SDS Lab 03

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Miller-Motte Technical College, Cary *

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Biology

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Feb 20, 2024

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1 EXERCISE 5: Separation and detection of proteins I. OBJECTIVES A. To separate red blood cell cytoskeletal proteins on an SDS-PAGE gel, B. To visualize these proteins with Coomassie Brilliant Blue staining, C. To transfer these proteins to a nitrocellulose membrane for subsequent immunoblot detection (Western blot), D. To probe for albumin by immunoblot (Western blot). II. BACKGROUND: ENRICHING FOR AND DETECTING THE PROTEIN OF INTEREST A. Introduction The thousands of different proteins found in cells play important roles in most cellular functions including metabolism, interactions between cells, and structural support. As scientists, we often wish to study specific proteins to understand their functions within the cell. Tools for identifying and visualizing proteins prove invaluable to these efforts. The laboratory exercises that you complete over the next two weeks will help you explore some of the possibilities and limitations of working with proteins. Parts B and C (described below) have been completed for you due to time constraints. To understand the parts of this exercise that you complete, however, you need to understand what we did on your behalf. B. Isolating Red Blood Cells For this lab, we started with sheep’s blood. Whole blood consists of red blood cells, white blood cells of various kinds, platelets, and plasma. All of these components have large protein pools, but we are only interested in red blood cell cytoskeletal proteins. The first step required for preparing this lab, then, was to isolate red blood cells. The different densities of the various blood components provided a mechanism for doing so. In the presence of an anti-coagulant, tubes containing whole blood were

2 spun rapidly in a centrifuge. Centrifugation exerts strong forces on samples placed in the centrifuge. The sheep’s blood was exposed to forces on the order of 15,000 times the force of gravity for about 30 minutes. This separated the whole blood into layers based on the densities of the various blood components. This process is called differential centrifugation. The blood layer with the greatest density contained the red blood cells. To isolate red blood cells, we discarded the upper layers and washed the red blood cells several times. Because of its abundance, a 68 kD (kiloDalton) plasma protein called albumin likely contaminates the preparation. We will measure this contamination during the second week of this exercise. C. Isolating Membrane Proteins The proteins we will examine are the cytoskeletal proteins. These give the cells their shape and play roles in transporting material within cells. Recognizing that all of the cytoskeletal proteins are linked directly or indirectly to the plasma membrane, we purified our protein pool by isolating just the plasma membrane ‘fraction’ of the cell. The process we used, called fractionation, began with lysing (i.e. breaking open) the cells. Cells can be lysed using various methods including detergents, mechanical disruption, and osmotic shock. To remove the cell contents without breaking up individual organelles or the plasma membrane itself, we used the relatively mild osmotic shock method. Placing the cells in a hypotonic buffer caused the cells to swell until the membranes developed holes. (You should be able to explain why this happened. Review osmosis in your textbook if this is unclear.) The holes in the plasma membrane allowed the cellular contents to flow out, without removing plasma membrane associated proteins or lipids. Cells prepared in this way are referred to as “ghosts” because they retain a cellular shape but are empty. Isolating the ghosts away from the organelles involved differential centrifugation. The washed red blood cell ghosts comprise the sample that you will work with in lab.

3 D. Size Separation of Proteins SDS-PAGE (S odium d odecyl s ulfate p olya crylamide g el e lectrophoresis) is a technique which allows us to separate proteins based on molecular size. The initial difficulty lies in the fact that proteins are composed of different amino acids. Each amino acid has a characteristic three-dimensional shape and charge (negative, positive or neutral). Linking these amino acids together covalently in a polypeptide gives each protein a characteristic three-dimensional shape and charge. This raises the question of how SDS-PAGE uses an electric current to separate proteins only on the basis of size. The answer involves the methods used to prepare the samples for SDS- PAGE. Sodium dodecyl sulfate (SDS) is a strong anionic detergent that destroys all secondary and tertiary protein structure by disrupting non-covalent interactions (hydrophobic interactions, ionic bonds, and van der Waals forces) within a protein. Second, the protein samples are heated to 100° C which disrupts hydrogen bonding. Finally, a reducing agent (dithiothreitol in our case) is necessary to break disulfide bridges within and between polypeptide chains that help maintain the protein folds. Preparing samples in this way linearizes polypeptide chains. SDS has an additional essential role of coating the linearized polypeptide with a uniform negative charge, overwhelming the charge differences due to the amino acid charges. The size separation of proteins (or any other macromolecule) is accomplished by resolving them through a semi-solid matrix, which in our case is made of polyacrylamide. Migration of the negatively charged proteins through the polyacrylamide gel is accomplished by applying an electrical current. The denatured proteins are introduced to the gel at the cathode (negative pole) and as they are negatively charged due to the SDS, they will migrate towards the anode (positive pole). Smaller molecular weight proteins migrate quickly toward the anode, while larger proteins are retained near the top of the gel. The study of red blood cell membrane and cytoskeletal proteins will utilize 12% acrylamide gels. These gels resolve proteins in the molecular weight range between 15 and

