Sunday, 27 October 2019

Protein Techniques

Protein Techniques


As with the other appendices, it should be noted that the information presented here merely aims to assist the reader with the text and the end- of-article questions; there are no laboratory protocols.

SDS- PAGE

SDS- PAGE

SDS- PAGE for stands the Sodium Dodecyl Sulfate PolyAcrylamide Gel Electrophoresis. As with DNA agarose gel electrophoresis, in this process, proteins will be moving through polyacrylamide gels in an electrophoretic field (although nucleic acids can also be run in polyacrylamide gels instead of an agarose gel if precise separation is required). Ammonium persulfate (APS) and tetramethylethylenediamine (TEMED) are commonly used as polymerization stabilizers. SDS is used as a chemical denaturation agent, thereby unfolding the protein, and thus eliminating conformation- related mobility differences.

SDS also applies a negative charge to proteins, which is crucial to further eliminate mobility differences in proteins due to charges in the amino acid side chains. In addition to SDS, protein samples are usually also heated, which helps complete unfolding, and DTT (dithiothreitol) and/ or 2-mercaptoethanol are used to completely get rid of disulfide linkages or any remaining secondary structures. Therefore, SDS- PAGE can be used to separate proteins purely based on size (charge and conformation effects are eliminated).

To ensure that all proteins are well separated, a two-tier gel system is used—on the top tier there is a low percentage “stacking” gel, typically around 5 to 6% polyacrylamide, and on the bottom tier there is a higher percentage “separating” or “resolving” gel, typically between 9 to 20% polyacrylamide, depending on the size range of the proteins to be separated and studied (higher percentage gels are required for better separation of smaller proteins) (Figure .1). Visualization of proteins can be done using specific dyes such as Coomassie Brilliant Blue R-250 (commonly referred to as Coomassie blue) or silver stain.

SDS- PAGE
Figure .1 A basic scheme for SDS- PAGE for the size-based separation of proteins. See the text for details.

"Western Blotting"

Western Blotting

Western blotting or immunoblotting is a method used to identify specific proteins in a sample, using the antibody-antigen affinity principle (named after the Southern blot). Usually, proteins are separated by SDS- PAGE electrophoresis and thereupon transferred to a blotting membrane (which could be nitrocellulose, polyvinylidene fluoride, PVDF, or similar) using again an electrical field for transfer (hence, called electroblotting) (Figure .2). There are wet blots, semi-dry blots, or dry- blots from different commercial companies, all of which could be used according to the manufacturer’s instructions and based on personal preference and experience.

Detection is done using a combination of primary and secondary antibodies, as shown in the article (also see Figure .2). After incubation with primary and secondary antibodies, detection is usually done through an enzymatic or biochemical reaction involving the conjugate molecule of the secondary antibody (Figure .2). The most common enzyme conjugates used with secondary antibodies are the alkaline phosphatase (AP) and the horseradish peroxidase (HRP), although many more are available.


Western Blotting


Figure .2 A basic schematic of the Western blotting principle. Proteins separated by SDS- PAGE are electroblotted onto an appropriate blotting membrane, after which the blot is first blocked using BSA, nonfat dry milk or similar, so as to avoid any nonspecific binding of primary antibody to random proteins. After washing with a phosphate-buffered, saline-based, or similar wash solution, the blot is immediately incubated with the appropriate primary (1°) antibody that is specific to the target protein. Again, after a wash step, the blot is then incubated with the appropriate secondary (2°) antibody that recognizes the 1° antibody and is conjugated (typically) with an enzyme, which can then be used to detect the signal through electrochemiluminescence or a color reaction.

AP is a typical phosphatase enzyme that removes phosphate groups from substrates. The most common AP substrate used in Western blots is the 5-Bromo-4-chloro-3-indolyl phosphate (BCIP) and nitroblue tetrazolium (NBT), which are processed by AP to yield an indigoid dye (purple) and an insoluble formazan precipitate (blue), respectively, in colorimetric reactions. HRP, on the other hand, is a typical peroxidase that can be used both in colorimetric detection and chemiluminescent reaction: when 4-chloro-1-naphthol (4CN) and 3,3′-diaminobenzidine (DAB) are used as HRP substrates in Western blots, the enzyme converts them to an insoluble purple product and an insoluble brown precipitate, respectively. When HRP reacts with luminal, a chemiluminescent substrate, on the other hand, it will produce light, which can be detected and quantified by chemiluminescence detecting equipment available from various manufacturers. (Chemiluminescent substrates for AP are also available from some companies, however, HRP conjugate has been far more common in this detection method.)

2D Gel Electrophoresis and Proteomics

2D Gel Electrophoresis and Proteomics

Another analytical technique in biochemistry, which readers can find more detailed information about in the aforementioned classical biochemistry books, is 2- dimensional (2D) gel electrophoresis. The two dimensions are required for higher resolution separation of proteins: in the first dimension there is a separation of nondenatured proteins (based mainly on a charge, called isoelectric focusing), and in the second dimension there is a purely size-based separation in the denaturing gel (similar to SDS- PAGE).

