Gel Electrophoresis - Biochemistry PDF

## Gel Electrophoresis ### Introduction This book has presented several principles of biochemistry: the nature of amino acids, their formation into proteins, the ability of many proteins to act as enzymes, structural features of various biomolecules, and metabolic reactions. These principles were...

## Gel Electrophoresis ### Introduction This book has presented several principles of biochemistry: the nature of amino acids, their formation into proteins, the ability of many proteins to act as enzymes, structural features of various biomolecules, and metabolic reactions. These principles were not always known, however, and had to be determined experimentally. This chapter presents some of the techniques used to probe biomolecules. In particular, it explores several techniques used to purify and characterize proteins and other biomolecules. One of the most important tools in biochemistry is electrophoresis. This technique includes a broad range of methods that separate biomolecules based on their size, electric charge, and other characteristics by generating an electric field that forces molecules to migrate through a gel. This lesson discusses multiple types of electrophoresis and the principles that govern this technique. ### 14.1.01 Principles of Electrophoresis Broadly speaking, electrophoresis involves the use of an electric field to cause charged particles to migrate through a gel or some other substance. In a biochemical context, electrophoresis is used to separate various charged particles (typically proteins, DNA, or RNA) from each other. Separation occurs because different proteins (or different DNA or RNA molecules) have different abilities to migrate through the gel. The most common substances used to make gels for electrophoresis are agarose (a complex carbohydrate) and polyacrylamide (an organic polymer). Both substances can form a porous matrix. Larger molecules migrate through the matrix more slowly than smaller molecules because it is more difficult for larger molecules to navigate the pores, as shown in Figure 14.1. **Figure 14.1** Relative motion of large and small particles through a gel in an electric field. * Highly crosslinked gel matrix hinders movement of molecules * Positive pole of electric field attracts negatively charged molecules to end of gel * Large molecules get tangled in matrix and move slowly * Small molecules can navigate matrix more easily In addition, molecules with a higher magnitude of charge (positive or negative) experience a greater force when exposed to an electric field, so they move more quickly through the gel. Accordingly, electrophoresis may separate molecules by size, charge, or both. A gel typically consists of a relatively small percentage of the matrix-forming agent (agarose or polyacrylamide) and a much larger percentage of aqueous solution (eg, a Tris-HCI buffer adjusted to a desired pH). Varying the percentage of agarose or polyacrylamide alters the size of the pores within the gel. For example, a 10% polyacrylamide gel forms smaller pores than a 5% polyacrylamide gel. Typically, agarose gels form larger pores than polyacrylamide gels. Consequently, agarose gels are most often used to separate larger molecules (eg, large proteins, DNA), and polyacrylamide gels are used to separate smaller proteins. Most proteins are small enough that they can be separated by polyacrylamide gels. For this reason, protein separation is often carried out by polyacrylamide gel electrophoresis (PAGE), whereas DNA and RNA separations are more commonly carried out by agarose gel electrophoresis. These gels are shown in Figure 14.2. **Figure 14.2** Polyacrylamide and agarose gels, and the molecules they separate by electrophoresis. * Polyacrylamide gel: Smaller pores accommodate small to large protein. * Agarose gel: Larger pores accommodate extremely large protein, RNA, and DNA. Electrophoresis occurs in an electrolytic cell. Unlike galvanic cells, electrolytic cells use a power source to drive electrons toward the negative electrode, where they accumulate and are eventually gained by electrolytes, causing a reduction reaction. By definition, reduction always occurs at the cathode (see General Chemistry Chapter 9). Therefore, in electrolytic cells, the negatively charged electrode (ie, the electrode that the power source moves electrons to) is the cathode and the positively charged electrode is the anode. Once the current is applied, negatively charged particles within the gel begin migrating toward the positively charged anode. Positively