Tris and SDS are two chemicals that, when used together in polyacrylamide gel electrophoresis, allow researchers to separate proteins purely by size. Tris (tris(hydroxymethyl)aminomethane) acts as the buffer that maintains a stable pH, while SDS (sodium dodecyl sulfate) is a detergent that unfolds proteins and coats them with a uniform negative charge. The combination was developed in the early 1970s and quickly became one of the most widely used techniques in molecular biology and biochemistry, a workhorse method that appears in thousands of published studies every year.1PubMed Central. Using T4 genetics and Laemmli’s development of high-resolution SDS gel electrophoresis to reveal structural protein interactions controlling protein folding and phage self-assembly Understanding how each component works, and where the system breaks down, matters for anyone running gels in a lab or interpreting the banded patterns that come out of them.
What SDS Does to Proteins
SDS is a powerful anionic detergent. When you dissolve a protein in an SDS-containing buffer, the detergent molecules bind along the polypeptide chain. Above a certain concentration, this binding ratio becomes remarkably consistent across different proteins: roughly the same amount of SDS per gram of protein, regardless of what that protein is.2Journal of Biological Chemistry. The interaction of sodium dodecyl sulfate with proteins. 1. Denatured protein-polypeptide complexes That uniformity is the key to the whole technique. Because the detergent contributes a large, consistent negative charge per unit mass, differences in a protein’s own native charge become irrelevant. A protein that would normally carry a slight positive charge at a given pH ends up just as negatively charged per gram as one that was already negative.
SDS also unfolds proteins. Native proteins are folded into elaborate three-dimensional shapes, compact globules or sprawling multi-subunit complexes. SDS strips most of that structure away, converting each polypeptide into an elongated, rod-like particle whose length scales with molecular weight.3Journal of Biological Chemistry. The interaction of sodium dodecyl sulfate with proteins. 1. Denatured protein-polypeptide complexes The practical result is that when these SDS-protein complexes are pushed through a gel by an electric field, the only variable left controlling how fast each one moves is its size. Small proteins slip through the gel pores quickly. Large ones get held back. That is why the method can estimate molecular weight.
The Role of Tris as a Buffer
Maintaining a stable pH during electrophoresis is not optional. Proteins change their charge behavior at different pH values, and the chemical reactions involved in running current through a gel produce ions that would shift pH rapidly without a buffer to absorb them. Tris is the buffer of choice in the classic system developed by Ulrich Laemmli at the MRC Laboratory of Molecular Biology in Cambridge.4PubMed Central. Using T4 genetics and Laemmli’s development of high-resolution SDS gel electrophoresis to reveal structural protein interactions controlling protein folding and phage self-assembly The Laemmli system uses Tris at two different pH values: one in the gel itself (around pH 8.8 in the resolving gel, around pH 6.8 in the stacking gel) and glycine in the running buffer. This discontinuous buffer system creates a stacking effect that compresses all the proteins into a tight band before they enter the resolving gel, which dramatically sharpens the final bands.
The pH difference between the stacking and resolving gels is what makes the system “discontinuous.” In the stacking gel, glycine exists mostly in an uncharged form and moves slowly, while chloride ions from the gel buffer move quickly. Proteins get sandwiched between these two ion fronts and concentrated into a thin layer. Once they hit the higher-pH resolving gel, glycine picks up charge, the ion front effect dissipates, and proteins separate based on size as they sieve through the polyacrylamide matrix.
How the Gel Matrix Works
Polyacrylamide gels act as molecular sieves. The gel is formed by polymerizing acrylamide monomers with a crosslinker, creating a mesh of pores. Higher acrylamide concentrations produce smaller pores, which slow down large proteins more and improve resolution in the lower-molecular-weight range. Lower acrylamide concentrations produce larger pores that let big proteins pass and resolve better in the high-molecular-weight range.5PubMed. One-dimensional SDS gel electrophoresis of proteins Choosing the right percentage is one of the first decisions a researcher makes when setting up a gel. A 10% gel is a common all-purpose choice, while a 15% gel resolves small proteins better and a 6% or 7% gel handles very large ones.
Gradient gels, where the acrylamide concentration increases from top to bottom, offer a wider separation range in a single run. As proteins migrate downward, they encounter progressively smaller pores until they effectively slow to a crawl at the point where pore size matches their effective diameter. This compresses the useful separation range into one gel rather than forcing you to run multiple gels at different percentages.
