Técnicas Baseadas em Imunoprecipitação: Purificação de Proteínas Endógenas Usando Esferas de Agarose
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Fonte: Susannah C. Shissler1, Tonya J. Webb1
1 Departamento de Microbiologia e Imunologia, Universidade de Maryland, Baltimore, MD 21201
Imunoprecipitação (IP, também conhecido como ensaio ‘pull-down’) é uma técnica amplamente utilizada que tem aplicações em uma variedade de campos. Concebido pela primeira vez em 1984, foi refinado em 1988 (1, 2). O objetivo fundamental do IP é a purificação e isolamento de uma proteína específica usando um anticorpo contra essa proteína. A palavra “imuno” refere-se ao uso de um anticorpo, enquanto a palavra “precipitação” refere-se a retirar uma substância específica de uma solução. A proteína alvo pode ser endógena ou recombinante. A maioria das proteínas recombinantes tem uma tag de epítope (ou seja, myc ou flag) anexada a elas para simplificar a purificação subsequente. Normalmente, é mais fácil otimizar a proteína recombinante IP porque os anticorpos contra tags de epitope recombinantes são muito fortes e eficazes. Anticorpos contra proteínas endógenas têm eficácia extremamente variável – tornando muito mais difícil otimizar esses IPs. Um passo necessário após a imunoprecipitação é a verificação da purificação. A proteína isolada é resolvida usando SDS-PAGE e posteriormente sondada para pureza por manchas ocidentais (Figura 1). Um controle importante é o uso de um anticorpo diferente durante a mancha ocidental para verificar a retirada da proteína correta. A combinação de IP com técnicas subsequentes é uma poderosa ferramenta de análise. O objetivo após a purificação pode ser a caracterização da proteína em si por NMR, espectrometria de massa e ensaios in vitro, ou análise dos parceiros interativos da proteína (ou seja, proteína, DNA, RNA) (3, 4, 5).
Figura 1: Visão geral do Procedimento de Imunoprecipitação. Imunoprecipitação é o isolamento de uma proteína específica usando um anticorpo. Após a produção de lise das células, há dois passos principais: pré-limpar e puxar para baixo. Durante a etapa de pré-limpeza, os lysatos celulares são pré-eliminados de proteínas que se ligam a anticorpos não especificamente usando um anticorpo de controle de isótipo. Em pull down step, a proteína alvo é puxada para baixo usando um anticorpo específico de proteína. A proteína isolada é então analisada pela mancha ocidental. Anticorpos isótipos e anticorpos específicos de proteína têm o mesmo domínio constante, mas diferentes domínios de ligação de antígeno. Um componente-chave deste protocolo são as contas de agarose da Proteína A/G que ligam o domínio constante dos anticorpos, permitindo a imunoprecipitação da proteína alvo. Clique aqui para ver uma versão maior desta figura.
Os anticorpos são o componente-chave de uma imunoprecipitação que o diferencia de outras formas de purificação de proteínas (ou seja, purificação da coluna de afinidade de níquel). Anticorpos são moléculas feitas por células B que podem reconhecer epítopos proteicos específicos. Os anticorpos possuem dois domínios: ligação constante (Fc) e antígeno (Fab) (Figura 1). O domínio constante identifica o tipo de anticorpo e dita a função in vivo. Normalmente, os domínios constantes dos anticorpos usados para IP são mouse, rat ou rabbit IgG. A porção de ligação de antígeno do anticorpo reconhece um epítope específico de uma proteína específica. Anticorpos podem reconhecer epítopos em proteínas dobradas que podem não existir quando a proteína é desnaturada e vice-versa. Portanto, a disponibilidade do epítope depende da dobra de proteínas – identificando um fator importante a considerar na escolha de anticorpos e condições para IP.
