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How does SDS-PAGE separate based on mass?

How does SDS-PAGE separate based on mass?


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In SDS-PAGE, electric field and mass-to-charge ratio are approximated to be constant for all proteins. Also, if $F=qE =ma$, then $frac{m}{q}a=E$.

Thus, all proteins must migrate with a constant acceleration.

So what is the driving force in SDS-PAGE that resolves proteins of different mass? Is it that bigger proteins have a larger size and thus experience larger friction force from the gel sieves? But even if that was true the larger proteins feeling larger resistance forces should be counteracted by their larger migrating force from increased total charge.

Would appreciate if someone could clear up this confusion. Thank you.


You are correct that molecule mobility depends on the mass-to-charge ratio, and this means that different sized molecules with the same $frac{m}{q}$ will have the same acceleration. However, the velocity of a molecule moving through a gel matrix depends on the point at which the force exerted by the electric field is in equilibrium with the frictional forces acting on the molecule. Modeling different sized molecules as spheres of different sizes, we can apply Stokes' Law

$F = 6pi mu Rv$

where $mu$ is viscosity of the matrix (constant), $R$ is the radius of our molecule, and $v$ is the flow velocity.

Rewriting your equation, we see that

$F = qE = ma = 6pi mu Rv$

and

$v = frac {ma}{6pi mu R}$

For constant $a$ and $mu$, we see that velocity varies with the ratio of mass to radius

$v sim frac{m}{R}$

For our model spheres, assuming uniform density, radius scales cubically with mass, meaning that $frac{m}{R}$ decreases as molecule size increases.

($d = frac{m}{V}$ and $V = frac{4}{3}pi R^3$ where $d$ is density and $V$ is volume)

So, larger molecules will reach force equilibrium with the opposing frictional resistance at a lower velocity compared to smaller molecules, thus explaining how different sized molecules with the same $frac{m}{q}$ will separate on a gel. One caveat to this explanation is that polypeptides and nucleic acids are poorly modeled by spheres, and that the mass of such molecules scales linearly with length.


How Does SDS-PAGE Work?

Electrophoresis is a major technique for separating proteins and other substances such as nucleic acids, purines, pyrimidines, some organic compounds and even inorganic ions. Most of the current electrophoresis is to separate the sample into the mobile phase in an immobilized medium. Polyacrylamide gel is one of the main media. It is a porous gel whose pore size is close to the size of protein molecules, which improves the resolution of proteins. Moreover, the polyacrylamide gel has good chemical stability, strong repeatability, stability to changes in pH and temperature, and easy color observation. SDS polyacrylamide gel electrophoresis (SDS-PAGE) has the advantages of simple operation and good reproducibility in the determination of protein molecular weight, detection of specific proteins, and identification of strain species.

Polyacrylamide gel is composed of acrylamide and cross-linking agent N, N'-methylenebisacrylamide under the action of catalysts ammonium persulfate (AP) and N, N, N', N'-Tetramethylethylenediamine (TEMED). It is a gel with a three-dimensional network structure. PAGE can separate proteins into several bands according to the different mobility caused by the different charge and molecular weight of protein molecules. SDS is an anionic surfactant, which can break the hydrogen and hydrophobic bonds of proteins in the presence of reducing agents (β-mercaptoethanol or dithiothreitol, DTT), and combine with protein molecules in a certain ratio to form short rod-shaped composites of the same density. Positively correlated with the molecular weight of the protein, the length of the complex formed by proteins of different molecular weights is different. SDS makes the amount of negatively charged protein far exceed its original charge, masking the natural charge difference between various protein molecules. Therefore, the mobility of various protein-SDS complexes during electrophoresis is no longer affected by the original charge and molecular shape, but only depends on the relative molecular mass.