charged particles do the opposite. It may be helpful to remember that the anode attracts anions in these cells. The smallest molecules and those with the largest charge migrate the farthest through the gel while the current is applied. In many electrophoresis experiments, the molecules of interest are either intrinsically negatively charged (eg, DNA, RNA) or they are given a negative charge by altering experimental conditions. For this reason, samples are usually loaded at the end of the gel nearer to the cathode (see Figure 14.3). Certain variations on electrophoresis in which some or all of the molecules are positively charged require a different setup. However, if such a setup is not specified, it can generally be assumed that the sample is loaded into the gel near the cathode. **Figure 14.3** Electrophoresis of negatively charged particles: Samples are loaded near the cathode and migrate toward the anode, with small molecules migrating faster. * Cathode * Sample of different molecules * Gel * Anode * Negatively charged molecules migrate toward the anode. * Small molecules migrate faster * Large molecules * Small molecules Once the applied electric current is turned off, the molecules in the gel stop migrating. In general, at this point the molecules are invisible and must be visualized for analysis. Proteins are commonly visualized by applying a staining molecule called Coomassie, which binds to proteins and colors them blue. The gel is soaked in Coomassie stain and then rinsed, and the proteins appear in bands (Figure 14.4). The width or color density of a band corresponds to the amount of protein in that band. Note that electrophoresis and staining are only effective for detecting sufficiently large proteins (typically around 70 amino acids or larger). **Figure 14.4** Coomassie stain makes proteins in a gel visible. * Unstained proteins invisible in the gel * Stain added * Stained protein visible in the gel * Less protein, thinner, lighter band * More protein, thicker, darker band * Coomassie stain In DNA and RNA gels, a fluorescent dye such as ethidium bromide is often included. These dyes bind to nucleic acids, which significantly increases the dye's fluorescence intensity. DNA gels of this nature are generally visualized by exposing them to ultraviolet light, which causes the dye to fluoresce bright orange, as shown in Figure 14.5. **Figure 14.5** Visualization of DNA in a gel. * DNA fluoresces under UV light when treated with ethidium bromide Whether a gel is used to analyze DNA, RNA, or proteins, it often contains multiple lanes, each of which represents a different sample. At least one lane contains a "ladder," which is a mixture of molecules of known size and charge (see Figure 14.6). The positions of the bands in the ladder can be used to estimate the sizes of the molecules in the other lanes. **Figure 14.6** A typical gel with a ladder lane and several sample lanes. * Ladder (molecules of known size) * Samples of interest By analyzing the intensity of bands at a given position (ie, distance of migration), the relative amounts of a specific protein or DNA strand from each sample may be compared. This is commonly used to determine how protein expression varies in different conditions. An increase in band density, for example, corresponds to an increase in gene expression. Similarly, decreases in intensity may correspond to decreased expression or increased degradation. ### 14.1.02 Native Electrophoresis Native electrophoresis refers to electrophoretic techniques that preserve the functional structure of the molecule of interest. DNA and RNA electrophoresis methods are usually native. Because all DNA and RNA molecules are negatively charged due to their phosphate groups, all migrate toward the anode without any modifications. In addition, because all nucleic acids contain one phosphate group for every nitrogenous base, the charge:mass ratio of every nucleic acid is approximately the same. Therefore, although a longer DNA strand has more negative charge and experiences more force in an electric field, it also has more mass that must migrate. Consequently, the primary factor limiting how quickly nucleic acids migrate through a gel is the size of the pores. Accordingly, native gels separate linear DNA predominantly by size, with larger molecules migrating more slowly (and a shorter distance) than smaller molecules, as shown in Figure 14.7. **Figure 14.7** DNA strands of different sizes migrate different distances through a gel. * Cathode * Least migration, largest, least compact DNA molecule * Farthest migration, smallest, most compact DNA molecule * Anode Importantly, circularized DNA molecules and folded RNA molecules may behave differently than their mass alone would suggest. Folded RNA and circularized DNA (especially supercoiled DNA) are more compact, allowing them to navigate the pores of the gel more easily and migrate faster. Binding of nucleic acids to other molecules (eg, proteins) generally hinders migration through the gel by making the complex more massive. When comparing individual nitrogenous bases, guanine is larger than adenine, which is larger than thymine or uracil. Cytosine is the smallest of the nitrogenous bases. Consequently, guanine alone migrates more slowly through a gel than cytosine alone. However, a base pair consisting of A and T is approximately the same molecular weight as a pair consisting of C and G. Therefore, it is convenient to measure the size of double-stranded DNA in base pairs (bp) or kilobase pairs (kbp), as shown in Figure 14.8. **Figure 14.8** A standard DNA ladder with sizes of each band marked in both base pairs and kilobase pairs. * kbp | bp ---|--- 10 | 10,000 8 | 8,000 6 | 6,000 5 | 5,000 4 | 4,000 3 | 3,000 2.5 | 2,500 2 | 2,000 1.5 | 1,500 1 | 1,000 0.75 | 750 0.5 | 500 0.25 | 250 Native electrophoresis may also be used for proteins. This often involves polyacrylamide gels; therefore, this type of electrophoresis is commonly called native PAGE. Native PAGE for proteins is often more complicated than electrophoresis of DNA because proteins may have varied charges depending on their isoelectric points (see Chapter 2). Because native PAGE is meant to preserve protein structure, it usually occurs at physiological pH (ie, pH 7-7.4). Proteins with a pl higher than this pH range are positively charged; those with a lower pl are negatively charged. Positively charged proteins migrate toward the cathode, whereas negatively charged proteins migrate toward the anode. If two proteins are the same size and have the same sign but different magnitude charges, the one with the greater charge migrates faster. For example, a 50 kDa protein with a -2 charge tends to move toward the anode more quickly than a 50 kDa protein with a -1 charge. Similarly, two proteins of equal charge migrate differently if they are different sizes. Even two proteins of identical molecular weight and charge may migrate differently if one is folded to be more compact than the other, because more compact molecules navigate the pores of the gel more easily. Different factors that affect migration are shown in Figure 14.9. **Figure 14.9** Charge, mass, and shape (ie, compactness) all affect how proteins migrate in native PAGE. * Proteins of equal mass, folded into equally compact shapes * Charge: -3 -1 * Charges differ * Proteins with equal charge folded into equally compact shapes * Larger protein -1 * Smaller protein -1 * Masses differ * Proteins of equal mass and charge * More compact protein -1 * Less compact protein -1 * Compactness differs These factors together make it difficult to predict how far a protein will migrate in a native gel. However, once the protein of interest is identified, it remains intact, including any quaternary structure. Because protein structure is preserved, the protein can continue to function, even within the gel. Consequently, native gels may be used to assess binding interactions, because interactions with a ligand alter the mobility of the protein. This is called a mobility shift (Figure 14.10). **Figure 14.10** Native PAGE preserves secondary, tertiary, and quaternary protein structure. Binding to a ligand can result in a mobility shift. * Native PAGE * Mobility shift * Functional protein * Ligand * Protein-ligand complex ### 14.1.03 Denaturing Electrophoresis Denaturing electrophoresis is most commonly used to separate proteins. Denaturing electrophoresis is typically carried out by adding a denaturing agent, usually sodium dodecyl sulfate (SDS), to the protein mixture. This is generally used with polyacrylamide gel electrophoresis, and the technique is often referred to as SDS-PAGE. Other denaturing agents are occasionally used, but SDS illustrates the overall principles of denaturing electrophoresis. SDS is a detergent. It consists of a sodium ion and a dodecyl sulfate molecule. Dodecyl sulfate consists of a negatively charged head group (the sulfate portion), which interacts with water, and a hydrophobic 12-carbon chain. The hydrophobic tail interacts favorably with hydrophobic side chains in a protein, which disrupts the hydrophobic effect (Lesson 