here, we present a protocol for coimmunoprecipitation, a technique suitable for the isolation of whole protein complexes in vivo and their subsequent identification by either immunoblotting or mass spectrometric sequencing combined to database search.

Protein–Protein Interactions in Abiotic Stress Signaling: An Overview of Biochemical and Biophysical Methods of Characterization

The identification and characterization of bona fide abiotic stress signaling proteins can occur at different levels of the complete in vivo signaling cascade or network. Knowledge of a particular abiotic stress signaling protein could theoretically

The identification and characterization of bona fide abiotic stress signaling proteins can occur at different levels of the complete in vivo signaling cascade or network. Knowledge of a particular abiotic stress signaling protein could theoretically lead to the characterization of complete networks through the analysis of unknown proteins that interact with the previously known protein. Such signaling proteins of interest can indeed be experimentally used as bait proteins to catch interacting prey proteins, provided that the association of bait proteins and prey proteins should yield a biochemical or biophysical signal that can be detected. To this end, several biochemical and biophysical techniques are available to provide experimental evidence for specific protein–protein interactions, such as co-immunoprecipitation, bimolecular fluorescence complementation, tandem affinity purification coupled to mass spectrometry, yeast two hybrid, protein microarrays, Förster resonance energy transfer, or fluorescence correlation spectroscopy. This array of methods can be implemented to establish the biochemical reality of putative protein–protein interactions between two proteins of interest or to identify previously unknown partners related to an initially known protein of interest. The ultimate validity of these methods however depends on the in vitro/in vivo nature of the approach and on the heterologous/homologous context of the analysis. This chapter will review the application and success of some classical methods of protein–protein interaction analysis in the field of plant abiotic stress signaling.

Wnt-Frizzled Interactions in Xenopus

The Wnt signaling cascades are regulatory modules which are involved in embryonic patterning, cell differentiation, morphogenesis, and diseases (1, 2). The Wnt pathways are activated when secreted Wnt ligands interact with 7-trans-membrane receptors

The Wnt signaling cascades are regulatory modules which are involved in embryonic patterning, cell differentiation, morphogenesis, and diseases (1, 2). The Wnt pathways are activated when secreted Wnt ligands interact with 7-trans-membrane receptors of the Frizzled (Fz) family. Specific readouts are determined by the ligand/receptor combinations and the cellular context. Here we describe two methods for the analysis of Wnt/Frizzled interactions in Xenopus embryos. Physical interaction of ligand and receptor are demonstrated by co-immunoprecipitation assays. The activation of Wnt targets in Xenopus animal cap tissue provides a versatile test system for activating and inhibitory components of the Wnt/ β -catenin pathway.

Study of GPCR Homo- and Heteroreceptor Complexes in Specific Neuronal Cell Populations Using the In Situ Proximity Ligation Assay

Membrane receptor, for example, G-protein-coupled receptors (GPCRs) , operates via coordinated changes between the receptor expression, their modifications , and interactions between each other. Perturbation in specific heteroreceptor complexes and/or

Membrane receptor, for example, G-protein-coupled receptors (GPCRs) , operates via coordinated changes between the receptor expression, their modifications , and interactions between each other. Perturbation in specific heteroreceptor complexes and/or their balance/equilibrium with other heteroreceptor complexes and corresponding homoreceptor complexes is considered to have a role in pathogenic mechanisms, including drug addiction, depression, Parkinson ’ s disease , and schizophrenia. To understand the associations of GPCRs and to unravel the global picture of their receptor – receptor interactions in the brain, different experimental detection techniques for receptor – receptor interactions have been established (e.g., co-immunoprecipitation - based approach). However, they have been criticized for not reflecting the cellular situation or the dynamic nature of receptor – receptor interactions. Therefore, the detection and visualization of GPCR homo- and heteroreceptor complexes in the brain remained largely unknown until recent years, when a well-characterized in situ proximity ligation assay (in situ PLA) was adapted to validate the receptor complexes in their native environment. The in situ PLA protocol presented here can be used to visualize GPCR receptor – receptor interactions in cells and tissues in a highly sensitive and specific manner. We have developed a combined method using immunohistochemistry and PLA, particularly aimed to monitor interactions between GPCRs in specific neuronal cell populations. This allows the analysis of homo - and heteroreceptor complexes at a cellular and subcellular level. The method has the advantage that it can be used in clinical specimens, providing localized, quantifiable homo and heteroreceptor complexes detected in single cells. We compare the advantages and limitations of the methods, underlining recent progress and the growing importance of these techniques in basic research. We discuss also their potential as tools for drug development and diagnostics.

Computational Prediction of Protein-Protein Interactions

One of the current goals of proteomics is to map the protein interaction networks of a large number of model organisms (1). Protein-protein interaction information allows the function of a protein to be defined by its position in a complex web of

One of the current goals of proteomics is to map the protein interaction networks of a large number of model organisms (1). Protein-protein interaction information allows the function of a protein to be defined by its position in a complex web of interacting proteins. Access to such information will greatly aid biological research and poten- tially make the discovery of novel drug targets much easier. Previously, the detection of protein-protein interactions was limited to labor-intensive experimental techniques such as co-immunoprecipitation or affinity chromatography. High-throughput experi- mental techniques such as yeast two-hybrid and mass spectrometry have now also become available for large-scale detection of protein interactions. These methods, how- ever, may not be generally applicable to all proteins in all organisms, and may also be prone to systematic error. Recently, a number of complementary computational approaches have been developed for the large-scale prediction of protein-protein inter- actions based on protein sequence, structure, and evolutionary relationships in com- plete genomes.

Chemical Cross-Linking for Protein–Protein Interaction Studies

Most proteins function through protein complex assemblies. Defining and mapping protein complex networks are crucial elements in the fundamental understanding of biological processes. The ability to measure protein–protein interactions in biological

Most proteins function through protein complex assemblies. Defining and mapping protein complex networks are crucial elements in the fundamental understanding of biological processes. The ability to measure protein–protein interactions in biological systems has undergone significant advances in the past decade due to emergence and growth of numerous new molecular biology and mass spectrometry technologies. Chemical cross-linking, along with yeast two-hybrid, fluorescence resonant energy transfer (FRET), and co-immunoprecipitation have become important tools for detection and characterization of protein–protein interactions. Individual protein members in a noncovalent complex assembly remain in close proximity which is within the reach of the two reactive groups of a cross-linker. Thus cross-linking reactions have potential for linking two interacting proteins which exist in close proximity. In general, chemical cross-linking experiments are carried out by first linking the interacting proteins through covalent bonds followed by a series of well-established protocols–SDS-PAGE, in-gel digestion, and shotgun LC/ MS/MS for identification of the cross-linked proteins. These approaches have been employed for both mapping topology of protein complex in vitro and determining the protein interaction partners in vivo .

Proteins participate in many processes of the organism and are very important for maintaining the health of the organism. However, proteins cannot function independently in the body. They must interact with proteins, DNA, RNA, and other substances to

Proteins participate in many processes of the organism and are very important for maintaining the health of the organism. However, proteins cannot function independently in the body. They must interact with proteins, DNA, RNA, and other substances to perform biological functions and maintain the body’s health. At present, there are many experimental methods and software tools that can detect and predict the interaction between proteins and other substances. There are also many databases that record the interaction between proteins and other substances. This article mainly describes protein–protein, protein–DNA, and protein–RNA interactions in detail by introducing some commonly used experimental methods, the software tools produced with the accumulation of experimental data and the rapid development of machine learning, and the related databases that record the relationship between proteins and some substances. By this review, we hope that through the analysis and summary of various aspects, it will be convenient for researchers to conduct further research on protein interactions.