The polyacrylamide gel is usually composed of a stacking gel in the upper layer and a separating gel in the lower layer. The difference between the upper and lower gels is the concentration of acrylamide and the pH of Tris-HCl. During electrophoresis, an electric field is applied to the gel, and negatively charged proteins migrate across the gel from the negative electrode to the positive electrode. The most common electrophoresis buffer consists of Tris and glycine. The pH in the stacking gel is 6.8, and only a few glycine molecules dissociate. Therefore, the SDS-treated protein molecules move between the upper glycine molecule and the lower Cl- ion. This process compresses the protein sample in the gel into bands that are much smaller than the volume initially loaded. As the electrophoresis progresses, the protein moves to the separating gel (pH 8.8), where of the glycine molecules dissociate. The speed of the movement increases and exceeds the protein. In the separating gel, the speed of movement of each protein depends on its molecular weight. Proteins with small molecular weights can pass through the pores in the gel easily, while those with large molecular weights have more difficulty passing through. After a period of time, proteins reach different distances according to the sizes, achieving the purpose of protein separation.

Figure 1. Schematic diagram of polyacrylamide gel electrophoresis (Gülay, et al, 2018).

How to Determine Molecular Weight of Protein by SDS-PAGE?

SDS-PAGE is the main method to determine the molecular weight of unknown proteins. A protein with known molecular weight and an unknown sample are electrophoresed at the same time. After staining, according to the relative mobility of the standard protein and the logarithm of the molecular weight, a line can be obtained and determine the molecular weight of the unknown sample using its relative mobility. In the laboratory, a standard molecular weight protein covalently coupled to a dye is used as a reference protein to roughly indicate the size of the unknown protein. This pre-stained protein marker can be directly observed during electrophoresis or when transferring membranes.

How to Read SDS-PAGE Results?

After electrophoresis, protein separation cannot be directly observed by the naked eye, and subsequent staining techniques are needed. Coomassie brilliant blue staining and silver staining are common methods for routine detection and quantification of proteins separated by electrophoresis. After simple processing such as fixation-staining-decolorization, the distribution of protein can be clearly observed. With the improvement of high-sensitivity protein analysis methods and protein identification technologies, new staining methods such as fluorescent labeling and isotope labeling technology have greatly improved sensitivity, and are also compatible with automated proteome platform gel cutting technology. More high sensitivity and automated dyeing technologies are been developed.

How to Store SDS-PAGE Gel?

Freshly SDS-PAGE gels are usually prepared before each experiment. However, gels can also be stored in clean water at 4°C for about a week. If the gel cannot be photographed in time after dyeing, it needs to be placed in water to prevent drying and shrinking of the gel. It is advised to photograph the staining results as soon as possible. Band will disperse if the gel is soaked in water for a long time.

References
1. Smith B J. SDS Polyacrylamide Gel Electrophoresis of Proteins. Methods in Molecular Biology, 1984, 1(4):41-55.
2. Duffy M F, Noormohammadi A H, Baseggio N, et al. Polyacrylamide gel-electrophoresis separation of whole-cell proteins. Methods in Molecular Biology, 1998, 104:267.

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Molecular mass versus molecular weight

Molecular mass (symbol m) is expressed in Daltons (Da). One Dalton is defined as 1/12 the mass of carbon 12. Most macromolecules are large enough to use the kiloDalton (kDa) to describe molecular mass. Molecular weight is not the same as molecular mass. It is also known as relative molecular mass (symbol Mr, where r is a subscript). Molecular weight is defined as the ratio of the mass of a macromolecule to 1/12 the mass of a carbon 12 atom. It is a dimensionless quantity.

When the literature gives a mass in Da or kDa it refers to molecular mass. It is incorrect to express molecular weight (relative molecular mass) in Daltons. Nevertheless you will find the term molecular weight used with Daltons or kiloDaltons in some literature, often using the abbreviation MW for molecular weight.


How does SDS-PAGE separate based on mass? - Biology

BISC411
EXPERIMENTAL MOLECULAR BIOLOGY OF THE CELL

Principles of Polyacrylamide Gel Electrophoresis (PAGE)

Powerful electrophoretic techniques have been developed to separate macromolecules on the basis of molecular weight. The mobility of a molecule in an electric field is inversely proportional to molecular friction which is the result of its molecular size and shape, and directly proportional to the voltage and the charge of the molecule. Proteins could be resolved electrophoretically in a semi-solid matrix strictly on the basis of molecular weight if, at a set voltage, a way could be found to charge these molecules to the same degree and to the same sign. Under these conditions, the mobility of the molecules would be simply inversely proportional to their size.