2.3) and causes the protein to unfold. The head group confers a uniform negative charge on the proteins in the sample. Figure 14.11 shows interactions between SDS and proteins. **Figure 14.11** Sodium dodecyl sulfate (SDS) denatures proteins and confers a uniform negative charge, allowing separation of proteins by size only. * Na+ * Sodium dodecyl sulfate (SDS) * Native protein with hydrophobic interior * Denaturing by SDS * Native shape and net charge are lost * SDS-PAGE * SDS-PAGE separates proteins by size only * Large protein * Small protein As a denaturing agent, SDS disrupts secondary, tertiary, and quaternary structure and renders proteins nonfunctional. Note, however, that SDS by itself does not break disulfide bonds; additional reagents, described in Concept 14.1.04, can be used together with SDS to break disulfide bonds. Because SDS is a denaturing agent, proteins separated by SDS-PAGE cannot be used to assess protein activity unless the proteins are first allowed to renature. ### 14.1.04 Reducing Gels The previous concept described how denaturing agents such as SDS disrupt secondary, tertiary, and quaternary structure but noted that SDS does not break disulfide bonds. Disulfide bonds form in proteins when the thiol (-SH) groups of two cysteine residues undergo oxidation to form an -S-S- bond. This converts the cysteine residues to cystine. Disulfide bonds can be broken by adding a reducing agent to the SDS-PAGE experiment. The most common reducing agents for this purpose are dithiothreitol (DTT) and ẞ-mercaptoethanol (BME). Each of these agents reduces the disulfide bonds in proteins by becoming oxidized and forming disulfide bonds themselves (Figure 14.15). The resulting reaction converts cystine (oxidized cysteine) back to two separate cysteine residues. **Figure 14.15** Reduction of disulfide bonds converts cystine to two cysteine side chains while oxidizing either DTT or BME. * Disulfide bond * Thiol group * COO- * DTT * COO- * Oxidized DTT * *H3N-CH * *H3N-CH * S-S * HS * CH2 * CH2 * SH * OH * HO * OH * S * SH * + * or * + * or * SH * S * Oxidized BME * 2 BME * CH2 * CH2 * SH * S-S * CH-NH3+ * HO * CH-NH3+ * HO * OH * SH * HO * COO- * COO- * Cystine * 2 Cysteine SDS-PAGE experiments that use DTT or BME are typically designated specifically as reducing SDS-PAGE. When DTT and BME are absent, it is often designated as nonreducing SDS-PAGE. Reducing and nonreducing gels yield different results when the proteins of interest contain disulfide bonds. A protein with subunits held together by one or more disulfide bonds appears as a single band in nonreducing SDS-PAGE because the disulfide bond keeps the subunits together, although they are denatured. In contrast, the subunits separate when a reducing agent is added. If the protein is a homodimer (ie, identical subunits), the gel will still show a single band, but it will appear further down the gel (closer to the anode) because each individual subunit is smaller than the combined subunits. If the protein is a heterodimer, two distinct bands will appear, with the band closer to the cathode corresponding to the larger subunit. Figure 14.16 shows the separation of disulfide-linked homo- and heterodimers by addition of a reducing agent. **Figure 14.16** A reducing agent separates disulfide-linked homodimers to produce a single, faster-migrating band and separates disulfide-linked heterodimers to produce two distinct bands. * Disulfide bond * Nonreducing SDS-PAGE * Reducing SDS-PAGE * Individual, identical subunits (disulfide bond broken) * Large subunit * Disulfide bond * Nonreducing SDS-PAGE * Reducing SDS-PAGE * Small subunit * Individual, distinct subunits (disulfide bond broken) ### 14.1.05 Isoelectric Focusing In addition to separation based on size or charge, proteins can be separated based on their isoelectric point (pl) by performing an isoelectric focusing experiment. To accomplish this, a polyacrylamide gel is prepared with a stable pH gradient. In other words, one end of the gel contains a low pH, the other end contains a high pH, and the pH gradually increases from the low end to the high end. Typically, these gels do not contain SDS or other denaturing agents and allow the proteins to maintain their native structures. The end of the gel with a high pH (ie, low H+ concentration) is placed near the cathode (negative charge), and the end with a low pH (ie, high H+ concentration) is placed near the anode (positive charge). Proteins may then be loaded onto the gel at any position (near the anode, near the cathode, or in the middle). In other