Sample Preparation and Common Pitfalls
Before loading proteins onto a gel, you mix them with a sample buffer that typically contains SDS, a reducing agent like beta-mercaptoethanol or dithiothreitol (DTT), glycerol for density, and a tracking dye. The SDS coats the proteins, the reducing agent breaks disulfide bonds that hold subunits together or stabilize internal loops, the glycerol ensures the sample sinks into the well, and the dye lets you watch the electrophoretic front travel through the gel.
Heating the sample is standard practice, usually to around 95°C for a few minutes, which helps SDS fully denature the protein. But this step is not always benign. Certain membrane proteins aggregate irreversibly when boiled in SDS-containing buffers. The vesicular monoamine transporter, for instance, forms SDS-resistant aggregates at high temperatures, apparently because boiling unfolds residual structure in a way that exposes sticky hydrophobic surfaces.6PubMed Central. SDS-resistant aggregation of membrane proteins: application to the purification of the vesicular monoamine transporter The SARS-CoV membrane protein shows a similar problem: boiling causes aggregation in the stacking gel, and the aggregated protein never makes it into the resolving gel at all. The hydrophobic regions of the protein drive the aggregation, and even strong detergents or denaturants cannot reverse it once it forms.7PubMed Central. Thermal aggregation of SARS-CoV membrane protein A lens membrane protein shows the same vulnerability, with aggregation worsened by higher ionic strength in the buffer.8Biochemical and Biophysical Research Communications. Heat induced aggregation of the sodium dodecyl sulfate-solubilized main intrinsic polypeptide isolated from bovine lens plasma membrane
The practical takeaway is that if your target protein is a hydrophobic membrane protein and it keeps disappearing from gels, try incubating the sample at a lower temperature, perhaps 37°C or room temperature, rather than boiling. This is one of the more common troubleshooting steps for membrane protein work.
Reducing Agents and Disulfide Bonds
Reducing agents break disulfide bonds so that multi-subunit proteins fall apart into their individual chains, and internal disulfide loops are opened. But both beta-mercaptoethanol and DTT migrate through the gel at the speed of the dye front rather than staying with the proteins throughout the run.9Analytical Biochemistry. Factors affecting polyacrylamide gel electrophoresis and electroblotting of high-molecular-weight myofibrillar proteins myofibrillar proteins Once they race ahead, proteins with free cysteine residues can re-form disulfide bonds with their neighbors during electrophoresis. This creates spurious high-molecular-weight bands that do not represent real protein complexes. Adding a small amount of reducing agent to the upper reservoir buffer, or chemically blocking free cysteines with an alkylating agent before the run, prevents this artifact.10Analytical Biochemistry. Factors affecting polyacrylamide gel electrophoresis and electroblotting of high-molecular-weight myofibrillar proteins myofibrillar proteins
Alkylation Before Running Gels
Alkylating cysteine residues before electrophoresis serves a second purpose beyond preventing disulfide re-formation. Unmodified free cysteines can react with acrylamide monomers in the gel itself, producing acrylamide adducts that add mass to the protein. If you plan to identify proteins by mass spectrometry afterward, these unintended modifications create confusing extra peaks. Alkylating cysteines ahead of time blocks this reaction.11PubMed. Modification of cysteine residues by alkylation. A tool in peptide mapping and protein identification
Alternative Buffer Systems
The classic Tris-glycine system works well for most proteins in the middle of the molecular weight range, but it has limits. Proteins smaller than about 10 kDa often do not resolve well because they migrate near the dye front, and very large proteins can be hard to separate. This has led to several modified buffer systems that still use SDS but swap out one or more of the other components.
Tricine-SDS-PAGE
Replacing glycine with tricine in the running buffer dramatically improves resolution of small proteins and peptides. Tricine-SDS-PAGE is the preferred system for separating proteins smaller than 30 kDa, and it can handle a range from about 1 to 100 kDa.12Nature Protocols. Tricine–SDS-PAGE One variation combines a tricine cathode buffer with modifications to gel ionic strength, achieving resolution from 3.5 to 200 kDa on a single low-percentage gel.13PubMed. An improved SDS-polyacrylamide gel electrophoresis for resolution of peptides in the range of 3.5-200kDa If you work with peptide fragments, small cytokines, or histones, tricine gels are usually a better choice than standard Tris-glycine.