It is precisely this idea which is exploited in PAGE to separate polypeptides according to their molecular weights. In PAGE, proteins charged negatively by the binding of the anionic detergent SDS (sodium dodecyl sulfate) separate within a matrix of polyacrylamide gel in an electric field according to their molecular weights.
Polyacrylamide is formed by the polymerization of the monomer molecule-acrylamide crosslinked by N,N'-methylene-bis-acrylamide (abbreviated BIS). Free radicals generated by ammonium persulfate (APS) and a catalyst acting as an oxygen scavenger (-N,N,N',N'-tetramethylethylene diamine [TEMED]) are required to start the polymerization since acrylamide and BIS are nonreactive by themselves or when mixed together.

The distinct advantage of acrylamide gel systems is that the initial concentrations of acrylamide and BIS control the hardness and degree of crosslinking of the gel. The hardness of a gel in turn controls the friction that macromolecules experience as they move through the gel in an electric field, and therefore affects the resolution of the components to be separated. Hard gels (12-20% acrylamide) retard the migration of large molecules more than they do small ones. In certain cases, high concentration acrylamide gels are so tight that they exclude large molecules from entering the gel but allow the migration and resolution of low molecular weight components of a complex mixture. Alternatively, in a loose gel (4-8% acrylamide), high molecular weight molecules migrate much farther down the gel and, in some instances, can move right out of the matrix.

SDS Polyacrylamide Gel Electrophoresis (SDS-PAGE)

Sodium dodecyl sulfate (SDS or sodium lauryl sulfate) is an anionic detergent which denatures proteins molecules without breaking peptide bonds. It binds strongly to all proteins and creates a very high and constant charge:mass ratio for all denatured proteins. After treatment with SDS, irrespective of their native charges, all proteins acquire a high negative charge.

Denaturation of proteins is performed by heating them in a buffer containing a soluble thiol reducing agent (e.g. 2-mercaptoethanol dithiothreitol) and SDS. Mercaptoethanol reduces all disulfide bonds of cysteine residues to free sulfhydryl groups, and heating in SDS disrupts all intra- and intermolecular protein interactions. This treatment yields individual polypeptide chains which carry an excess negative charge induced by the binding of the detergent, and an identical charge:mass ratio. Thereafter, the denatured proteins can be resolved electrophoretically strictly on the basis of size in a buffered polyacrylamide gel which contains SDS and thiol reducing agents.

SDS-PAGE gel systems are exceedingly useful in analyzing and resolving complex protein mixtures. Many applications and modifications of this technique are relevant to modern experimental biologists. Some are mentioned below. They are employed to monitor enzyme purification, to determine the subunit composition of oligomeric proteins, to characterize the protein components of subcellular organelles and membranes, and to assign specific proteins to specific genes by comparing protein extracts of wild-type organisms and suppressible mutants. In addition, the mobility of polypeptides in SDS-PAGE gel systems is proportional to the inverse of the log of their molecular weights. This property makes it possible to measure the molecular weight of an unknown protein with an accuracy of +/- 5%, quickly, cheaply and reproducibly.

Discontinuous SDS Polyacrylamide Gel Electrophoresis

Disc gels are constructed with two different acrylamide gels, one on top of the other. The upper or stacking gel contains 4-5% acrylamide (a very loose gel) weakly buffered at pH 9.0. The lower resolving gel (often called the running gel), contains a higher acrylamide concentration, or a gradient of acrylamide, strongly buffered at pH 9.0. Both gels can be cast as tubes in glass or plastic cylinders (tube gels), or as thin slabs within glass plates, an arrangement which improves resolution considerably, and which makes it possible to analyze and compare many protein samples at once, and on the same gel (slab gels). Today, you will be constructing and running slab gels.

