Biochem 14.2 Blotting Techniques PDF

## Lesson 14.2: Blotting Techniques ### Chapter 14: Biomolecule Purification and Characterization ### Introduction Lesson 14.1 discusses gel electrophoresis as a way of separating a mixture of biomolecules based on some physical or chemical property, such as size or isoelectric point. The stains...

## Lesson 14.2: Blotting Techniques ### Chapter 14: Biomolecule Purification and Characterization ### Introduction Lesson 14.1 discusses gel electrophoresis as a way of separating a mixture of biomolecules based on some physical or chemical property, such as size or isoelectric point. The stains that detect these molecules, however, are generally nonspecific. In other words, Coomassie stains both the protein of interest and all other proteins in the sample. Similarly, ethidium bromide stains both the RNA or DNA of interest and all other nucleic acids in the sample. Although these stains can be useful when working with purified samples or monitoring the progress of a purification experiment, the nonspecific nature of the dyes makes it difficult to interpret data with impure samples, such as crude lysates. To visualize and analyze only a specific protein or nucleic acid of interest, blotting techniques have been developed that use stains that target only particular biomolecules, as shown in Figure 14.20. This lesson provides an overview of the principles of these blotting techniques and discusses in detail three blotting techniques that are likely to appear on the exam. * **Coomassie-stained SDS-PAGE gel:** All proteins are stained. Specific protein of interest is hidden by other proteins. * **Western blot:** Only protein of interest is stained. ### Figure 14.20: Blotting techniques allow for the visualization of specific biomolecules within a mixed sample ### 14.2.01: Principles of Blotting Techniques The generalized process of blotting involves the transfer of biomolecules from the sample to a membrane, followed by visualization of specific target molecules. Often, the sample had been previously processed through gel electrophoresis to allow for the separation of biomolecules by size (eg, Southern, northern, and western blots); however, blots can also be performed without electrophoresis, as discussed at the end of this concept. #### Transfer of Biomolecules to Membranes The membranes used with blotting techniques are "sticky" for their respective biomolecules. This means that once a biomolecule adheres to the membrane, it is immobilized and unlikely to diffuse or be washed away during the subsequent staining, blocking, and washing steps of the visualization procedure, discussed later in this concept. Blots for analyzing DNA or RNA often use a positively charged nylon membrane, which binds to the negatively charged sugar phosphate backbone of nucleic acids. In contrast, blots used to analyze proteins often use a nitrocellulose or polyvinylidene fluoride (PVDF) membrane, which binds proteins through a combination of hydrophobic and electrostatic interactions. Two common ways to transfer samples from an electrophoretic gel to a membrane are capillary transfer and electroblotting. These methods are illustrated in Figure 14.21. Capillary transfer is discussed in more detail with Southern blots in Concept 14.2.02, and electroblotting is discussed in more detail with western blots in Concept 14.2.04. ### Figure 14.21: Capillary transfer and electroblotting are two methods of transferring biomolecules to a membrane. ### Probes and Blocking To specifically stain a biomolecule of interest, the membrane must be incubated with probes that specifically bind to the target molecule, but not to any other molecules. Blots intended to visualize nucleic acids often use single-stranded DNA primers that hybridize with the target sequence. Blots intended to visualize proteins often use antibodies that specifically recognize the target protein. Note that these probes are the same type of molecule that the membrane is "sticky" to. In other words, DNA primers are nucleic acids and can nonspecifically stick to nylon; antibodies are proteins and can nonspecifically stick to nitrocellulose or PVDF. To prevent this, the membrane must be blocked prior to incubation with probes. Blocking involves incubating the membrane with a solution that contains nucleic acid or protein to saturate the nonspecific binding sites. By performing blocking prior to adding any probe, experimenters can ensure that the probe binds only to the target molecule, rather than binding the membrane itself. Ultimately, this helps minimize background noise and improves signal clarity. The effect of blocking for a western blot is shown in Figure 14.22. A common blocking reagent for Southern and northern blots is salmon sperm DNA; two common blocking reagents for western blots are purified solutions of bovine serum albumin (BSA) and fat-free cow's milk. ### Figure 14.22: Blocking reagents bind nonspecifically to blotting