Related Techniques

Co-immunoprecipitation occurring with Yeast Two-hybrid Assay

Co-immunoprecipitation for Assessing Protein–Protein Interactions in Agrobacterium- Mediated Transient Expression System in Nicotiana benthamiana

The characterization of protein–protein interactions (PPI) often provides functional information about a target protein. Yeast-two-hybrid (Y2H) and luminescence/fluorescence-based detections, therefore, have been widely utilized for assessing PPI. In

The characterization of protein–protein interactions (PPI) often provides functional information about a target protein. Yeast-two-hybrid (Y2H) and luminescence/fluorescence-based detections, therefore, have been widely utilized for assessing PPI. In addition, a co-immunoprecipitation (co-IP) method has also been adopted with transient protein expression in Nicotiana benthamiana ( N. benthamiana ) infiltrated with Agrobacterium tumefaciens . Herein, we describe a co-IP procedure in which structural maintenance of chromosome 1 (SMC1), identified from a Y2H screening, was verified as an interacting partner for microchidia 1 (MORC1), a protein well known for its function in plant immunity and epigenetics. SMC1 and MORC1 were transiently expressed in N. benthamiana when infiltrated by Agrobacterium with the respective genes. From this approach, we identified a region of SMC1 responsible for interacting with MORC1. The co-IP method, of which outputs are mainly from immunoblot analysis, provided information about target protein expression as well, which is often useful for troubleshooting. Using this feature, we showcased a PPI confirmation from our SMC1–MORC1 study in which a full-length SMC1 protein was not detectable, and, therefore, a subsequent truncated mutant analysis had to be employed for PPI verification.

Co-immunoprecipitation occurring with Transfection

Co-immunoprecipitation occurring with Western Blot

Co-immunoprecipitation occurring with RNA Immunoprecipitation Sequencing

Co-Immunoprecipitation of Long Noncoding RNAs

It is now estimated that the human genome encodes thousands of long noncoding (lnc)RNAs. These novel molecules are causing a paradigm shift in the field of molecular biology as a number of lncRNAs have been shown to be involved in a wide range of

It is now estimated that the human genome encodes thousands of long noncoding (lnc)RNAs. These novel molecules are causing a paradigm shift in the field of molecular biology as a number of lncRNAs have been shown to be involved in a wide range of biological functions including regulation of gene expression. Also, misregulation of lncRNAs has been observed in human diseases such as cancer and neurological disorders. These findings have spurred a huge interest in elucidating the functions and mechanisms of lncRNAs; and therefore, the need for new methods to do so. In this chapter, we discuss RIP-Seq, a method that is utilized to discover the lncRNA partners of a specific protein. This procedure involves immunoprecipitation of a protein from cross-linked cell lysate followed by reverse-cross-linking, isolation, and deep sequencing of RNAs, leading to the identification of all lncRNAs that are associated with a specific protein complex.

Investigating RNA–Protein Interactions in Neisseria meningitidis by RIP-Seq Analysis

Deep sequencing technology has revolutionized transcriptome analyses of both prokaryotes and eukaryotes. RNA-sequencing (RNA-seq), which is based on massively parallel sequencing of cDNAs, has been used to annotate transcript boundaries and has

Deep sequencing technology has revolutionized transcriptome analyses of both prokaryotes and eukaryotes. RNA-sequencing (RNA-seq), which is based on massively parallel sequencing of cDNAs, has been used to annotate transcript boundaries and has revealed widespread antisense transcription as well as a wealth of novel noncoding transcripts in many bacterial pathogens. Moreover, RNA-seq is nowadays also widely used to comprehensively explore the interaction between RNA-binding proteins and their RNA targets on a genome-wide level in many human-pathogenic bacteria. In particular, immunoprecipitation of an RNA-binding protein (RBP) of interest followed by isolation and analysis of all bound RNAs (RNA immunoprecipitation (RIP)) allows rapid characterization of its RNA regulon. Here, we describe an experimental approach which employs co-immunoprecipitation (coIP) of the RNA-binding chaperone Hfq along with bound RNAs followed by deep-sequencing of co-purified RNAs (RIP-Seq) from a genetically modified strain of Neisseria meningitidis expressing a chromosomally encoded Hfq-3×FLAG protein. This approach allowed us to comprehensively identify both mRNAs and sRNAs as targets of Hfq and served as an excellent starting point for sRNA research in this human pathogenic bacterium.

Co-immunoprecipitation occurring with SDS-PAGE

Isolation of Cytosolic Ribosomes Associated with Plant Mitochondria and Chloroplasts

Excluding the few dozen proteins encoded by the chloroplast and mitochondrial genomes, the majority of plant cell proteins are synthesized by cytosolic ribosomes. Most of these nuclear-encoded proteins are then targeted to specific cell compartments

Excluding the few dozen proteins encoded by the chloroplast and mitochondrial genomes, the majority of plant cell proteins are synthesized by cytosolic ribosomes. Most of these nuclear-encoded proteins are then targeted to specific cell compartments thanks to localization signals present in their amino acid sequence. These signals can be specific amino acid sequences known as transit peptides, or post-translational modifications, ability to interact with specific proteins or other more complex regulatory processes. Furthermore, in eukaryotic cells, protein synthesis can be regulated so that certain proteins are synthesized close to their destination site, thus enabling local protein synthesis in specific compartments of the cell. Previous studies have revealed that such locally translating cytosolic ribosomes are present in the vicinity of mitochondria and emerging views suggest that localized translation near chloroplasts could also occur. However, in higher plants, very little information is available on molecular mechanisms controlling these processes and there is a need to characterize cytosolic ribosomes associated with organelles membranes. To this goal, this protocol describes the purification of higher plant chloroplast and mitochondria and the organelle-associated cytosolic ribosomes.

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How to conduct a Co-immunoprecipitation (Co-IP)

Co-IP describes the immunoprecipitation of a protein of interest and its interacting partners.

Immunoprecipitation (IP ) is a technique used to isolate a protein from of an extract using a Nanobody or antibody (Ab). In co-immunoprecipitation (Co-IP), besides the IP of a specific protein, its interaction partner(s) are also pulled down and analyzed. In Co-IP, the protein that is pulled down is called the “bait” protein, while the interaction partner is called the “prey”. In Co-IP, the bait is directly precipitated with a Nanobody or Ab, which is coupled to (magnetic) beads. The prey is indirectly precipitated together with the bait. Therefore, Co-IP is used to enrich and/or identify interaction partners of the bait protein.

In the following blog, we focus on the precipitation of a GFP -tagged bait and its interacting prey protein. The Co-IP is conducted with GFP-Trap® comprising an anti-GFP Nanobody conjugated to beads. However, the principles described in this blog also apply to tags other than GFP or for Co-IPs using conventional antibodies.

The GFP-bait is a fusion protein of GFP (light green) and bait protein (dark green). The bait interacts with the prey protein (red). During Co-IP binds the added GFP-Trap® to the GFP bait..

A Co-IP consists of 5 steps.

The typical steps of a Co-IP are:

1. Preparation of the cell lysate

The preparation method of the cell lysate depends on the organism and bait and prey protein.

2. Binding of the bait and prey by GFP-Trap®

GFP-Trap® is added to the cell lysate and incubated for 1 h at +4°C.

3. Washing off unbound molecules, prey should remain bound to bait

The beads are washed a few times to remove unbound molecules while the prey protein should remain bound to the bait protein.

4. Elution of bound bait and prey from the beads

Bait and prey protein are released from the beads using SDS-sample buffer or acidic elution buffer.

5. Analysis of bait and prey on SDS-PAGE, Western blot, or by mass spectrometry

The eluate containing the bait and prey proteins is analyzed by SDS-PAGE, Western blot (WB) or mass spectrometry (MS).

Western blot analysis of Co-IP, probed with anti-GFP antibody (left lane) and anti-prey antibody (right lane).