The ionic strength discontinuity between the loose stacking gel and the hard running gel leads to a voltage discontinuity as current is applied. The goal of these gels is to maximize resolution of protein molecules by reducing and concentrating the sample to an ultrathin zone (1-100 nm) at the stacking gel:running gel boundary. The protein sample is applied in a well within the stacking gel as a rather long liquid column (0.2-0.5 cm) depending on the amount and the thickness of the gel or tube. The protein sample contains glycerol or sucrose so that it can be overlaid with a running buffer. This buffer is called the running buffer, and the arrangement is such that the top and bottom of the gel are in running buffer to make a circuit.

As current is applied, the proteins start to migrate downward through the stacking gel toward the positive pole, since they are negatively charged by the bound SDS. Since the stacking gel is very loose, low and average molecular weight proteins are not impeded in their migration and move much more quickly than in the running gel. In addition, the lower ionic strength of the stacking gel (weak buffer) creates a high electrical resistance, (i.e., a high electric field V/cm) to make proteins move faster than in the running gel (high ionic strength, lower resistance, hence lower electric field, V/cm). Remember that applied voltage results in current flow in the gel through the migration of ions. Hence low ionic strength means high resistance because fewer ions are present to dissipate the voltage and the electric field (V/cm) is increased causing the highly polyanionic proteins to migrate rapidly.

The rapid migration of proteins through the stacking gel causes them to accumulate and stack as a very thin zone at the stacking gel/running gel boundary, and most importantly, since the 4-5% stacking gel affects the mobility of the large components only slightly, the stack is arranged in order of mobility of the proteins in the mixture. This stacking effect results in superior resolution within the running gel, where polypeptides enter and migrate much more slowly, according to their size and shape.

In all gel systems, a tracking dye (usually Bromophenol blue) is introduced with the protein sample to determine the time at which the operation should be stopped. Bromophenol blue is a small molecule which travels essentially unimpeded just behind the ion front moving down toward the bottom of the gel. Few protein molecules travel ahead of this tracking dye. When the dye front reaches the bottom of the running gel, the current is turned off to make sure that proteins do not electrophorese out of the gel into the buffer tank.

Visualizing the Proteins

Gels are removed from tubes or from the glass plates and stained with a dye, Coomassie Brilliant Blue. Coomassie blue binds strongly to all proteins. Unbound dye is removed by extensive washing of the gel. Blue protein bands can thereafter be located and quantified since the amount of bound dye is proportional to protein content. Stained gels can be dried and preserved, photographed or scanned with a recording densitometer to measure the intensity of the color in each protein band. Alternatively, if the proteins are radioactive, the protein bands can be detected by autoradiography, a technique that is widely used in modern cell and molecular biology. When gels are prepared as thin slabs to maximize resolution as you will do today, the slabs of acrylamide are removed from the support glass plates and dried on filter paper. A piece of X-ray film is placed and clamped tight over the dried slab in a light-proof box. The X-ray film is exposed by the radioactivity in the protein bands and, after developing, dark spots or bands can be seen on the film. These dark bands can in turn be quantified since their intensity is proportional to the amount of radioactivity and hence to protein content.


How does SDS-PAGE separate based on mass? - Biology

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SDS-PAGE – Gel Based Protein Separation

SDS-PAGE is a method of separating proteins based on their molecular mass. SDS (sodium dodecyl sulfate) is a detergent that binds proteins and covers them with a negative charge. In general, one SDS molecule binds to two amino acids. After exposure to SDS different proteins will have very similar charge to mass ratios because SDS coats the protein in a negative charge overwhelming whatever intrinsic charge the protein originally had. The acronym PAGE stands for polyacrylamide gel electrophoresis. The protein mixture is denatured by adding SDS and beta-mercaptoethanol (used to break disulfide bonds) and then heated. The denatured protein mixture is added to a polyacrylamide gel where an electric field is applied and the proteins, which are now coated with a negative charge, move towards the positive electrode (the anode).