membranes, preventing nonspecific binding of the probe (eg, an antibody) to the membrane. This reduces background noise and improves signal detection. ### Visualization To determine both the location of the target molecule on the membrane and the relative amount of it, the probe molecule that binds to it must be labeled. This label can be incorporated either directly (ie, the label is covalently incorporated onto the primary probe molecule), or it can be indirect (ie, the label is covalently incorporated in a secondary probe that then binds the first probe). Indirect methods are discussed in more detail with western blots in Concept 14.2.04. Several types of labels are common in modern biology and biochemistry, including radioactive, chemiluminescent, and fluorescent labels. In radioactive labeling, certain atoms in the probe molecule are replaced with radioactive isotopes of that element. For example, the 31P atom in the sugar-phosphate backbone of DNA probes can be replaced with a radioactive isotope such as 32P. When the 32P isotope undergoes ẞ- decay (becoming 32S), it emits a high-energy particle that reacts with radiographic film (sometimes called x-ray film). Like photographs, the film can be developed to allow for visualization of the location and position of the radioactive labels on the blotting membrane; the developed signals appear as bands on the film. The detection of radioactive labels using radiographic film is known as autoradiography. An example of an autoradiograph is shown in Figure 14.23. ### Figure 14.23: Autoradiography uses radiographic film to capture radioactive decay of isotopically labeled probes. Once the film is developed, locations where decay was detected appear as dark bands. Unlike other labeling methods, radioactive labels have the benefit of causing minimal disruption to the chemical properties of the probe molecule. This greatly reduces the risk of any effect on binding interactions. However, radioactive labels do pose a danger to the experimenter due to radioactive emissions. To avoid these risks, chemiluminescent or fluorescent molecules may instead be linked to the probe molecule. These molecules can then be detected by film or specialized digital cameras, yielding similar results to autoradiography without releasing high-energy particles or photons. ### Loading Controls Blotting is considered a semiquantitative technique. Although blots are not generally used to calculate precise quantities or percent changes, they can indicate whether the expression of the target molecule has increased or decreased. This can be done by comparing the size and intensity of the bands between two experimental conditions, as in gel electrophoresis (Lesson 14.1). When using blots in a semiquantitative way, it is critically important to ensure that samples are loaded in a consistent manner. Differences in sample collection may result in inconsistent concentrations of total protein or total nucleic acid. If these differences are not accounted for, it is impossible to determine if an increased band size is due to an increase in expression or due to the experimenter loading a higher concentration of cell lysate material, as shown in Figure 14.24. ### Figure 14.24: Differences in blot band size may be due to an expression change or due to an error in sample collection and loading. To prevent this ambiguity, the contents of the cell lysate must be quantified prior to loading (biomolecule quantitation is discussed in Concept 14.4.02). Based on the results of the biomolecule quantitation, equal masses of the analyte (ie, protein, nucleic acid) can be loaded onto the gel or blot. Proper loading can be verified through the probing of particular molecules in addition to the target of interest. Tubulin, ẞ-actin, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) are some commonly probed additional targets because they are expressed by all cells and because expression levels are typically unaffected by experimental conditions. Consequently, the genes that express tubulin, ẞ-actin, GAPDH, and other similar proteins are often called housekeeping genes. When interpreting a change in band size for a target of interest, it is common practice to compare it to an unchanged band size for the housekeeping gene (or gene product). This serves as a **loading control** and ensures any change in the target protein is due to changes in the experimental conditions and not due to sample collection errors. The use of loading controls is shown in Figure 14.25. ### Figure 14.25: The use of loading controls helps ensure interpretation of a scientifically valid result. ### Concept Check 14.5 An experimenter is about to load an SDS-PAGE gel in preparation for a western blot. She is preparing her samples and has 10 µL of crude lysate available per gel lane. She quantifies her two samples of interest and finds Sample A has a total protein concentration of 2 mg/mL and Sample B has a total protein concentration