Controls are crucial for a successful Co-IP experiment

Controls are an essential part of any Co-IP experiments – they are very important for the interpretation of your results and can help you to identify potential problems. In our Co-IP example, three different components should be analyzed: the bait-GFP, prey, and GFP only. Please note that a loading control is also often analyzed in addition, e.g., GAPDH , which is not shown here. In a comprehensive Co-IP experiment, different combinations of the components are usually examined to avoid false positive and false negative results.

1. Positive control

The positive control consists of a pulldown of the GFP-bait protein and GFP protein in the absence of the prey protein. GFP-bait and GFP are supposed to be found in the input and bound (IP:GFP) fractions. This experiment is conducted to confirm that the immunoprecipitation of the GFP-bait protein works under the chosen conditions, which is a pre-requisite of every Co-IP.

2. Negative control

The negative control consists of a pulldown of the prey protein in the absence of the bait protein. If the bait protein contains a larger tag like GFP, often a second pulldown is conducted with the prey protein and the tag only. The negative control experiments should confirm that the prey protein is not precipitated in the absence of GFP-bait or if only GFP is present. The prey protein is supposed to be present in the input fraction but not in the bound fraction.

3. Co-IP experiment

If the positive control and negative control experiments show the expected results, the final Co-IP experiment can be conducted. In the final experiment, both the GFP-tagged bait and prey protein are incubated together in the input. If the prey and GFP-bait are both found in the bound fraction, it can be concluded that the prey is only precipitated if the bait protein is present. This experiment, along with the positive and negative controls, demonstrates that the prey and bait proteins interact with each other.

A complete Co-IP experiment consists of all three parts. In publications, typical Co-IP data looks like this:

Controls are essential for meaningful results. Only by looking at all controls can you identify and resolve issues such as no pulldown of the prey or unspecific pulldown of the prey protein to avoid false results .

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Learn how to save precious hours on your IP, IF, and western blotting experiments

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Co-immunoprecipitation Co-IP

The complete co-ip technique guide..

By Ryan Hamnett, PhD

Co-immunoprecipitation (co-IP) is a powerful technique for the study of protein-protein interactions, based on the principle of immunoprecipitation (IP). A target protein is purified from a complex mixture along with its interacting partners using specific antibodies that are immobilized on a solid bead support. Protein complexes isolated by co-IP can be further investigated by western blot (WB), mass spectrometry (MS), and functional assessments to confirm hypothesized binding partners or identify new interacting proteins.

This guide aims to provide an overview of co-IP experimental design, important controls, protocols, and troubleshooting. The critical technical aspects of co-IPs will be discussed, providing an approach that can then be tailored to your specific research objectives.

Table of Contents

Co-IP Workflow

IP with Tagged Proteins

Co-IP Applications and Limitations

Sample types and preparation.

Lysis Buffers

Pre-clearing and Pre-blocking

Antibody selection.

Application Compatibility

Unidirectional Co-IP

• Bead Support

Agarose Beads

Magnetic Beads

Immobilizing Antibodies to Beads

Protein A and Protein G

• Direct immobilization
• Other immobilization methods

Immunoprecipitation

SDS-PAGE and Western Blot

Mass Spectrometry

Blue Native PAGE

Co-IP is one of a number of variations on the original IP format , which precipitates only a single protein target. Other IP formats enable the study of protein-protein (Co-IP), protein-DNA (ChIP) and protein-RNA (RIP) interactions. An overview of each method can be found in our IP Guide , while a comparison of the main features of each is below in Table 1.

Table 1: Key applications and differences between IP formats.

Co-IP follows a similar workflow to the individual IP workflow , but instead of purifying a single known protein (the “bait”), the aim is to co-precipitate unknown proteins (the “prey”) that are bound to the target antigen, on the basis that binding partners are likely related to, and may be required for, the function of the antigen. These prey proteins may include activators or inhibitors, kinases and other mediators of post-translational modifications (PTMs), ligands, and so on. Thus, protein complexes are precipitated and can be studied in similar ways to IP products, facilitating protein-protein interaction discovery.

As with IP, there are two standard approaches when performing a co-IP (Figure 1). In the pre-immobilized antibody method, also known as the direct method, an antibody specific for the target antigen is first immobilized onto a bead support. Once immobilized, the bead-antibody complex is added to the sample in order to capture the antigen. In the free antibody, or indirect, method, the antibody is added to the sample first, allowing antigen-antibody complexes to form. Then the beads are added to capture the immune complex.

Regardless of which approach is taken, the sample is then centrifuged in order to collect the beads in a pellet at the bottom of the tube, thus precipitating out the target antigen and any interacting proteins. This pellet is washed and re-pelleted several times to remove as much non-specifically bound material as possible. The target antigen and interacting proteins are then eluted by dissociating the precipitated proteins (and often the capture antibody) from the bead. The proteins are then ready to be used for downstream application such as sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) and WB, or MS.

Figure 1: Schematic of co-immunoprecipitation (co-IP) workflow.

Pull-down Assay

Pull-downs involve precipitating protein complexes based on protein-protein interactions captured on a solid bead substrate, and are therefore very similar conceptually to co-IPs. However, pull-downs do not use antibodies and are therefore not a type of immunoassay. Instead, the bait protein itself is immobilized to a bead without the antibody bridge.

IPs rely on the highly specific binding of an antibody to an antigen. In the absence of an antibody that exhibits strong specificity towards a protein of interest, one solution is to instead ‘tag’ the protein of interest with a peptide sequence or fluorescent protein for which a high affinity antibody is available. Tagging, which involves placing the DNA sequence for the tag at either the C- or N-terminal of the target protein sequence, can be achieved via genetic modification in model organisms, or expression of a recombinant protein from a plasmid or viral vector. The disadvantage of protein tagging is that the tags themselves may affect molecular interactions. Nevertheless, it has become a standard method for protein purification and co-IPs in particular, enabling researchers to express bait and prey proteins of interest in a cellular system of choice that is simpler to work with than live tissue.

• FLAG: DYKDDDDK
• c-Myc: EQKLISEEDL
• Hemagglutinin (HA): YPYDVPDYA
• V5: GKPIPNPLLGLDST

Co-IP can be used to detect both known and unknown binding partners. MS is often used to find previously undiscovered protein interactions, because it identifies proteins based on sequence information without requiring a prior hypothesis. 1 MS can also be used to confirm the presence and therefore binding of hypothesized (known) partners following a co-IP, but a simpler and more cost-effective method is usually to run the purified protein complex on a WB and probed using an antibody against the prey protein. A positive band will appear in both the input and the co-IP lanes if an interaction exists, whereas if there is no interaction, a band for the prey will only appear in the input lane. By systematically performing co-IP using different bait proteins and targeting the same or multiple prey proteins, protein-protein interaction networks can be elucidated.

Understanding how protein interactions change in different conditions, such as following drug treatment, can also be investigated by co-IP. Changes in the strength of the interaction may be inferred by a reduced or increased amount of prey protein appearing on a WB after co-IP, while biophysical and functional techniques can be used to more precisely measure changes in stoichiometry and activity.

Finally, co-IP followed by WB or MS is a useful tool for validating interactions observed using other techniques, such as the yeast 2-hybrid system, particularly given that interactions are detected in a physiologically relevant environment.

Despite its uses, co-IP is not without limitations. Co-IPs require considerable optimization to maintain protein-protein interactions, which can be easily disrupted by handling and buffer conditions. As a result, low-affinity and transient protein interactions may not be detected. Whether the interactions detected by co-IP are direct or indirect (e.g. through a third, intermediary protein) will not be immediately clear, and must be determined by careful use of controls and probing with different antibodies, and ideally validated with other assays. Finally, the antibody for the bait protein might bind to the site of interaction between the bait and prey, disrupting or blocking the interaction and leading to false negative results.