Proteins move through the gel based on their size. The polyacrylamide gel can be thought of as a series of nets stacked on top of the each other. The higher the percentage of polyacrylamide the more nets there are and the size of the holes between the polyacrylamide chains becomes smaller making it more difficult for proteins to move through. If you want to separate proteins that are relatively small, a higher percentage of polyacrylamide should be used. If one needs to separate larger proteins a smaller percentage of polyacrylamide would be used. Because the sample buffer contains blue dye which binds to proteins, the proteins that are separated by SDS-PAGE form blue bands which are easily visualized.

SDS-PAGE is often used as a quality control measure for biopharmaceuticals. A pure protein product with a single chain should show only one band using SDS-PAGE. Any additional bands would be contaminants or degradation products. SDS-PAGE is also the first step in creating a western blot which can be used to identify a particular protein within a mixture.


Variants

SDS-PAGE is the most widely used method for gel electrophoretic separation of proteins. Two-dimensional gel electrophoresis sequentially combines isoelectric focusing or BAC-PAGE with a SDS-PAGE. Native PAGE is used if native protein folding is to be maintained. For separation of membrane proteins, BAC-PAGE or CTAB-PAGE may be used as an alternative to SDS-PAGE. For electrophoretic separation of larger protein complexes, agarose gel electrophoresis can be used, e.g. the SDD-AGE. Some enzymes can be detected via their enzyme activity by zymography.


When proteins are separated by electrophoresis through a gel matrix, smaller proteins migrate faster due to less resistance from the gel matrix. Other influences on the rate of migration through the gel matrix include the structure and charge of the proteins.

In SDS-PAGE, the use of sodium dodecyl sulfate (SDS, also known as sodium lauryl sulfate) and polyacrylamide gel largely eliminates the influence of the structure and charge, and proteins are separated solely based on polypeptide chain length.

SDS is a detergent with a strong protein-denaturing effect and binds to the protein backbone at a constant molar ratio. In the presence of SDS and a reducing agent that cleaves disulfide bonds critical for proper folding, proteins unfold into linear chains with negative charge proportional to the polypeptide chain length.

Polymerized acrylamide (polyacrylamide) forms a mesh-like matrix suitable for the separation of proteins of typical size. The strength of the gel allows easy handling. Polyacrylamide gel electrophoresis of SDS-treated proteins allows researchers to separate proteins based on their length in an easy, inexpensive, and relatively accurate manner.


How gel concentration affects relative mobility

So what happens if you want to characterize all of the proteins in a sample? . run more than one gel, of course! A gel with low density will resolve the larger polypeptides while cutting off the lighter ones, and one of higher density will reveal the smaller polypeptides, while compressing and possibly distorting the larger ones.

Here are some examples of the effect of acrylamide concentration on relative mobility. Molecular weight standards are identified by number. A typical erythrocyte membrane protein sample is also presented, with band 3 protein labeled as a reference.

Standard 1 = myosin (205,000) std. 2 = beta-galactosidase (116,000) std. 3 = phosphorylase B (92,000) std. 4 = bovine serum albumin (66,000) std. 5 = egg albumin (45,000) std. 6 = carbonic anhydrase (29,000).

On the gels from 6 to 10% there is a distinct dark doublet at the top of the membrane lane. Notice in the 12% gel the doublet is jammed together and appears as one band. One could estimate MW of band 3 from the first five gels although the best estimate comes from the 6%, which produced the greatest separation between the standards on either side of band 3. The 12% gel did not resolve bands well at all above the fourth standard (serum albumin, 66,000). Analysis should be confined to the part of the gel below 66,000 or even lower if the same sample was run on a lower density gel.

Frequently, students analyze the upper part of a high density acrylamide gel that overlaps part of a lower density gel. That practice suggests that the student missed the point of the anaysis and/or did not understand the limitations of the method. Interpret only that part of the denser gel that doesn't overlap the other one. To put it another way, use high density gels to study proteins (or parts of proteins) of relatively low molecular weight, and lower density gels to resolve proteins of higher molecular weight.

Don't forget that different polypeptides can have similar or even identical molecular masses. One band on a gel can therefore consist of one or more polypeptides. This is most likely to happen toward the top of a gel, and especially in higher density gels.