of 4 mg/mL. If the experimenter wants to maximize the total amount of protein loaded per well, how many microliters of Sample A should she add to the Sample A gel lane? How many microliters of Sample B should she add to the Sample B gel lane? ### Solution ### Nonelectrophoretic Blots: The Dot Blot The following concepts in this lesson discuss the most commonly used blotting methods, which all follow a gel electrophoresis procedure. However, blots can also be performed without first running an electrophoretic gel. These blots are known as dot blots. Although dot blots are less informative because of the lack of electrophoresis data (eg, molecular size), the presence of a signal and the relative signal intensities can still provide valuable information in a quicker and less labor-intensive experiment. Because dot blots do not involve gels, the transfer process simply involves spotting dissolved sample onto the blotting membrane, similar to spotting thin-layer chromatography (TLC) plates (see Organic Chemistry Lesson 13.3). The spots can air dry, or the solvent can be removed through vacuum filtration. Either way, the target molecule adheres to the dry membrane, leaving "dots" that can be visualized through one of the methods described previously (Figure 14.26). ### Figure 14.26: A dot blot is an example of a nonelectrophoretic blotting technique. ### 14.2.02: Southern Blots The first of the electrophoretic blotting methods is the Southern blot. The Southern blot is named after its developer, Edwin Southern, and is used to detect DNA. The Southern blotting method begins with agarose gel electrophoresis (Lesson 14.1) to separate DNA fragments based on their size. DNA agarose gels are typically run in native conditions, which keeps the DNA double stranded. This double-stranded DNA must be denatured before it is transferred to a membrane so the probes can base pair with the denatured single strands. Typically, denaturation is done by incubating the gel in an alkaline (ie, high-pH) solution. Once the DNA has been denatured into single strands, it must then be transferred from the gel to the blotting membrane. The classical way to perform transfers from agarose gels is through upward capillary transfer. In upward capillary transfer, shown in Figure 14.27, the membrane is placed on top of the gel, and the gel-membrane pair is sandwiched between two sheets of wet filter paper to prevent either from drying out. The bottom piece of filter paper is connected to a reservoir tank filled with buffer; the upper piece of filter paper is overlaid with dry paper towels. A weight is placed upon the whole stack to add pressure, which maintains contact between the membrane and the gel. ### Figure 14.27: In upward capillary transfer, single-stranded nucleic acids are carried upward by buffer, transferring them from an agarose gel to a nylon membrane. The dry paper towels near the top of the stack draw buffer up toward them, resulting in buffer flow from the tank to the lower filter paper and then through the entire setup. As the buffer moves upward, it also carries the DNA in the gel upward. The DNA stops migrating once it adheres to the "sticky" nylon membrane. Once the DNA is immobilized on the membrane, the membrane must be blocked (see Concept 14.2.01) before being incubated with probe. The oligonucleotide probe has a sequence that is the reverse complement of a specific region in the target sequence, allowing the probe to base pair with the target DNA. This annealing of probe to target DNA is called hybridization. After washing off unbound probe, the hybridized target-probe pair can then be visualized as described in Concept 14.2.01. An overview of the Southern blot procedure is shown in Figure 14.28. ### Figure 14.28: Overview of the Southern blot procedure. Southern blots can be used in genomic analysis, such as testing for gene duplication, deletion, or rearrangement events. Southern blots can also be used to examine smaller point mutations (eg, insertions, deletions, substitutions); however, PCR-based methods (see Biology Lesson 4.1) are much more common for this purpose in modern biochemistry. ### 14.2.03: Northern Blots Unlike the Southern blot, which targets DNA, the second electrophoretic blotting method targets RNA. This blot is not named after a scientist; however, the direction-based naming system was kept by naming the RNA blot a northern blot. The process of northern blotting is very similar to the process of Southern blotting because both blots target nucleic acids. However, there are some important differences. Although RNA is not typically double stranded, it can still have local regions of secondary structure due to internal base pairing, which means that RNA still needs to be denatured. Because RNA is much more reactive than DNA, alkaline conditions (which would hydrolyze RNA) cannot be used. Instead, regions of RNA base pairing are denatured with a chemical denaturant such as formaldehyde. To