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Why my proteins are migrating like this on SDS-page?

Why my proteins are migrating like this on SDS-page?


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I'm trying to make a SDS-page of total proteins extracted from algae but I'm encountering two problems. First, when I depose my proteins in the wells the sample goes down and then it starts to move up the sides of the well. Second the result of my run looks like this:

so my bands are not visible and I have like a vertical smear on the sides of the lane.

The buffer used to extract proteins contains Tris, NaCl, EDTA DTT and Glycerol and my loading buffer is a 5x commercial dye with B-mer.

Thank you!


It's a guess, as I haven't run a algae sample on SDS-PAGE, but your migration problems might be due the algal polysaccharides that are present in your sample. Check if your protein purification protocol is specific for algal proteins and see if there is any step dealing with the algal polysaccharides.


Analysing protein using SDS-page

Hello, am a mature student hoping to tap into your lovely brains. Am trying to write a paper but I am struggling to analyse protein using an SDS-page. Its about purification of protein. If I have lots bands that are not my protein of interest, where might these protein bands come from and why might my process fail to pick it up and how can it be rectified?

I shall be grateful if anyone can explain it to me or point me into any directions of useful materials

I think this could have something to do with the sample you're running in the first place. If it's something like a total cell protein extract then that would have other proteins besides your protein of interest (PoI), which can then show up on your PAGE. If your PoI is the same mol wt as another random protein, it will definitely hinder analysis/purification.

I would say if you know the size and mass or other properties of your PoI you could purify your sample before running a gel. Chromatographic techniques work, dialysis works sometimes.


Detergent binding explains anomalous SDS-PAGE migration of membrane proteins

Migration on sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) that does not correlate with formula molecular weights, termed “gel shifting,” appears to be common for membrane proteins but has yet to be conclusively explained. In the present work, we investigate the anomalous gel mobility of helical membrane proteins using a library of wild-type and mutant helix-loop-helix (“hairpin”) sequences derived from transmembrane segments 3 and 4 of the human cystic fibrosis transmembrane conductance regulator (CFTR), including disease-phenotypic residue substitutions. We find that these hairpins migrate at rates of −10% to +30% vs. their actual formula weights on SDS-PAGE and load detergent at ratios ranging from 3.4–10 g SDS/g protein. We additionally demonstrate that mutant gel shifts strongly correlate with changes in hairpin SDS loading capacity (R 2 = 0.8), and with hairpin helicity (R 2 = 0.9), indicating that gel shift behavior originates in altered detergent binding. In some cases, this differential solvation by SDS may result from replacing protein-detergent contacts with protein-protein contacts, implying that detergent binding and folding are intimately linked. The CF-phenotypic V232D mutant included in our library may thus disrupt CFTR function via altered protein-lipid interactions. The observed interdependence between hairpin migration, SDS aggregation number, and conformation additionally suggests that detergent binding may provide a rapid and economical screen for identifying membrane proteins with robust tertiary and/or quaternary structures.

Determination of protein molecular weight (MW) via polyacrylamide gel electrophoresis (PAGE) in the presence of sodium dodecyl sulfate (SDS) is a universally used method in biomedical research. The technique requires that the proteins under investigation on the gel and the proteins used for MW calibration adopt equivalent shapes after SDS treatment. This condition is achieved not by the complete unfolding of the proteins but rather by the aggregation of SDS molecules at hydrophobic protein sites to induce “reconstructive denaturation,” where proteins adopt a conformational mixture of α-helix and random coil [reviewed in (1)]. The resultant protein-detergent complexes consist of helical SDS-coated polypeptide regions spatially separated by flexible and uncoated linkers, termed “necklace and bead” structures (2). Individual sizes of the micellar “beads” can vary and seem to be determined by amino acid sequence (3). A further proviso for MW estimation by SDS-PAGE is a consistent amount of detergent binding among proteins (4). Maximum SDS-binding levels are generally quoted at 1.4 g SDS/g protein, following the work of Reynolds and Tanford that identified this weight ratio as nearly invariant for a variety of globular and erythrocyte ghost proteins (4). Subsequent investigations of SDS loading have refined this saturation value for globular proteins to as much as 1.5–2 g detergent/g protein (5).

Peptide segments that contain an abundance of hydrophobic residues—such as the transmembrane (TM) sequences of membrane proteins—have helical regions expected to be embedded within the SDS micelle interior, while amphipathic segments common to globular polypeptides orient their hydrophobic faces toward the detergent tails and hydrophilic portions toward the micelle surface [reviewed in (1)]. Membrane proteins in some cases load 2-fold greater amounts of SDS than globular polypeptides examples include the 3.4 g SDS/g stoichiometry of human erythrocyte membrane glycoprotein glycophorin (6) and 4.5 g SDS/g bound to CP-B2, a membrane protein from R. rubrum chromatophores (7). Large SDS/protein aggregate stoichiometry is nevertheless not universal among membrane-soluble polypeptides: intact cytochrome b5 binds 1.2 g SDS/g protein (8) the KcsA potassium channel tetramer 0.7 g SDS/g (9) and the human red cell glucose transporter Glut1 1.7 g SDS/g (10).