Ambos os sistemas procarióticos e eucarióticos têm proteínas de ligação de anticorpos. Nos sistemas eucarióticos, o objetivo é a proteção imune contra bactérias enquanto em sistemas procarióticos, o objetivo é a proteção do sistema imunológico. Proteínas de ligação de anticorpos afetam a metodologia IP de duas maneiras. Em primeiro lugar, há uma etapa de pré-compensação necessária (Figura 1) para livrar o lise de proteínas que ligam anticorpos – reduzindo assim a vinculação não específica no produto final. Esta etapa usa um anticorpo isótipo que tem o mesmo domínio constante que, mas um domínio de ligação de anticorpos diferente do seu anticorpo específico para proteínas. Proteínas bacterianas de ligação de anticorpos são o segundo componente-chave deste método. Depois que o anticorpo específico da proteína liga a proteína alvo, o anticorpo: complexo proteico deve ser puxado para baixo (Figura 1). Proteínas A, G e L são proteínas bacterianas que ligam o domínio constante dos anticorpos. Embora as bactérias usem isso para subverter o sistema imunológico, os pesquisadores têm cooptado este sistema para uma fácil purificação de anticorpos, e ele é usado durante os passos pré-clareira e pull-down. Essas proteínas têm diferentes afinidades de ligação para diferentes espécies e diferentes subtipos de domínio constantes – outro fator a considerar ao escolher condições para IP. Muitas empresas vendem contas de agarose rotuladas protein A/G (Figura 1), colunas de spin pré-feitas ou resinas para fazer colunas. Em geral, contas e colunas de spin são usadas para tamanhos amostrais menores, enquanto resinas são usadas para purificação a granel.
Neste exercício de laboratório, demonstramos como purificar a proteína endógena c-myc, dos timócitos murinas primários, utilizando contas de proteína A/G Plus baseadas em imunoprecipitação básica. O protocolo começa a partir da preparação de lisecelulares e termina com a verificação de retirada de proteínas bem sucedidas usando a análise de manchas ocidentais.
Procedure
Results
The results of the procedure detailed above are shown in Figure 2. From left to right, the lanes contain the control group (isotype), the test group (c-myc), the pre-cleared lysate (lysate), and the molecular weight ladder (ladder). The 25 and 75 kDa ladder bands are marked. The two prominent bands at ~25 kDa and 50 kDa are the light and heavy chain of the binding antibody, respectively and are non-specific to the IP or the samples. c-myc protein which runs around 67kDa on Western blots and is usually visible just below the 75 kDa ladder band. In this blot, the c-myc band is visible in the second lane, but absent in the first lane, indicating that the IP antibody successfully pulled down c-myc. There is no visible band in the pre-cleared lysate lane, suggesting that this protein has low endogenous expression levels.
Figure 2: Results of a Western Blot Analysis, used to assess the purification of c-myc by immunoprecipitation. A band at 67 kDa, corresponding to c-myc, is visible in the anti-c-myc lane, but not the isotype control lane. Note that c-myc levels were not high enough to be visualized in the lysate lane. Please click here to view a larger version of this figure.
Applications and Summary
In short, immunoprecipitation is the isolation of a specific protein using an antibody. In this example, the results of the immunoprecipitation were analyzed by Western blot to assess the purity. The isolated protein could be used in a number of applications afterwards including: NMR for protein structure, Mass Spectrometry for amino acid sequence, or in vitro assays for enzymatic characterization. IPs can also characterize the interacting partners of proteins. For instance, following isolation, DNA or RNA could be isolated for sequencing. Co-immunoprecipitations assess protein-protein interactions. When the target protein is pulled down during an IP, interacting proteins can also be pulled down. These interacting partners can be assessed by mass spectrometry and Western blot. Immunoprecipitation is a powerful technique for studying protein biology.
References
- Olliver, C. L. and Boyd, C. D. (1984). Immunoprecipitation of In Vitro Translation Products with Protein A Bound to Sepharose. In J. M. Walker (eds), Nucleic Acids. Methods in Molecular Biology (pp. 157-160). New Jersey: Humana Press.
- Thurston, C. F. and Henley, L. F. (1988). Direct Immunoprecipitation of Protein. In J. M. Walker (eds), New Protein Techniques. Methods in Molecular Biology (pp. 149-158). New Jersey: Humana Press.
- Anderson, N. G. (1998). Co-immunoprecipitation: Identification of Interacting Proteins. In R. A. Clegg (eds), Protein Targeting Protocols.Methods in Molecular Biology (pp. 35-45). New Jersey: Humana Press.
- Jackson, D. I. and Dickson, C. (1999). Protein Techniques: Immunoprecipitation, In Vitro Kinase Assays, and Western Blotting. In P.T. Sharpe and I. Mason (eds), Molecular Embryology. Methods in Molecular Biology (pp. 699-708). New Jersey: Humana Press.
- Trieu, E. P. and Targoff, I. N. (2015). Immunoprecipitation: Western Blot for Proteins of Low Abundance. In B. Kurien and R. Scofield (eds), Western Blotting. Methods in Molecular Biology (pp. 327-342). New York, NY: Humana Press.
Transcript
Immunoprecipitation, or IP, is a widely used technique to isolate a protein of interest from a cell or tissue lysate or a body fluid for protein characterization or to investigate protein-protein interactions.