prevent these structured and folded regions from affecting electrophoretic migration, formaldehyde is also included in the agarose gel and buffer during electrophoresis, in which it acts similarly to the denaturing effect of SDS in SDS-PAGE gels. The electrophoresed RNA is then transferred, hybridized, and visualized in a manner similar to that of Southern blots (Concept 14.2.02). An overview of the northern blot procedure is shown in Figure 14.29. ### Figure 14.29: Overview of the northern blot procedure. Northern blots can be used to analyze expression and sequence various RNA subtypes (eg, mRNA, tRNA, miRNA) and can be used in combination with PCR-based methods. ### 14.2.04: Western Blots The third electrophoretic blotting method targets proteins. To maintain the directional theme, but to distinguish itself from the north-south nucleic acid axis, protein blots are called western blots. Like Southern and northern blots, western blots begin with running an electrophoretic gel. In the case of western blots, this is typically a polyacrylamide gel. Instead of transferring to the membrane via upward capillary motion, protein gels are typically transferred using an electric field (electroblotting). As with gel electrophoresis (Lesson 14.1), negatively charged proteins travel toward the positively charged anode of an electrolytic cell. If the protein gel was run with SDS (as in SDS-PAGE), typically enough SDS remains bound to protein to maintain this directional transfer. Proteins in native gels require additional treatment, either during or after electrophoresis, to ensure the proteins migrate toward the anode during transfer to the membrane. A nitrocellulose or PVDF membrane is placed on the anode-facing surface of the gel, and proteins bind to this membrane as they leave the gel, stopping their migration. A typical electroblotting setup is shown in Figure 14.30. ### Figure 14.30: A wet tank electroblotting setup. Like the upward capillary transfer method described in Concept 14.2.02, contact between the gel and the membrane is maintained, in this case helped by the pressure from the foam pads (or sponges) in a closed cassette. Unlike upward transfer, electroblotting requires electric power to drive molecular movement. This process is faster than capillary transfer but also generates much more heat. Because polyacrylamide gels are much thinner than agarose gels, heat buildup is less of an issue; however, transfers are still usually performed with ice blocks or in a refrigerated room to minimize heating. Once protein transfer to the membrane is complete, the membrane must be blocked (typically with a solution of bovine serum albumin [BSA] or with fat-free cow's milk) before being incubated with probe. In western blots, the probe is usually an antibody that recognizes the target protein as its antigen. The antibody that directly binds the target protein is known as the primary antibody. The primary antibody may be directly (ie, covalently) linked to a label for visualization, or another antibody (a secondary antibody) may be used. The secondary antibody recognizes and binds to the primary antibody. Typically, a secondary antibody binds any primary antibody produced by a specific organism (eg, mouse, goat). If a secondary antibody is used, it serves as the molecule that the label is linked to. Primary and secondary antibodies are shown in Figure 14.31. ### Figure 14.31: Primary and secondary antibodies. The use of secondary antibodies brings many advantages, the first of which is cost reduction. Chemically linking target-specific antibodies to labels is costly, labor intensive and may produce low yields. By using mass-produced secondary antibodies, different specialized primary antibodies produced by the same species can be used in a variety of experiments (including immunohistochemistry, immunofluorescence, immunoprecipitations, and ELISAs, in addition to western blots), and each can be detected by the same secondary antibody. Second, the use of secondary antibodies can improve signal detection. This is because it is possible for multiple secondary antibody molecules to bind to a single primary antibody molecule, resulting in multiple labels per primary antibody. The potential for multiple secondary antibodies to bind is due partly to the dimeric nature of antibodies and partly due to the use of polyclonal secondary antibodies (ie, different antibodies in the same mixture that bind different epitopes on the target molecule). ### Visualization of Antibody Labels For visualization, the relevant antibody is typically linked either to an enzyme or to a fluorescent molecule (a fluorophore). The enzyme (typically horseradish peroxidase) can be used to catalyze either a chemiluminescent or a chromogenic reaction, whereas the fluorophore can be visualized spectroscopically. An overview of the western blotting process is illustrated in Figure 14.32. ### Figure 14.32: Overview of the western blot procedure.