Co-IP begins with separating soluble proteins from a lysate, typically from cells or tissue. Tissue usually needs to be homogenized in order to make cells fully accessible to the lysis buffer, while lysis buffer can be directly added to cell culture after washing. Brief sonication of samples can sometimes help to disrupt the nuclear membrane to release nuclear proteins, but often agitation of cells or tissue homogenate in lysis buffer for 30 minutes on ice is sufficient to release soluble proteins. Insoluble material can then be pelleted, while the supernatant will be taken forward for co-IP.

The amount of total protein needed for successful IP will depend on the abundance of the protein and the affinity of the antibody. For a cell culture lysate, approximately 300 µg of total protein is a useful starting point. This can be increased up to 2 mg for low abundance proteins and may make visualization of the protein interaction easier, but more starting material can also increase background. If the target protein is only present in one region of the cell, such as the nucleus, a more refined option is to perform subcellular fractionation first in order to increase the abundance of the target as a proportion of the total input pool. This is useful to increase the relative abundance, but it may result in missing or overstating a given interaction if that interaction only occurs in a specific part of the cell.

Something that is essential to do in any co-IP protocol is to set aside 1-10% of the lysate (before the addition of any antibody, beads etc.). This is termed the input, and represents the starting sample material. The input will be run alongside the precipitate at the end of the experiment (e.g. by WB). This is a useful positive control to determine if the IP has worked: if a band is seen in the input but not the IP, then the IP did not successfully precipitate the protein.

In co-IP, the input has an additional function in that it confirms negative results: if a prey protein is probed for and seen in the input but not in the co-IP lanes, then an interaction between prey and bait has not been detected. A prey band that is significantly weaker in the co-IP lane than the input may also suggest a relatively weak interaction, although this can also depend on abundance and antibody efficiency. Efficiency of the antibody can be determined by comparing the strength of the target band in the IP lane to the band in the input lane, while specificity can be gauged by comparing the strength of consistent non-specific bands between the lanes. Finally, the input lane serves as quality control to ensure that the starting material is consistent across different experiments or samples.

One of the most important technical aspects of co-IP is the lysis buffer, the choice of which will depend on the sample type and purpose of the experiment, and typically requires significant optimization. Lysis buffers should stabilize native protein conformation, inhibit enzyme activity to decrease degradation and PTM modification, and rupture membranes for protein release from cells. The location of the protein in a cell (e.g. in the cytosol, nucleus) can affect how easily a protein will be released, which in turn will affect the choice of lysis buffer.

The most important lysis buffer consideration is whether the buffer used contains ionic or non-ionic detergent. Ionic detergents contain a charged head group and have a much stronger denaturing effect, which can result in altered protein conformations and protein-protein interactions. Non-ionic detergents are non-denaturing and less harsh than ionic detergents, meaning they are less likely to affect protein-protein interactions. Given that co-IP relies on protein-protein interactions, ionic detergents generally cannot be used.

RIPA (radio-immunoprecipitation assay) buffer is commonly used in WBs and sometimes recommended for individual protein IP, but it contains sodium deoxycholate and SDS (0.01-0.5%), which are ionic and will disrupt protein-protein interactions. Is is therefore rarely used for co-IP, though it should be stated that strong protein interactions may be maintained even in RIPA buffer at 4°C.

Buffers containing NP40 or Triton X-100 (0.1-2%) are useful, non-ionic alternatives to RIPA, but may result in slightly higher background. These weaker detergents are also not quite as effective at extracting all proteins from a cell. Even if the target proteins are effectively released, an antibody previously used for IP might only recognize the denatured form of the protein, in which case an alternative will need to be sought.

Buffers that completely lack detergent are also available for proteins that can be released using only physical disruption, usually consisting of just EDTA in phosphate buffered saline (PBS), though physical disruption strong enough to lyse a cell may also be capable of disrupting protein interactions.

Both ionic and non-ionic lysis buffers tend to contain NaCl and Tris-HCl, and usually have a slightly basic pH (7.4 to 8), though this can be optimized (between pH 6-9). Other buffer components that can be optimized include: salts (0-1 M) for maintaining ionic strength and correct tonicity for easy cell lysis; divalent cations such as Mg 2+ (0-10 mM) which can help to prevent DNA causing the solution to become viscous; and EDTA (0-5 mM) for chelating ions such as Zn 2+ that proteases require for proteolytic function.

Regardless of the composition of the buffer, performing all steps of a co-IP at 4°C or on ice is strongly recommended in order to minimize disruption to protein interactions.

Enzyme Inhibitors

The final components of lysis buffers are inhibitors of proteases and enzymes that alter protein PTMs, particularly phosphatases. Protease inhibitors prevent the degradation of target proteins, while PTMs are often necessary for protein-protein interactions and so must be maintained for co-IP. Preserving PTMs is obviously essential if the target of the IP is to understand the PTM state of target proteins.

Inhibitors should be added fresh, immediately before lysis buffer use. Table 2 and Table 3 contain common inhibitors used in lysis buffers.

Table 2: Commonly used protease inhibitors in co-IP. Note that protease inhibitor cocktails are commercially available (often as tablets) and provide a convenient way to ensure protease inhibition.

Table 3: Commonly used phosphatase Inhibitors in co-IP. Phosphatase inhibitor cocktails are commercially available (often as tablets) and provide a convenient way to ensure phosphatase inhibition. Note that while phosphatase inhibitors are typically included in IP buffers, inhibitors of other PTMs, such as ubiquitination and methylation, are included only as needed, determined by the aims of the experiment.

Pre-clearing is an optional but often worthwhile step to reduce non-specific binding in co-IP. Pre-clearing refers to incubating samples with the beads (including Protein A/G or other attachment substrates), or with a nonspecific antibody from the same host species as the IP antibody immobilized to the beads, in order to remove lysate components that bind to beads or immunoglobulins non-specifically. This prevents these components from being carried through and ultimately eluted, therefore resulting in a purer final product containing predominantly the target antigen.

Reducing non-specific binding can also be achieved by pre-blocking the bead. This works in a similar way to blocking in immunostaining, western blotting and ELISA. The bead is incubated with a mild blocking buffer containing 1-5% BSA, non-fat milk, 1% gelatin or 0.1-1% Tween-20 to block sites of non-specific binding on the bead, preventing lysate components from binding.

Pre-clearing is not necessary if the target is particularly abundant, and is less important if using magnetic beads over agarose beads, because magnetic beads are less prone to non-specific binding.

The supernatant from a pre-clearing step can be kept for WB analysis to confirm that no target antigen was removed during the pre-clearing step.

Antibodies are generated by the immune system of a host organism (e.g. mouse, rabbit) that has been repeatedly immunized with a specific antigen. The antibodies recognize and specifically bind to that antigen with a high affinity, allowing the protein to be precipitated from the lysate. High specificity is essential for a successful co-IP, as non-specific binding can result in high background or false positive results.

To confirm the specificity of the antibody-antigen interaction, an isotype control should be included. The isotype control consists of a non-immune antibody of the same isotype as the experimental antibody. This should not have any affinity for the target antigen, but it may non-specifically bind to other factors, therefore confirming that the precipitated protein band in a WB is specifically recognized by the chosen antibody. Any bands in both the co-IP and isotype control lanes in WB analysis is likely to represent a protein that binds immunoglobulins non-specifically.

The following factors should be taken into account when selecting an antibody.

Primary antibodies will recognize antigens only from certain species, which will be the species that the immunizing antigen was originally from, as well as closely related species. For example, an antibody that recognizes a mouse protein target will often also recognize the same protein in rat, but possibly not in fish. The key determinant of this is how similar the epitope sequence is between species, allowing researchers to predict if an available antibody will work in a non-validated species.