Bis-Tris Gels
Bis-Tris systems operate at a lower, more neutral pH than Tris-glycine gels. This seemingly small change has real practical benefits. Chemical modifications that proteins can undergo during electrophoresis, such as deamidation and oxidation, happen at significantly lower rates in Bis-Tris gels compared to Tris-glycine gels.14PubMed. Models of protein modification in Tris-glycine and neutral pH Bis-Tris gels during electrophoresis: effect of gel pH The sample buffer used with commercial Bis-Tris gels also operates at a higher pH (around 8.5 versus 6.8 for Tris-glycine), which reduces protein degradation during the heating step.15PubMed. Models of protein modification in Tris-glycine and neutral pH Bis-Tris gels during electrophoresis: effect of gel pH For researchers who need to preserve the exact chemical state of their proteins, such as when looking for specific post-translational modifications, this matters.
When Proteins Migrate Anomalously
The whole point of SDS-PAGE is that migration correlates with molecular weight. But some proteins break this rule. Membrane proteins are the most common offenders. A database of 168 helical membrane proteins with known structures found that more than three-quarters of them migrate anomalously on standard SDS gels, running either faster or slower than their true molecular weight would predict.16PubMed. Correction factors for membrane protein molecular weight readouts on sodium dodecyl sulfate-polyacrylamide gel electrophoresis
The underlying cause has been traced to differences in how much SDS these proteins actually bind. Remember, the technique relies on a uniform coating of SDS. Helical membrane proteins, with their hydrophobic transmembrane regions, bind SDS at variable ratios rather than the standard amount. Research using hairpin sequences derived from the cystic fibrosis transmembrane conductance regulator (CFTR) showed that these fragments loaded anywhere from 3.4 to 10 grams of SDS per gram of protein, and their gel migration correlated strongly with how much SDS they carried.17PubMed Central. Detergent binding explains anomalous SDS-PAGE migration of membrane proteins In practical terms, a membrane protein might appear on a gel at a position suggesting it weighs 37 kDa when its actual mass is 55 kDa.
The cystine/glutamate antiporter SLC7A11 provides a concrete example. This highly hydrophobic protein consistently migrates at about 37 kDa on standard gels despite having a predicted mass of 55.4 kDa. Replacing nonpolar amino acids with polar ones reduced the hydrophobicity and restored normal migration. Using higher acrylamide concentrations (12% or 15% instead of lower percentages) also eliminated the shift.18PubMed Central. Hydrophobicity causes anomalous migration of cystine/glutamate antiporter SLC7A11 in SDS-PAGE with low acrylamide concentration If you are working with a membrane protein and the band keeps showing up at the wrong place, adjusting the gel percentage is one of the first things to try.
Glycoproteins are another class of proteins that often run anomalously. The sugar chains attached to glycosylated proteins do not bind SDS the way the polypeptide backbone does, so the charge-to-mass ratio is thrown off. Heavily glycosylated proteins typically run slower than expected and produce broad, fuzzy bands rather than the sharp lines you see with unmodified proteins.
Staining and Visualization
After electrophoresis, the proteins in the gel are invisible. You need a stain to see them. The two most common options are Coomassie blue and silver staining. Coomassie is simpler, faster, and gives reasonably quantitative results, but it requires a minimum amount of protein per band to produce a visible signal, usually in the low nanogram range. Silver staining is far more sensitive, detecting down to the femtomole level, but the protocol is more involved and can be less reproducible.19PubMed. Silver- and Coomassie-staining protocols: detection limits and compatibility with ESI MS Interestingly, proteins stained with sulfonic acid dyes (like Coomassie) before silver staining show enhanced silver detection, up to eight-fold improvement, suggesting that the dye provides nucleation sites for silver deposition.20PubMed. Mechanism studies of coomassie blue and silver staining of proteins
Fluorescent stains have become increasingly popular as an alternative. Products like SYPRO Ruby offer sensitivity approaching silver staining with the linear dynamic range and ease of use closer to Coomassie. They also avoid the fixation chemistry of silver staining, which can interfere with downstream analyses like mass spectrometry. The choice of stain often depends on what you plan to do with the gel afterward, not just how much protein you have.