The process begins with an antibody, which has a high affinity and specificity for the target protein. This antibody is mixed with the sample, allowing antibody-target complexes to form. Any protein bound to the target protein also gets indirectly attached to the antibody in the process. Next, the solution is incubated with agarose beads, conjugated to a bacterial protein, which has a strong affinity for the constant region of antibodies. The bacterial protein binds to the antibody and connects the antibody- target complexes to the beads. Then, the solution is centrifuged to precipitate the beads, thereby extracting the entire complex containing the binding antibody, the target protein, and any interacting proteins. Finally, the bound proteins are extracted from the beads and released from each other and are used for further analysis by techniques such as Western blotting.
Several variations of different parts of this technique are commonly used, like pre-clearing, using peptide tags or magnetic beads, or analyzing other non-protein binding partners. IP can be preceeded by a pre-clearing step, to remove non-specific antibody-binding proteins in the sample and minimize background. This involves first incubating the sample with isotype control antibodies, allowing them to bind to these proteins, and then using agarose beads to precipitate the complexes. The sample is then ready to proceed to the actual IP.
Peptide tags are useful if a specific antibody is not available for IP. Here, the target protein can be genetically modified to contain a peptide epitope tag and an antibody against the tag is able to pull out the protein of interest. Magnetic beads are often used instead of agarose to precipitate the target. After binding to the antibody-target complex, the sample tube is placed in a strong magnetic field, which extracts the beads from the solution. This eliminates the need for centrifugation and improves speed and convenience.
Immunoprecipitation is also used for studying DNA or RNA binding proteins and are known as chromatin immunoprecipitation and RNA immunoprecipitation, respectively. These variations are useful for troubleshooting and adapting the method for different experimental applications. In this video, you will observe how to pre-clear a cell lysate and perform immunoprecipitation to extract a protein of interest, followed by Western blot analysis to validate the experiment.
To begin, place the pre-collected cells in a microcentrifuge and spin at 13 thousand rpm for three minutes. Following the spin, remove the supernatant and then resuspend the cells in 500 microliters of lysis buffer RIPA with PMSF. Now, disrupt the cells using a few quick pulses with a vortex and then aspirate the lysate a few times with a 25 gauge needle attached to a syringe, taking care to avoid creating bubbles. Place the cells on ice for 15 minutes. After incubating the samples on ice, centrifuge the lysate for 15 minutes at four degrees celsius.
Label a new 1.5 milliliter microcentrifuge tube. Following the spin, transfer the supernatant to the freshly labeled tube and discard the pellet. Next, pre-clear the lysate of contaminants that bind non-specifically to either the agarose beads or the primary antibody by adding 20 microliters of the Protein A/G PLUS-agarose beads and one microgram of an isotype control antibody to the lysate, which in this example is a mouse IgG1 isotype control. Incubate the tube on a rotator in a cold room for 30 minutes. After rotating the lysate in the cold room for 30 minutes, centrifuge the sample at 3200 rpm for 30 seconds at four degrees celsius. Remove the tube from the centrifuge and transfer the pre-cleared supernatant to a fresh labeled 1.5 milliliter microcentrifuge tube. Discard the pellet.
Now, determine the protein concentration of the cell lysate by performing a Bradford assay. Label seven 1. 5-milliliter microcentrifuge tubes one through six and sample and aliquot 1000 microliters of the Bradford reagent into each tube. Six of the tubes will be used to make a standard curve by adding various amounts of known quantities of BSA to each tube. The amounts to add are listed in this table. In the seventh sample tube, add one microliter of the pre-cleared lysate. Place 200 microliters from each of the seven tubes into individual wells of a flat-bottom 96-well plate, repeating each sample in triplicate so that there are three columns of seven samples. Read the plate on a plate reader, using a wavelength of 595 nanometers. After creating a standard curve in Excel, calculate the protein concentration of the pre-cleared lysate.
Next, label two 1.5-milliliter microcentrifuge tubes- one as control and the other as test, which in this example, will be the c-myc antibody. Place 500 micrograms of the pre-cleared lysate into each of these tubes and then bring the total volume for each tube up to 500 microliters using lysis buffer. Next, add two micrograms of the anti-c-myc antibody to the test group tube. For the control, add two micrograms of the mouse IgG1 isotype control antibody. Once the antibodies are added to the tubes, place the samples on a rotator in a cold room and incubate for two hours. Now, add the agarose beads. To do this, it is recommended to cut off the end of a pipette tip and then, using this modified tip, add 200 microliters of the Protein A/G PLUS-agarose beads to each tube. Incubate the tubes on a rotator in the cold room overnight.