Both monoclonal and polyclonal primary antibodies can be used to detect the protein of interest in co-IPs. Monoclonal antibodies correspond to a single epitope for a given antigen, whereas polyclonal antibodies may recognize multiple epitopes. As a result, polyclonal antibodies are often preferred to monoclonals for co-IP because they offer a high chance of capturing the target, and a smaller chance that the interaction site between bait and prey will be blocked.

Clonality can also be a factor when considering analysis (see also the Analysis section). Using two different antibodies for the precipitation and WB stages can be advantageous to ensure high specificity, using one polyclonal and one monoclonal. A polyclonal antibody for co-IP gives the highest capture efficiency, while a monoclonal antibody in the WB gives the highest detection specificity.

Co-IP, ChIP and RIP always seek to isolate proteins in their native conformation, which is essential for physiologically relevant molecular interactions, so a chosen antibody should recognize this form rather than denatured protein (as is common in western blotting). This contrasts to individual protein IPs, which can be performed on denatured samples. If an IP-validated antibody is not available, it is usually acceptable to use one that has been validated for IHC or ELISA, which also work with native proteins.

Immunoprecipitation of a protein complex may only be observed when one of the interacting proteins is the bait, but not in the reciprocal arrangement where the other protein is the bait. This can be due to the abundances of the two proteins and how common being in the bound state is for each partner. If this occurs, it is recommend to use a capture antibody specific to the protein that has the highest percentage of its total population bound to the partner. However, this is obviously dependent on the aims of the experiment: if searching for novel binding partners of a protein of interest, changing the target protein is not an option. In this case, trying alternative antibodies for the protein of interest and optimizing the co-IP conditions should be considered.

Bead support

The type of beads chosen when performing co-IP is an important consideration and will dictate certain aspects of the overall protocol. Beads are usually either agarose (or Sepharose, which is a tradename for a crosslinked, beaded-form of agarose) or magnetic beads. Table 4 summarizes the key differences between agarose and magnetic beads, which are described further below.

Table 4: Key features of agarose and magnetic beads in co-IP.

Agarose beads tend to be 50-150 μm in size, and they have a very high binding capacity because they are porous, creating a very high surface area per bead that is available for binding to antibodies. However, this can create a requirement to use a large amount of antibody to ensure that every surface of the agarose beads is covered in antibody, lest any unoccupied agarose non-specifically bind to lysate components and increase background signal. To solve this issue, it is possible to back-calculate from the amount of expected analyte to the amount of antibody needed for detection, to the amount of agarose needed to hold the antibody. The downside is that this approach requires significant knowledge or prior experience to estimate such quantities. An alternative would be to simply saturate the beads with excess antibody, but antibody is typically expensive and limiting. The best option is therefore to pre-clear the lysate to remove anything that will non-specifically bind to the agarose.

For co-IP, agarose beads may have a disadvantage in that protein complexes (as opposed to individual proteins) may not fit into the porous beads, leaving only the outer surface available for binding.

Magnetic beads lack the porosity of agarose because they are solid spheres, resulting in only the external surface being available for binding. While this may at first glance suggest that magnetic beads have much lower binding capacity, they are much smaller than agarose particles at 1-4 μm, meaning that there can be far more beads per unit volume, so the ultimate binding capacities of the two are similar, with agarose maintaining a small advantage.

Magnetic beads offer several other advantages over agarose for use in co-IP, however. They are highly uniform in size and have a consistent binding capacity, which can make experiments more consistent between runs. Magnetic beads are manipulated using powerful magnets that pull the beads to one side of a tube so that the supernatant can be removed, obviating the need for centrifugation. Centrifugation can be stressful for protein complexes and can lead to a loss of yield compared to the gentler magnet-based washing steps, thus final yield can be higher with magnetic beads. Magnet-based manipulation is faster and makes co-IPs more amenable to higher throughput applications.

Immobilizing antibodies to the solid phase (beads) is essential for co-IP, enabling the precipitation of the target protein complexes. While the most common method is to use Protein A or G to capture the co-IP antibody, various approaches can be used, each of which has its own benefits and applications. These approaches are illustrated in Figure 2 and explained in more detail below.

Figure 2: Approaches to IP antibody immobilization on beads. A , Protein A, G or A/G bind to the Fc region of antibodies. B , Antibodies can be crosslinked to Protein A, G or A/G to prevent eluting the antibodies during antigen elution. C , Protein L binds to the light chain of antibodies. D , Direct immobilization of antibodies to beads obviates the need for Protein A, G or L, and prevents antibody elution. E , Streptavidin-conjugated beads capture biotinylated IP antibodies with very high affinity. F , Secondary antibodies directly bound to beads can be used to capture IP antibodies from a specific host or of a specific isotype, but offer less flexibility than Protein A or G.

The most common approach to immobilize the antibody to the chosen bead is with Protein A 2 or Protein G, 3 which are covalently bound to the beads following bead activation with a coupling agent such as cyanogen bromide or N-hydroxysuccinimide (NHS). Protein A and Protein G are derived from bacteria, and bind to the heavy chains of the antibody’s Fc region (Figure 2a). Binding to this site has the advantage of orienting the antibody such that the Fab region is clear and directed away from the bead to bind to the target protein.

Due to their promiscuity in generally binding immunoglobulin Fc regions, care must be taken when performing co-IP on a sample that contains immunoglobulins besides the added capture antibody, such as serum. In this instance, the capture antibody would need to be added to the beads first (pre-immobilized/direct method) and could be covalently bound to Protein A or G (see Covalently Linking Capture Antibodies to Protein A/G below). 4

The choice between Protein A and G will depend on the capture antibody’s host species and isotype, because Protein A and G exhibit different binding affinities for different immunoglobulins (Table 5). To circumvent this issue, the recombinant Protein A/G was engineered to contain four Protein A and two Protein G binding sites, and which binds all of the subtypes that Protein A and G bind individually.

Table 5: Binding affinities of Protein A and Protein G for different immunoglobulins. ++++ refers to strong binding, + is weak binding, X is no binding affinity.

The interaction between antibodies and Protein A/G will occur in most physiological buffers, but binding, and therefore capacity, can be increased by using dedicated Protein A or Protein G binding buffers. Protein A binds IgG best at pH 8.2, whereas protein G binds best at pH 5. However, these buffers may then not be suited for antigen binding, thus optimization of binding buffers is always necessary.

Covalently Linking Capture Antibodies to Protein A/G

Binding of a capture antibody to the solid phase via Protein A/G is not covalent, meaning the antibody will be eluted along with the target antigen at the end of the experiment. If this is followed by a WB, two antibody bands can be visible on the blot at 25 kDa and at 50-55 kDa, corresponding to the light and heavy chains after denaturing. These bands will obscure the signal from the target antigen if the antigen is a similar molecular weight. One method of preventing this (others are discussed in the Analysis section), the IP antibody can be covalently linked to Protein A/G (Figure 2b). Covalent linkage is achieved using a cross-linker such as disuccinimidyl suberate (DSS) or bissulfosuccinimidyl suberate (BS3). These are simple carbon spacers with NHS ester groups at either end, which react with primary amines on lysine residues in the proteins to form a covalent link between them.

Protein L is derived from Peptostreptococcus magnus and binds to the kappa light chain in the variable domain of antibodies (Figure 2c). 5 Protein L is most commonly used with mouse and rat IgM capture antibodies, which bind poorly to Protein A/G. This is due to the heavy chains of IgMs interacting with each other, forming their classic pentameric structure and blocking Protein A/G binding. Protein L is also useful because it does not bind to goat, sheep or cow antibodies, and so can be used for cell culture lysates that contains serum from those species.

Direct Immobilization

Direct immobilization refers to covalent bonding of the antibody to the beads without Protein A, G or L acting as an intermediary (Figure 2d). This is achieved by activating beads with aldehyde groups, which are highly reactive with primary amine groups on antibodies, followed by reduction with sodium cyanoborohydride to form a stable secondary amine bond. This approach can be useful if Protein A/G/L are not compatible with the subclass of IP antibody.