From Gel to Western Blot and Mass Spectrometry
SDS-PAGE is often just the first step. Western blotting transfers the separated proteins from the gel onto a membrane, where specific proteins can be detected with antibodies. The transfer typically uses a Tris-glycine transfer buffer with methanol added to help proteins bind to the membrane. For large proteins, though, methanol is less critical. Testing on a range of proteins of different sizes showed that methanol could be reduced or even removed from the transfer buffer without hurting the signal for large proteins, while medium and small proteins benefited from around 10% methanol.21PubMed Central. Optimization of western blotting for the detection of proteins of different molecular weight
Mass spectrometry presents a different challenge. SDS is an exceptionally effective detergent for solubilizing and separating proteins, but it is also notoriously harmful to mass spectrometry instruments and suppresses ionization of peptides at very low concentrations. Before proteins or peptides can be analyzed by mass spectrometry, the SDS must be removed almost completely. Multiple depletion strategies exist, and comparative testing showed that filter-aided sample preparation (FASP) and in-gel digestion achieved the highest SDS removal, over 99.99%. Other methods like acetone precipitation and strong cation exchange also bring SDS below the threshold where it interferes with the analysis.22PubMed. Comparison of sodium dodecyl sulfate depletion techniques for proteome analysis by mass spectrometry Various cleanup approaches including bead-based purification, filtration, and protein precipitation are used routinely in proteomics workflows to handle this problem.23PubMed Central. Optimal Conditions for Carrying Out Trypsin Digestions on Complex Proteomes: From Bulk Samples to Single Cells
Native SDS-PAGE and Preserving Protein Activity
Standard SDS-PAGE deliberately destroys a protein’s native shape. That is the price of separating by size. But researchers sometimes want the sizing advantage of SDS while keeping proteins at least partially functional. A modified protocol called native SDS-PAGE (NSDS-PAGE) uses gentler conditions: lower SDS concentrations, no boiling, and no reducing agents. Under these conditions, many proteins retain their multimeric associations and even their enzymatic activity. In one study, seven of nine model enzymes remained active after NSDS-PAGE, and proteins migrated as their intact complexes rather than dissociated subunits. Zinc retention in proteins jumped from 26% under standard denaturing conditions to 98% under the native SDS protocol.24PubMed Central. Native SDS-PAGE: High Resolution Electrophoretic Separation of Proteins With Retention of Native Properties Including Bound Metal Ions
This is a niche technique, not a general replacement for standard SDS-PAGE. The migration patterns are harder to interpret because proteins are running as complexes of varying shape, and the relationship between migration distance and molecular weight is less predictable. But for studying metal-binding proteins or confirming that a protein maintains its quaternary structure, NSDS-PAGE fills a gap between fully denaturing SDS-PAGE and Blue Native PAGE, which avoids SDS entirely.
Capillary Electrophoresis as a Modern Alternative
Slab gels are labor-intensive. You pour the gel, load each well by hand, run the current, stain, destain, and image. For quality control in the biopharmaceutical industry, where the same assay might be run thousands of times on therapeutic antibodies, a faster and more automated approach is appealing. Capillary electrophoresis with SDS (CE-SDS) uses the same basic principle, SDS-coated proteins separated by size, but in a narrow capillary rather than a slab gel. Detection happens automatically as proteins pass a detector. An interlaboratory validation study showed that CE-SDS performs reliably as a purity and size-heterogeneity assay for monoclonal antibody therapeutics across different laboratories.25PubMed. Interlaboratory method validation of capillary electrophoresis sodium dodecyl sulfate (CE-SDS) methodology for analysis of mAbs CE-SDS will not replace slab gels in research labs any time soon, since cutting out a band from a capillary for further analysis is not straightforward, but for standardized quality testing it is a genuine step forward.
Safety Considerations in the Lab
Of the chemicals involved in an SDS-PAGE experiment, acrylamide is the one that deserves the most respect. Before polymerization, acrylamide monomer is a documented neurotoxin in humans based on occupational exposure studies, and animal research raises concerns about reproductive toxicity and carcinogenicity.26ACS Publications. Chemistry, biochemistry, and safety of acrylamide. A review. Once the gel is polymerized, the crosslinked matrix is essentially inert, but handling unpolymerized acrylamide solutions demands gloves and caution. Pre-cast gels, which now dominate many labs, reduce this exposure substantially since you never handle the liquid monomer. SDS itself is an irritant, particularly to eyes and skin, but far less hazardous than acrylamide. The electrical currents used during electrophoresis are the other safety consideration. Power supplies can deliver enough voltage and current to be dangerous if you open the gel apparatus while it is running, so all modern gel boxes include safety interlocks that cut power when the lid is removed.
Beta-mercaptoethanol, the reducing agent many protocols call for, is volatile and has a distinctive, potent stench often compared to rotten eggs. Working with it outside a fume hood is unpleasant and inadvisable, since it is toxic by inhalation. DTT is a less odorous alternative that serves the same chemical function. Methanol, used in western blot transfer buffers, is another common toxin in the SDS-PAGE workflow, though recent work suggests it can often be reduced or eliminated from transfer buffers without sacrificing signal quality for many proteins.27PubMed Central. Optimization of western blotting for the detection of proteins of different molecular weight