4 80 kD. The gels you will use in class have been prepared for you. You will load your protein samples into the wells of the gel. After the gel is run, you will be looking in each lane (the long strip under the well) for the bands of protein. In order to determine the actual size of the proteins, we will be including a molecular weight marker, also called a size standard. The marker is a preparation of proteins of known sizes. After the gel is run, these proteins can be used as a ruler to determine the sizes of proteins from the experimental sample. For instance, a protein that resolves halfway between the 25 kD marker and the 32 kD marker is going to be ~ 29kD. Smaller (i.e. shorter) proteins migrate (or “run”) faster through the gel because the gel impedes their progress less than larger (i.e. longer) proteins. This means that the smallest marker will be farther down the gel and the largest will closer to the top. When analyzing your gel, notice that the markers are not equally spaced. Bands separate less near the top. Also be aware that sometime small proteins run off the bottom of the gel or are masked by the blue dye front. The proteins that you resolve by SDS-PAGE will be visualized using two different techniques. Coomassie Brilliant Blue binds to all polypeptides and will give the otherwise colorless proteins a blue color. This method of detecting proteins does not work for low abundance proteins. As described in the following section, you will also use a far more sensitive method for detecting specific proteins that involves antibodies. E. Immunodetection of Proteins A technique that is commonly used to identify the presence of a given protein within a preparation is called electroimmunoblot analysis (or Western blot analysis). Following SDS-PAGE, you will electrophoretically transfer the proteins to a nitrocellulose membrane. This membrane, or ‘blot’, provides a solid support for the proteins, whereas the acrylamide gel is both very delicate and prevents access to the proteins within. After transferring the proteins to the membrane, you will incubate it with an antibody that recognizes albumin. Antibodies are extremely specific for proteins and are produced by B-

5 Iymphocytes in the body in response to foreign proteins. Antibody binding to foreign proteins (antigens) targets the antigens for removal from the body. An antibody raised against a specific antigen is called the primary antibody . Albumin is the antigen for the primary antibody you will use. Primary antibodies have a variable region and a constant region . The variable region is what makes a particular antibody specific to a particular antigen. The constant region, on the other hand, is exactly the same for every antibody produced within an animal. Every antibody made in a rabbit, no matter what the variable region recognizes, will have an identical constant region. A primary antibody against albumin that is mixed in with our blots will bind exclusively to albumin through its variable region; the constant region will ‘stick out’ from the blot. Primary antibodies can be used alone (direct immunodetection), but generally another step using secondary antibodies is included. A secondary antibody is made by using the constant region of the primary antibody as the antigen for the secondary antibody. Use of the secondary antibody allows us to amplify the signal dramatically. For example, if 10 primary antibodies were able to stick to each strand of albumin, we would have 10-fold amplification of the signal. If 10 secondary antibodies stuck to each primary, we would have 100-fold amplification. Finally, we need some method to visualize this protein-antibody complex. The secondary antibody can be detected by one of several methods. The secondary can be tagged radioactively, fluorescently, with a gold bead (for electron microscopy), enzymatically. Enzyme tags (e.g. alkaline phosphatase or horseradish peroxidase) convert a colorless solution to a colored precipitate. Antigen detection using a labeled secondary antibody is termed indirect immunodetection. After washing off excess anti-albumin antibodies (primary antibody), you will add a secondary antibody (against the constant portion of the anti-albumin antibody) that is complexed with horseradish peroxidase. This enzyme, in the presence of the chemical H 2 0 2 (hydrogen peroxide) and the chromophore 4-chloro-napthol, forms a purple precipitate identifying the presence of albumin on the nitrocellulose membrane.

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