Similarly to covalent bonding of the antibody to a protein support, direct immobilization will prevent the antibody from co-eluting with the target antigen, so it will not be seen on the WB membrane. As a result, the antibody-bound beads can theoretically be used in multiple experimental runs because the antibody will remain intact and permanently bound. Removing Protein A/G as a component in the system also removes it as a source of non-specific binding during the co-IP.

The slight disadvantage of this approach is that antibodies are coupled to the bead in a random orientation, as opposed to the directed orientation that results from Protein A/G binding. However, this only has a minor effect on capacity and co-IP yield.

Other Immobilization Methods

Biotin-avidin binding

Avidin is a protein found in egg whites, while streptavidin is from purified from the bacterium Streptomyces avidinii, but they both have an extremely high and specific affinity for biotin. Biotin is a small 244 Da vitamin that is easy to covalently bond to protein (the carboxyl group in biotin can be modified with reactive groups such as NHS esters, maleimides or hydrazides that target amines (-NH 2 ), sulfhydryls (-SH) or aldehydes (>C=O), respectively, on proteins).

The affinity between biotin and avidin is extremely strong and specific, making it an ideal system for affinity purification, in which a biotinylated antibody is bound by streptavidin-conjugated beads (Figure 2e). Only harsh buffers can dissociate biotin-avidin, such as 8 M guanidine-HCl at pH 1.5, meaning the association will remain throughout all wash steps and not be eluted at the end.

Because any protein can be biotinylated, this system is also adaptable to general pull-downs in which antibodies are not used. The extremely high affinity of biotin-avidin offers a distinct advantage in pull-downs whereby prey proteins can be eluted in a regular elution buffer, but the bait protein will remain attached to the beads. The disadvantage when using biotin-streptavidin for co-IPs is that antibodies are not necessarily oriented in the optimal direction with the Fab region accessible, but this has only a minor effect on co-IP efficiency.

Secondary antibodies

An indirect co-IP method that is analogous to indirect methods used in IHC, ELISA and WBs uses secondary antibodies directly immobilized to beads (Figure 2f). These secondary antibodies recognize the host species of the IP capture antibody to ensure specificity. For example, a mouse IP antibody could be bound by a goat anti-mouse secondary antibody attached to the beads. The disadvantage of this approach is that it does not offer the flexibility of Protein A/G-conjugated beads, which will bind to an IP antibody from any host.

As illustrated in Co-IP Workflow , there are two approaches for IPs (Figure 1): the pre-immobilized antibody method in which the antibody is incubated with the bead first, and the free-antibody method in which the antibody is added to the sample lysate first before the beads are added. Using free antibodies can be beneficial in instances where the target protein is low abundance as it can give the highest yield, though it can also give higher background than the pre-immobilized approach. The pre-immobilized method must be used when using direct immobilization approaches, which is highly beneficial in preventing the antibody from being eluted with the antigen, and typically outweighs any minor reduction in yield.

Once the approach has been decided upon, the antibodies are added to the sample lysate and incubated to allow adequate binding. How long this incubation needs to be to ensure sufficient target capture depends on how abundant the antigen is, but anywhere between 1 hour and overnight at 4°C is recommended. Over-incubation, particularly at room temperature, may result in non-specific binding and therefore high background, and co-IPs should be performed at 4°C as much as possible to preserve protein interactions.

After the antibodies and beads have been added, the bead-antibody-antigen complexes are pelleted, allowing the supernatant to be removed. At this point, it is good practice to keep the supernatant until the co-IP has been verified as successful, in case the majority of the target protein complex is still contained within.

The beads then need to be washed to remove non-specific binding of other lysate components to the bead, immobilization substrate or antibody. Washing is done using either the original lysis buffer or a dedicated wash buffer, either of which should contain protease and phosphatase inhibitors as the original lysis buffer did. Washing must be carefully optimized for co-IP to find the right level of stringency that will remove non-specifically bound proteins but maintain target protein interactions, particularly if they are relatively weak.

Standard wash buffers consist of PBS or Tris-buffered saline (TBS), which contain physiological levels of salt at a physiological pH, and 0.5-1% of a mild detergent such as NP-40 or Triton X-100. Salt (NaCl) can be increased up to 1 M to increase the stringency of the wash by reducing ionic and electrostatic interactions, while reducing agents (e.g. 1-2 mM dithiothreitol (DTT) or β-mercaptoethanol (BME)) can reduce non-specific interactions mediated by disulfide bridges or nucleophilic attractions.

Elution refers to dissociating the target protein complex (and often the co-IP antibody) from the beads to obtain a pure protein sample. While there are some analysis options that beads are compatible with, resolution of protein size by SDS-PAGE and WB is affected by beads. Elution of the protein will also elute the antibody, which can result in bands being present on the subsequent SDS-PAGE and WB, unless the antibody was covalently linked to either Protein A/G or directly to the beads.

Given how prevalent western blotting is following co-IP, a standard elution buffer is SDS-PAGE sample buffer. While denaturing buffers have been avoided for co-IP up until now, probing for bait and prey proteins on a WB does not require the proteins to be in their native conformation or still bound to each other. SDS-PAGE sample buffer is highly denaturing and will therefore easily disrupt affinity-based interactions and prepare the bait and prey for WB.

SDS-PAGE sample buffer will not be suitable for all sample types or downstream applications due to its harshness. A more generally applicable, non-denaturing elution buffer is 0.1 M glycine buffer at pH 2.5-3. This can be useful for protein characterization, sequence determination, and crystallization, amongst others. The low pH will cause non-crosslinked Protein A/G-antibody and the antibody-antigen, and bait-prey interactions to dissociate, though it is not universally successful and even this can be harsh for some antigens, causing them to denature. Eluting using urea buffer (6-8 M urea, 20 mM Tris, 100 mM NaCl; pH 7.5) is another option, which is particularly useful for downstream MS. Finally, no elution is an option in some cases, keeping the antigen attached to the antibody and beads, because some bioassays and techniques to study protein-protein interactions are unaffected by the presence of the beads. 6

The proteins purified by co-IP can be used and studied by a large range of techniques. While the most common are WB and MS (discussed below), protein abundance may be measured by ELISA , function can be determined by various activity assays, and binding and physical characteristics can be studied through biophysical techniques, NMR and crystallization.

SDS-PAGE involves first denaturing proteins so that secondary and tertiary structure will not interfere with protein migration on a gel, and then running the denatured protein samples on a gel to determine size and abundance (see our full Western Blot guide ). Unlike the relatively pure sample resulting from individual protein IP, which may be visualized on the gel by a Coomassie stain, there are likely to be many different proteins contained within a co-IP eluate that would interfere visualization or interpretation. The proteins should therefore be transferred from the gel to a nitrocellulose or PVDF membrane and blotted using an antibody that is specific for either the bait protein or hypothesized prey proteins. Some scenarios for experimental set up and interpretation of co-IP results via WB can be seen in Figure 3 below. These results can be further complicated by the addition of extra conditions, such as drug treatment, and extra prey proteins of interest.

Alongside the Co-IP lane, several other samples should be included as important controls. An input lane of 1-10% of the starting lysate material is always included (Figure 3) to get a sense of co-IP efficiency and sensitivity (as described in Sample Types and Preparation ), and most importantly to confirm that negative results are true negatives rather than unsuccessful probing with antibodies against prey proteins. Another indicator of efficiency is to run the supernatant from the co-IP as a negative control; if the co-IP was successful and efficient, this lane should be negative. An isotype control, in which the co-IP is performed with an antibody of the same isotype but that does not recognize the target antigen, is also often included to exclude non-specific binding. For more information on controls, see Co-IP Controls .