Following the incubation, remove the tubes from the rotator and spin the lysates in the microcentrifuge to pull down the beads. After the spin is complete, remove the tubes from the centrifuge and aspirate the supernatant from each tube. Next, wash the beads using 500 microliters of 1X Dulbecco’s PBS. Place the tubes in a microcentrifuge and spin down for 30 seconds at four degrees celsius. Following this, remove the supernatant. Repeat the wash and centrifuge steps one more time for a total of two times. Remove the tubes from the microcentrifuge and aspirate the buffer from each tube. Using gel loading tips, remove any left over buffer from the beads, keeping the beads on ice to elute the bound protein.
In this example, the protein is eluted into SDS-PAGE running buffer by boiling for Western blot analysis. To do this, resuspend the beads in 20 microliters of SDS-PAGE loading dye containing beta-mercaptoethanol, or BME. Boil the samples at 95 degrees celsius for five minutes to dissociate the immunocomplexes from the beads. Then, centrifuge the beads at maximum speed for 10 seconds at room temperature. Remove the tubes from the microcentrifuge and hold them in a rack at room temperature. Using gel loading tips, carefully pipette the samples from the beads and load them into wells of a 4 to 15% gradient SDS-PAGE gel. In addition to the samples, load a lane with a protein ladder as well as a lane with the pre-cleared lysate to serve as a loading control. Once the gel is loaded, run the gel at 100 volts.
After the dye front has reached the bottom of the gel, which should take approximately one hour, stop the gel and make a Western blot sandwich, ensuring that the PVDF membrane is between the gel and the cathode. Place the Western blot sandwich in the transfer apparatus and transfer the proteins on the gel to the membrane for one hour at 100 volts. After the transfer is complete, place the membrane in five milliliters of block to prevent the antibodies from binding non-specifically to the membrane. Rock at a low setting for an hour at room temperature. When the timer sounds, remove the blocking buffer. Add five milliliters of the blocking buffer with the detection antibody to the membrane. Here, an anti-c-myc antibody, that is different than the one used for the pull down, is used.
Incubate the blot over night, at four degrees celsius on a rocker at a low setting. Following the incubation, remove the antibody and blocking buffer. Wash the blot, using five milliliters of TBST for five minutes at room temperature, on a rocker at a low setting. This wash step should be repeated two to five times for a total of three to six washes, using fresh TBST for each wash. Add five milliliters of one to 1000 secondary antibody and blocking buffer to the blot. In this case, the secondary antibody is HRP-tagged anti-rabbit light chain. Incubate the blot on a rocker at a low setting for one our at room temperature. Next, remove the buffer and wash the blot with five milliliters of TBST. Incubate this wash on a rocker at a low setting for five minutes at room temperature. Repeat this wash for a total of six to 12 washes, each with a fresh five milliliters of TBST. Remove the final wash by first pouring the liquid off of the blot. Then, using tweezers, dab the edge of the blot on a laboratory wipe to remove any excess liquid and then place the blot in a fresh container. Next, cover the blot with 1X Chemiluminescent Detection Reagent and incubate for one minute.
Working quickly, dab the edge of the blot on a laboratory wipe to remove any excess detection reagent and then place the blot on the imaging surface of the Imager tray. Image using the Chemiluminescent program to capture multiple time points from 10 to 30 seconds. After the blot is imaged, choose an image with optimal band visibility and then export that image. Prior to moving the blot, use the Imager to take a picture of the blot to capture the location of the ladder. Then, export that image also. Finally, using a slide preparation software, such as PowerPoint, align the bands and ladder images to form one image.
This image shows the Western blot result for immunoprecipitation of the protein c-myc from thymocyte cells. From left to right, the lanes represent the isotype control, the c-myc IP, and the pre-cleared lysate input. The lane on the extreme right is a merged image of the molecular weight ladder. The strong band, at around 25 kilodaltons is from the light chain and the one at 50 kilodaltons is from the heavy chain of the binding antibody and are non-specific to the IP or the samples. C-myc runs around 67 kilodaltons on Western blots and is usually visible just below the 75 kilodalton ladder band. In this blot, the c-myc band is visible in the second lane but absent in the first lane, indicating that the IP antibody successfully pulled down c-myc. There is no visible band in the pre-cleared lysate lane, suggesting that this protein has low endogenous expression levels.