Protein tertiary structure may affect both detergent-loading levels and polypeptide-SDS-PAGE migration rates. Disulfide bonds, for example, reduce SDS binding to globular proteins by up to 2-fold (11), and have been linked to the anomalously fast migration of unreduced vs. reduced lysozyme, presumably because the intact disulfide bonds impose a more compact shape on the enzyme (12). Similarly, in a previous study with helix-loop-helix or “hairpin” model membrane proteins based on portions of the TM domain of the human cystic fibrosis transmembrane conductance regulator (CFTR), we noted that increased SDS-PAGE migration rates were traceable to a compact S-S bridged conformation (13). The E. coli β-barrel membrane protein OmpA also binds less SDS as a folded than fully denatured species (14, 15), and moves through gels according to structural compactness, with the folded form migrating faster than the completely denatured protein (16).

While these overall observations suggest that there may be a specific relationship between protein structure and SDS loading, a unifying source for atypical membrane protein SDS-PAGE migration has not emerged, and questions as to whether this detergent “denatures” membrane proteins and/or their oligomeric forms are often posed (17). In the present work, we establish a quantitative relationship between gel migration behavior and SDS aggregate stoichiometry using a library of helical hairpins derived from CFTR. We find that gel shifts strongly correlate (R 2 = 0.8) with changes in the SDS-loading capacity of these miniature membrane proteins, indicating that altered detergent binding explains anomalous SDS-PAGE behavior. Our results reveal a distinction between two CF-phenotypic mutants studied: V232D binds significantly less SDS than the WT protein while P205S SDS binding is indistinguishable from WT, indicating that CFTR dysfunction may arise variously as a consequence of altered protein-lipid interactions or via altered intra-protein contacts.


Smeared gels &ndash example 3

Smearing can be a normal consequence of running membrane-associated proteins with a high lipid content in a discontinuous gel. In discontinuous PAGE, sample proteins are super-concentrated by a combination of two factors. First, the rate of migration is suddenly slowed as the sample moves from a non-restrictive stacking gel to a separating gel that restricts movement. Second (and more important), a pH change affects the electrophoretic mobility of proteins in the sample, causing the sample to check up abruptly. A sample of a half cm or so in height becomes compressed into a layer of bands a few micrometers thick, and the local concentration of protein goes up considerably.

Membrane-associated proteins tend to precipate at lower concentrations, thus lanes containing such proteins often have a dark band of precipitated proteins at the very top. As electrophoresis proceeds the precipates re-dissolve and enter the gel continuously, thus forming a continuous dark background of unresolved polypeptides. Many of the membrane samples shown on these gels will show that characteristic.

The resolved bands in the membrane protein lanes (4 and 7) appear to consist of similar amounts of protein, however in lane 4 the capacity of the gel for at least some of the membrane proteins was exceeded, resulting in a dark smear.


Help for understanding page/ sds page electrophoresis on native proteins

Thanks all, I have to write a research paper on electrophoresis and I'm trying to understand what you learn with running page on a native protein. I understand that it separates proteins via mass/ charge ratio, but why can you suddenly know the mass when they run it into sds page? Is it because now all the proteins have the same mass/ charge ratio when they go through sds page?

SDS denatures the protein, negating any effects of the folded protein. This means that the protein can be considered a linear poltpeptide with no high order structure that can lead to variable rates of migration. SDS also interacts directly with the protein, and the large negative charge of the detergent mostly masks or camels the individual charges in the protein, giving the protein a fairly uniform charge/mass ratio.

So, the SDS detergent removes the factors that lead to variability in migration rate, leaving molecular weight as the only variable that distinguishes between two different proteins, allowing their separation by size alone.


Disrupting dimers in SDS-PAGE

I expressed a his tagged protein in ecoli and isolated by nickel column. When I ran a SDS-PAGE of the elutions I got a major band of the expected size and another major band of about double that size. My protein of interest is well characterized as forming dimers through electrostatic interactions, so I suspect that the heavy band is just a dimer of my protein. I want to do an experiment to test this, so I looked at some approaches to disrupt protein dimerization before running the PAGE:

This paper describes hour long boiling times, guanidine HCl, and urea added to the loading buffer to disrupt dimers. It looks like guanidine HCl is not compatible with SDS-PAGE and needs to be removed in advance of running the gel. In the supplementary data they describe using urea to disrupt dimers, but don't describe removing the urea in any way.

Long story short, my question is: If I put urea in my loading buffer in an attempt to dirupt a suspected protein dimer, do I have to remove the urea in any way before running the sample of an SDS-PAGE


Resources

Protein Standards Selection Guide (PDF 291 KB)

For quick guidance on choosing the best standard for your application.