Figure 3: Interpretation of co-IP western blots. A , Co-IP performed on cells transfected with proteins under investigation. Cells were transfected with bait protein only (positive control, lanes 1 + 4), prey proteins only (negative control, lanes 2 + 5) or bait and prey (lanes 3 + 6). No proteins are precipitated in the absence of the bait protein (lane 5). Lane 6 shows the co-IP experimental condition, and suggests that the bait does not interact with Prey 2. Blotting for a protein known not to interact with the bait and which will be endogenously expressed in the cell culture system, such as GAPDH, is included as an additional negative control. B + C , Co-IP performed on tissue, in which endogenously expressed proteins are under investigation rather than transfected proteins. Here a negative control can consist of a bait knockout (KO; B ) or an IP using an isotype control antibody (IP:Neg; C ).

Avoiding Antibody Bands

When blotting for the bait protein, the primary antibody used for western blotting will ideally be different to the capture IP antibody. Not only will this increase specificity of the final detection, but it will also prevent detection of any eluted IP antibody. This is because the WB and IP antibodies can then be from different species, meaning the WB secondary antibody will only detect the WB primary antibody (Figure 4). For example, if antigen X is immunoprecipitated by mouse anti-X, but blotted for with rabbit anti-X, then the secondary goat anti-rabbit antibody will only recognize the WB antibody.

Nevertheless, the same antibody can be used for both IP and WB, so long as the target antigen is not the same size as the bands representing the light and heavy chains of the IP antibody after denaturing. These bands are at 25 kDa and 50-55 kDa, respectively, and will obscure the target signal if the target antigen is not sufficiently larger or smaller. Alternatively, covalently linking the antibody directly to the beads or to Protein A/G will prevent its elution and therefore prevent it from being detected on the membrane. A final option is to use secondary antibodies with greater specificity for primary antibody structure. Secondary antibodies are available that only recognize the intact antibody conformation, not the denatured form, or it is possible to use a light-chain-specific secondary antibody. This will recognize the WB primary antibody and only the light chain of the IP antibody, producing a band at 25 kDa but not 50-55 kDa.

Figure 4: Contaminating antibody bands on western blot after IP. A , Western blot (WB) primary antibody is the same antibody as the (now denatured) IP capture antibody, resulting in the WB secondary antibody recognizing both in the eluate. Bands for the antibody light chain (~25 kDa) and heavy chain (~50 kDa) appear on the WB (black bands) alongside the precipitated antigen (red band). B , WB is performed using a primary antibody raised in a different host (mouse) compared to the IP antibody (rabbit), so the WB secondary does not recognize the denatured IP antibody. No antibody bands appear on the WB. C , WB is performed using a secondary antibody that only recognizes the intact, non-denatured form of the primary antibody. No antibody bands appear on the WB. D , WB is performed using a secondary antibody that only recognizes the light chain of the primary antibody, so the band corresponding to the heavy chain does not appear on the WB.

MS is an extremely sensitive method of detecting, identifying and quantifying proteins in a sample using peptide sequences resulting from enzymatic digestion of proteins. For MS, proteins are first cleaved into smaller peptide fragments using an enzyme such as trypsin. The peptides are then ionized, accelerated through the mass spectrometer, deflected by a powerful magnet, and ultimately detected.

MS works by measuring the mass/charge ratio (m/z) of molecules, and ionization is necessary so that they are affected by the magnetic field. Heavier ions will be deflected less by the magnet than lighter ions, and ions with a greater charge will be deflected more than ions with less charge. Therefore, the combination of mass and charge determines how strong the magnetic field needs to be in order to deflect the particles to the detector. The abundance of particles with a given m/z is detected and that m/z can then be computationally assigned to a peptide sequence using known sequence information. Using short peptides, rather than intact protein, is important because it increases the chance of being able to assign a given m/z to a peptide sequence. In contrast, a single, large m/z value for an entire protein could represent any number of possible sequences. The disadvantage of this bottom-up approach is that the protein identity must then be computationally reconstructed based on the peptide fragments that are detected, which is achieved using bioinformatics.

Due to its ability to detect proteins based on sequence information, MS is commonly applied alongside co-IP to identify previously unknown protein interactors and their relative abundance, and establishing protein interaction networks. 1,7,8 MS can further enhance co-IP experiments by comparing samples in different experimental groups, for example determining if a drug treatment affects a protein’s binding partners, abundance, or PTMs.

Blue Native PAGE (BN-PAGE) is an alternative electrophoretic method to SDS-PAGE that separates protein complexes in their native form. 9-12 BN-PAGE relies on solubilizing protein complexes with non-ionic detergents before adding the anionic dye Coomassie blue G-250. Coomassie imparts a charge shift on the proteins that allows proteins to migrate to the anode during BN-PAGE, but, unlike SDS, Coomassie does not denature proteins. Separation therefore occurs based primarily on molecular weight, shape and size as they pass through progressively smaller acrylamide gel pores until they reach their size-dependent specific pore-size limit. 11 Combining BN-PAGE with MS can be a powerful approach for understanding the composition of large protein complexes. 13

Diagrams created with BioRender.com.

• Maccarrone, G., Bonfiglio, J. J., Silberstein, S., Turck, C. W. & Martins-de-Souza, D. Characterization of a Protein Interactome by Co-Immunoprecipitation and Shotgun Mass Spectrometry. in Multiplex Biomarker Techniques: Methods and Applications, 1546, 223–234.
• Hjelm, H., Hjelm, K. & Sjöquist, J. Protein a from Staphylococcus aureus. Its isolation by affinity chromatography and its use as an immunosorbent for isolation of immunoglobulins. FEBS Lett. 28, 73–76 (1972).
• Björck, L. & Kronvall, G. Purification and some properties of streptococcal protein G, a novel IgG-binding reagent. J. Immunol. Baltim. Md 1950 133, 969–974 (1984).
• Michielsen, E. C., Diris, J. H., Hackeng, C. M., Wodzig, W. K. & Van Dieijen-Visser, M. P. Highly Sensitive Immunoprecipitation Method for Extracting and Concentrating Low-Abundance Proteins from Human Serum. Clin. Chem. 51, 222–224 (2005).
• Nilson, B. H., Solomon, A., Björck, L. & Akerström, B. Protein L from Peptostreptococcus magnus binds to the kappa light chain variable domain. J. Biol. Chem. 267, 2234–2239 (1992).
• Peled, M., Strazza, M. & Mor, A. Co-immunoprecipitation Assay for Studying Functional Interactions Between Receptors and Enzymes. J. Vis. Exp. JoVE 58433 (2018).
• Lagundzin, D., Krieger, K. L., Law, H. C.-H. & Woods, N. T. An optimized co-immunoprecipitation protocol for the analysis of endogenous protein-protein interactions in cell lines using mass spectrometry. STAR Protoc. 3, (2022).
• Free, R. B., Hazelwood, L. A. & Sibley, D. R. Identifying Novel Protein-Protein Interactions Using Co-Immunoprecipitation and Mass Spectroscopy. Curr. Protoc. Neurosci. 46, 5.28.1-5.28.14 (2009).
• Schägger, H. & von Jagow, G. Blue native electrophoresis for isolation of membrane protein complexes in enzymatically active form. Anal. Biochem. 199, 223–231 (1991).
• Darie, C. C. Investigation of Protein-Protein Interactions by Blue Native-PAGE & Mass Spectrometry. Mod. Chem. Appl. 1, 1–3 (2013).
• Wittig, I., Braun, H.-P. & Schägger, H. Blue native PAGE. Nat. Protoc. 1, 418–428 (2006).
• Fiala, G. J., Schamel, W. W. A. & Blumenthal, B. Blue Native Polyacrylamide Gel Electrophoresis (BN-PAGE) for Analysis of Multiprotein Complexes from Cellular Lysates. J. Vis. Exp. JoVE 2164 (2011).
• Pardo, M., Bode, D., Yu, L. & Choudhary, J. S. Resolving Affinity Purified Protein Complexes by Blue Native PAGE and Protein Correlation Profiling. JoVE J. Vis. Exp. e55498 (2017).