words, isoelectric focusing is a case in which the sample does not have to be loaded near the cathode. Proteins at a pH that is less than their pl will pick up protons from the environment and gain a positive charge, whereas those at positions where the pH is greater than their pl will lose protons to their environment and become negatively charged. This is shown in Figure 14.17. **Figure 14.17** Proteins placed near the cathode in an isoelectric focusing gel tend to be negatively charged, while those near the anode tend to be positively charged. * Cathode * pH > protein pl: Protein is negatively charged * -NH2 * COO- * Θ * pH < protein pl: Protein is positively charged * NH3 * COO- * Θ * pH 14 13 12 11 10 9 8 7 6 5 4 3 2 1 * Anode When a current is applied and an electric field generated, positively charged proteins migrate closer to the negatively charged cathode, where the pH of the gel is higher. As the gel pH increases, these proteins lose protons and therefore become less positively charged. Eventually they reach a position in the gel where pH is equal to pl and the net charge of the protein drops to 0. Particles with 0 net charge do not migrate in an electric field because they experience no net force. Therefore, at this point migration stops. Similarly, proteins with a negative charge migrate toward the positively charged anode, where the pH is lower. These proteins pick up protons and become less negatively charged. When the pH of the gel matches the isoelectric point, the net charge becomes 0 and migration stops. Therefore, regardless of where a given protein is loaded onto an IEF gel, it migrates (or focuses) toward the point where the gel's pH equals its isoelectric point. This technique facilitates empirical determination of a protein's pl. As explained in Lesson 2.1, calculation of a protein's isoelectric point becomes difficult in complex, folded proteins because the pKa values of the side chains may change due to interactions with each other. Therefore, a calculation using the pKa values of free amino acid side chains may be inaccurate. Isoelectric focusing, however, directly measures pl: whichever pH stops the protein from migrating is the pl of that protein. Figure 14.18 shows a protein's migration toward its isoelectric point. **Figure 14.18** Migration through the isoelectric focusing gel occurs until the protein's net charge becomes 0. * Cathode * Negative protein migrates toward anode, gains protons * H3N * COO- * -NH2 * COO- * Θ * Positive protein migrates towards cathode, loses protons * H3N * COO- * -NH3 * COO- * Θ * pH 14 13 12 11 10 9 8 7 6 5 4 3 2 1 * Net charge = 0 (charges cancel) * Migration stops, pl ≈ 8 * Cathode * pH 14 13 12 11 10 9 8 7 6 5 4 3 2 1 * Protein band visualized * Anode * Anode ### 14.1.06 2D Gels SDS-PAGE separates proteins by size, and isoelectric focusing separates them by isoelectric point. However, in a mixture of proteins, it is possible that two different proteins will be of similar size and migrate to the same position in SDS-PAGE, or they may have the same (or nearly the same) isoelectric point and migrate to the same position in isoelectric focusing. To further separate these proteins, both techniques may be applied, one after the other. Because this technique involves two parameters, or dimensions, it is called **two-dimensional (2D) electrophoresis**. Typically, a sample of proteins is first subjected to isoelectric focusing. The resulting isoelectric focusing gel is treated with SDS to make all proteins within it negatively charged, and the treated gel is then aligned with the sample-loading edge of an SDS-PAGE gel. When a current is applied, the proteins migrate from the IEF gel into the SDS-PAGE gel, where they separate based on size. Unlike traditional gels, the result does not have evenly distributed lanes. Instead, 2D gels generally have multiple spots at various locations throughout the gel, each of which corresponds to a specific protein (Figure 14.19). The final location of a spot represents the isoelectric point along one axis and the molecular weight in kilodaltons on the other. **Figure 14.19** Example of a 2D gel using isoelectric focusing in one dimension and SDS-PAGE in the other. Other variations of 2D gels are also possible. For instance, isoelectric focusing may be followed by native PAGE instead of SDS-PAGE, or native PAGE may occur first, followed by SDS-PAGE. Therefore, the positions of each protein would show their isoelectric point and native size:charge ratio, or their native size:charge ratio and denatured size, respectively.