The principle is this: in the first dimension, the isoelectric focusing separates proteins based essentially on a charge, through a gel that contains a pH gradient (which can be adjusted according to the protein groups to be studied). Proteins move along the gel to either the positive or negative end, until they reach their isoelectric point, at which point the net charge on the protein will be zero, and hence the proteins will no longer have mobility. After the first dimension, the gel (with different proteins settled at their respective isoelectric points) will be mounted onto an SDS- PAGE denaturing gel, whereby proteins from the first dimension will now be separated according to size, as described above, in this second dimension (Figure .3). The incorporation of the first dimension greatly improves resolution, both separating proteins of similar size but different amino acid compositions, but also separating proteins that are post-translationally modified (such as phosphorylated or acetylated proteins) with higher precision than standard SDS- PAGE gels.

The proteins are then commonly detected by silver staining, or Coomassie staining described above, appearing as spots on the gel, which are then analyzed with the help of specialized computer programs. The spots of interest can then be excised from the gel, and analyzed by mass spectrometry (Figure .3).

2D Gel Electrophoresis and Proteomics


Figure .3 A diagram showing the basic principle of 2D gel electrophoresis. See the text for details.

Proteomics is the large- scale and global analysis of (almost) all proteins (and every post-translational variant thereof) in a cell or tissue. A 2D gel electrophoresis– mass spectrometry combination is perhaps the most commonly used approach to study whole proteome profiles, although protein microarrays can also be used. However, since proteins have to be detected with antibodies (with current technologies), and since both proteins and antibodies can be denatured quite easily, affecting their interaction, protein arrays or protein chips are more difficult to optimize, and can thus not be routinely employed. Differential gel electrophoresis (DIGE), which uses Cy2, Cy3, and/ or Cy5 labeling of different protein samples, allows for higher sensitivity of the fluorophore-labeled proteins. Other biochemical techniques such as liquid chromatography followed by tandem mass spectrometry (LC/ MS/ MS) are also used in proteomics.

Mass spectrometry-based targeted proteomics, which can detect specific proteins of interest with high sensitivity and reproducibility, has in principle been around for quite some time, and yet because of its widespread use and the quantitative information recently made possible, it was chosen the “Method of the Year 2012” (Editorial 2013). Tissue proteomics, from formalin-fixed and paraffin-embedded (FFPE) tissues, analyzed by matrix-assisted laser desorption/ ionization (MALDI) MS imaging, has become popular, as imaging of hundreds of compounds on a tissue section is possible, linking histology with molecular biology and proteomics (Lonquespee et al. 2014).

Immunofluorescence and Immunohistochemistry

Immunofluorescence and Immunohistochemistry

Both these techniques rely on the detection of proteins in cells or tissues using antibodies (immuno-), either by fluorescent or enzyme conjugate on the secondary antibody, that is, either by fluorescent or light microscopy. In immunohistochemistry, -histo refers to the staining of tissues; if dispersed cells (devoid of any extracellular matrix) are stained, then the technique is also called immunocytochemistry (-cyto- refers to staining of individual cells).

The basis of both immunofluorescence and immunohistochemistry is in principle the same, using primary antibodies as probes to recognize and identify target proteins, and secondary antibodies (in indirect fluorescence) against the primary antibody to amplify the signal (Figure .4). The only difference is the cells or tissues being used, which will slightly change the protocol (depending on whether the extracellular matrix is around or not; for example, please refer to detailed protocols).

In essence, the basic logic is the same as in Western blots— one is trying to detect the expression of a protein. However, since cells are used, information about the subcellular localization of proteins can also be obtained in addition to the level of expression. (Note: The technique is not limited to proteins— any molecule against which a specific antibody is available can be studied). The first step is the fixation of cells or tissues onto the microscope slide, followed by permeabilization. This step is essential, not only because cells must remain on the slide at all times and not get washed away, but also so that antibodies against target molecules can penetrate through the cell membrane (and cell wall, if yeast, fungi, or plants are studied). Following fixation/ permeabilization, a blocking step is necessary, for the same reasons as in Western blotting (i.e., to prevent nonspecific binding of antibodies).

Once slide preparations are blocked and excess reagents are washed away, the slides are incubated with primary antibodies against (and specific for) the target protein. (Direct immunofluorescence, in fact, uses conjugated primary antibodies and directly prepares slides for detection; however, this is not employed frequently.) Following the primary antibody incubation, the slides are briefly washed and incubated with secondary antibodies against the primary. This secondary antibody contains either a fluorescent conjugate, such as fluorescein isothiocyanate (FITC), which emits green fluorescence when excited, or an enzyme conjugate, such as HRP, which can be used for colorimetric detection in light microscopy.

Immunofluorescence and Immunohistochemistry

Figure .4 A brief overview of the steps of immunofluorescence and immunohistochemistry. Two samples are shown here: either a protein’s amount and localization can be studied in cell A versus cell B, or the level of expression and localization of a protein can be studied under two (or more) different conditions (cond. A and cond. B). See the text for details.

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