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Immunoprecipitation (IP) protocol

Immunoprecipitation is a protein purification method that allows us to isolate a specific protein from the mixture using antigen-antibody interaction.

An antibody for the protein of interest is incubated with a cell extract enabling the antibody to bind to the protein in solution. The antibody/antigen complex is then pulled out of the sample using protein A/G-coupled agarose or magnetic beads. This isolates the protein of interest from the rest of the sample. The protein of interest can then be analyzed by western blot, mass spectrometry, direct ELISA, or other analytical techniques.

Stage 1 - Preparing the lysates

The first step is to lyse the cell or tissue samples in a suitable buffer to release proteins into the solution. We provide a non-denaturing lysis buffer (ab152163) suitable for isolating most proteins from cells or tissues.

Some proteins are more difficult to isolate than others, so you may need to create and optimize your own buffer. The ideal lysis buffer will minimize protein denaturation while releasing enough proteins from the sample.

Non-ionic detergents, such as NP-40 and Triton X-100, are less harsh than ionic detergents, such as SDS and sodium deoxycholate. Other variables that can affect the success of immunoprecipitation include salt concentration, divalent cation concentration, and pH.

Materials required

• Suitable lysis buffer (example: ab152163 , or develop your own)
• Protease inhibitor cocktail (example: ab65621 )
• Phosphatase inhibitor cocktail (optional – example: ab201112 ) 

Prepare an appropriate lysis buffer for your protein.

Keep samples, buffers, and equipment on ice throughout the process.

Add protease inhibitors to the buffer.

• Include phosphatase inhibitors for phosphorylated proteins.

Isolate your cells and suspend them in a lysis buffer.

• Some adherent cells may require enzymatic or mechanical detachment. We detach adherent cells with TrypLE, spin to remove the media, resuspend the pellet in PBS and spin two more times, each time resuspending the pellet in PBS to wash the cells, leaving a washed cell pellet at the end.
• Suspension cells can be washed in PBS and spun down into a pellet before suspension in the ice-cold lysis buffer in the next step.

Add 300 µL more lysis buffer if cells do not resuspend well.

To lyse the cells, add ice-cold lysis buffer directly to the cell pellet and resuspend it.

• Incubate the cells with lysis buffer on ice for 10 mins (without agitation).
• Sonicate the lysate 3 times in ice-cold water.

Optimization may be required.

Only agitate - using vortex - if cells remain clumped.

Spin down the suspension to pellet insoluble contents.

• Keep the supernatant – this is your lysate.
• Centrifuge the suspension at 8,000 x g for 10 minutes at 4 °C. 
• Place the supernatant in a fresh tube on ice.

You may have to adjust the centrifugation force and time for your cell type. Leukocytes, for example, only need light centrifugation.

Determine the protein concentration in your lysate using a Bradford or BCA assay.

• Suppose the protein concentration at this stage is low, and your protein resides in the nucleus or mitochondria. In that case, you could consider fractionating your original sample to produce a more concentrated lysate. 
• We offer cell fractionation kits for this purpose.

If not using immediately, snap freeze aliquots in liquid nitrogen and store at -80°C.

Stage 2 - Pre-clearing the lysates (optional)

Pre-clearing lysates is an additional optional step that can help to increase the purity of proteins obtained by IP. This step involves incubating the lysate with only beads or beads plus an isotype control, depending on the beads used, to precipitate unwanted proteins. One can normally skip this step when using high-quality beads without severe bead adsorption effect.

Pre-clearing the lysate can help reduce non-specific binding and reduce background. However, if the final detection of the protein is by western blotting, pre-clearing may not be necessary unless a contaminating protein interferes with the visualization of the protein of interest.

Here we provide a protocol example for pre-clearing the lysates using beads combined with an isotype control.

• Isotype control antibody
• Your lysate
• Protein A/G magnetic beads (example ab214286 )
• Magnetic rack

Prepare the beads and mix them with isotype control antibodies.

• Follow the bead manufacturer’s advice to prepare the antibody-bead complexes.

Add a slurry of beads with isotype control antibodies to the lysate.

• Add ~100 µL of beads per 1 mL of lysate.
• Incubate for 10 – 30 mins at 4°C, with gentle agitation.

Remove the antibody-bead complexes.

• Place the tube on a magnetic rack for ~1 min; the beads should be pulled by the magnet to the bottom of the tube.
• Aspirate the solution, leaving the beads behind, and transfer this solution to a new tube.

Proceed to IP on your pre-cleared lysate.

Stage 3 - Immunoprecipitation and washing

Now we have lysed your cells and pre-cleared the lysate if necessary, so we’re ready to run the IP.

There are two main methods to immunoprecipitate proteins. The first approach is mixing the antibody with the lysate and then adding Protein A/G beads to the antibody-lysate complex. This method yields high purity of protein; however, the antibodies are also co-eluted with the protein of interest, which sometimes creates difficulties in western blot detection.

The second approach is preparing the antibody-bead complexes and then incubating them with your lysate to pull out the protein of interest. This method gives a lesser yield than the first one but avoids the problem of co-elution of antibodies.

We suggest you run IP according to the protocols below. If using an isotype control antibody, you can run the same procedure and compare this with your IP for the protein of interest when analyzing the results.

• Primary antibody
• Protein A/G magnetic beads (example: ab214286 )

Incubate your lysate with antibodies at 4°C overnight, with gentle agitation.

Add the Protein A/G magnetic beads to the antibody-lysate complex.

• Incubate for 1–4 hours at 4°C, with gentle shaking.

Follow the bead manufacturer’s advice on how to incubate beads with an antibody-lysate complex.

Volumes of lysate and beads solution might require optimization.

Wash the beads three times with PBS or another wash buffer of your choice to remove unbound impurities.

• Aspirate and discard the solution, keeping the beads in the tube.
• Add ~ 1 mL of fresh lysis buffer to the beads and repeat the steps above.

The protein of interest should now be specifically bound to the antibody coating the beads.

Stage 4 - Elution

After washing, we’re ready to elute the protein from the beads. Elution can be done in various buffer conditions, including glycine (non-denaturing), Laemmli (denaturing), and urea buffers.

Of all these buffers, Laemmli buffer containing SDS is the harshest, as it will also elute non-covalently bound antibodies and antibody fragments along with the protein of interest. In contrast, glycine buffer gently elutes the protein with a reduced amount of eluted antibody.

This method produces a slightly less concentrated sample but keeps the protein in its native state. Since this is a non-denaturing buffer, the beads can also be re-used once the protocol is complete.

• 0.1 – 0.2 M Glycine, pH 2.6
• Tris-HCl, pH 8.5
• Your samples bound to beads

Add an equal volume of glycine to the beads.

• Incubate for 10 mins at room temperature, with gentle agitation.
• The low pH should separate the protein of interest from the antibody beads.

Separate the beads from the solution using a magnetic rack (for magnetic beads) or light centrifugation (for agarose beads).

• For magnetic beads: place the tube on a magnetic rack for ~1 min; the beads should be pulled by the magnet to the bottom of the tube.
• For agarose beads: spin down the tube at the manufacturer's recommended speed for 2 mins.

You may wish to repeat Steps 1 – 2 several times to maximize the protein eluted.

The beads can be re-used by washing them in a lysis buffer to remove the glycine.

Neutralize the pH of the solution by adding Tris-HCl.

Optional: if not using immediately, aliquot samples, snap freeze, and store at -80°C.

Analyze the IP results using western blot or another technique of your choice.

• Compare the results of IP performed with your capture antibody and isotype control antibody.