Lysate preparation

Cell lysis is the first step in protein extraction, fractionation and purification. Numerous techniques have been developed to obtain the best possible yield and purity for different organisms, sample types (cells or tissue), subcellular structures or specific proteins. Both physical and reagent-based methods may be required to extract cellular proteins because of the diversity of cell types and cell membrane (or cell wall) composition .

Lysis

Physical lysis is a common method of cell disruption and extraction of cellular contents. However, it requires specialized equipment and protocols that are difficult to repeat because of variability in the apparatus (e.g., different dounce pestles or sonication settings). Also, traditional physical disruption methods are typically not conducive to small sample volumes and high-throughput sample handling. Finally, physical lysis methods alone are unable to solubilize membrane-associated proteins. In contrast, reagent-based lysis methods using detergents not only lyse cells but also solubilize proteins. By using different buffers, detergents, salts and reducing agents, cell lysis can be optimized to provide the best possible results for a particular cell type or protein fraction.

Protein stability

Cell lysis disrupts cellular compartments, which can activate endogenous proteases and phosphatases. To protect extracted proteins from degradation or artifactual modification by the activities of these enzymes, it is necessary to add protease and/or phosphatase inhibitors to the lysis reagents.

When the goal of cell lysis is to purify or test the function of a particular protein(s), special attention must be given to the effects that the lysis reagents have on the stability and function of the target proteins. Certain detergents will inactivate the function of particular enzymes or disrupt protein complexes. Downstream analysis of extracted/purified proteins may also require detergent removal in order to study proteins of interest or maintain long-term stability of the extracted protein.

Depletion and enrichment

Sample complexity negatively affects the ability to detect, identify and quantify low-abundance proteins by MS, because peptides from high-abundance proteins can mask detection of those from low-abundance proteins. Therefore, the more that a sample can be simplified and the greater that the dynamic range of protein concentrations can be reduced, the greater will be the ability to detect proteins at very low concentrations.

Depletion and enrichment strategies have been developed to remove high abundant proteins or isolate target proteins in the sample, respectively. Depletion is more often used to reduce the complexity of biological samples such as blood or serum, which contain high concentrations of albumin and immunoglobulins. Depletion strategies utilize immunoaffinity techniques such as immunoprecipitation and co-immunoprecipitation (IP and co-IP, respectively), and commercial kits are available to remove these and other high abundant proteins from samples. A significant drawback to this approach, though, is that abundant proteins often bind to other proteins, which could result in the depletion of complexes with low-abundance proteins.

Protein enrichment encompasses numerous techniques to isolate subclasses of cellular proteins based on unique biochemical activity, post-translational modifications (PTMs) or spatial localization within a cell. Post-translational modifications such as phosphorylation and glycosylation can be enriched using affinity ligands such as ion-metal affinity chromatography (IMAC) or immobilized lectins, respectively. In addition, PTM-specific antibodies have been used. Other techniques entail metabolic or enzymatic incorporation of modified amino acids or PTMs to introduce unique protein chemistries that can be used for enrichment. Finally, proteins can also be enriched using various enzyme class-specific compounds or cell-impermeable labeling reagents that selectively label cell surface proteins.

Separation of distinct subcellular fractions is another method of enrichment and can be achieved through the careful optimization of physical disruption techniques, detergent-buffer solutions and density gradient methods. For example, with the phase-separating detergents, hydrophobic membrane proteins can be solubilized and extracted from hydrophilic proteins. Density gradient centrifugation is another technique and can be used to isolate intact nuclei, mitochondria and other organelles before protein solubilization.

Dialysis and desalting

Whether they are simple or complex, samples often need to be processed in several ways to ensure they are compatible and optimized for digestion and analysis by mass spectrometry. For example, because MS measures charged ions, salts—especially sodium and phosphate salts—should be removed prior to MS to minimize their detection.

Dialysis and desalting products allow buffer exchange, desalting, or small molecule removal to prevent interference with downstream processes. Protein assays help monitor protein concentration for consistent control of experimental loading or yield.