Little Book of Standards (PDF 5.12 MB)

Complete reference information for all of Bio-Rad&rsquos protein standards and nucleic acid standards.

Protein Gel Migration Charts

View banding patterns for Bio-Rad protein standards on Bio-Rad and competitor gels.

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Electrophoresis Guide (PDF 8.35 MB)

Theory and techniques, Bio-Rad products, tips and troubleshooting.

Protein Blotting Guide (PDF 7.11 MB)

Details on blotting technology, available products, and tips and techniques.


Peggy's Intern Diary

Yesterday (January 31st, 2013), I went to RPI and worked with Eun Ji. But before that, I stopped by the Union Building, got a bit lost, and picked up my ID card (FINALLY. whew. ).

When we were heading to the lab, I told Eun Ji that I just did DNA electrophoresis in my bio class. Then, she said that she was just going to do an electrophoresis to confirm her previous data and asked me if I want to do it with her (Of course. D)!

Gel electrophoresis is a coomon used technique in biotechnology. It can separate and determine the size of the DNA fragments (cut by restriction enzymes) according to their moving rate. First we load the DNA fragments into a gel well. Then, a positive electric current passes through the other end of the gel. Since DNA fragments are negatively charged, they will slowly migrate to the positive end of the gel at a rate that are proportional to their sizes (the smaller the faster).

However, instead of doing DNA electrophoresis, Eun Ji and I were doing the protein electrophoresis, or SDS-PAGE (Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis). Unlike DNA, protein has a very specific 3D shape where all the amino acids are tightly packed. Thus, first of all, we have to make it into a straight chain of polypeptide by adding DTT or other substance, which break the hydrogen bonds between amino acids. Then, we have to add SDS to make the positive-charged protein negative charged because gel electrophoresis works for substance to move from negative end to positive end.

Yet, Eun Ji and I were working on the step before launching electrophoresis - the protein assay , a
simple and accurate procedure for determining concentration of a protein sample. This step is critical because when we are loading the protein into the gel, we want each lane to have the same amount of protein (μg) , not the same volume (μl). Those protein samples are extracted from E. coli. However, while most proteins exist in cytosol, some are in cellar compartments such as inclusion body that are hard to extract proteins. The reason why she wanted to double check her is because in order to get the protein, she has to add SDS to break the inclusion body, which would also affect the targeted protein itself.

Therefore, first of all, Eun Ji got all the materials and the samples she wanted to re-investigate Also, we want to make sure that we do wear gloves because since SDS affects protein, it can also destroy our skin (It smells really bad, and being exposed too long under it can cause health effects, like I got an headache for just smelling for a few seconds). We divided the 96-well plates into a few sections.

The next step would be to put the plate into spectrophotometer to measure the absorbance and calculate the concentration. However, because of the shortage of time, I wasn't able to get to this step:( Nonetheless, Eun Ji explained the following steps, converting volume, loading and running gel, and analyzing gel, in which the whole process may take up to 2 days!:O To get the specific target protein even involves more steps!

Later in the day Eun Ji sent me the data including some comments. In general, I did fine on it, but compared to hers, my numbers fluctuate a lot. One thing I would need to work on is the pipette technique, especially when dealing with such tiny amount of substance. Meanwhile, I asked Eun Ji why SDS-PAGE runs vertically where as DNA eletrophoresis runs horizontally. She was kind of surprise because she has never thought about this before!! In the email, she explained that SDS-PAGE is consist of two layers (upper layer (stacking gel) and lower layer (separating or running gel), and vertical direction just makes the process easier. Another reason that I found online is that since the gel runs best in anoxic environment, it's better to have the protein in sealed b/w 2 layers rather than open layer like that in horizontal one.


Why is SDS used in PAGE?

SDS is a detergent that is present in the SDS-PAGE sample buffer where, along with a bit of boiling. It&rsquos a reducing agent to break down protein-protein disulphide bonds that disrupts the tertiary structure of proteins, and brings the folded proteins down to linear molecules. SDS coats the protein with a uniform negative charge, which masks the intrinsic charges on the R-groups. SDS binds fairly uniformly to the linear proteins (around 1.4g SDS/ 1g protein), meaning that the charge of the protein is now approximately proportional to its molecular weight. SDS is also present in the gel to make sure that once the proteins are linearized and their charges masked, they stay that way throughout the run.

The dominant factor in determining an SDS-coated protein is it&rsquos molecular radius. SDS-coated proteins have been shown to be linear molecules, 18 Angstroms wide and with length proportional to their molecular weight, so the molecular radius (and hence their mobility in the gel) is determined by the molecular weight of the protein. Since the SDS-coated proteins have the same charge to mass ratio, there will be no differential migration based on charge.


Electrophoresis

Electrophoresis is one of the most important techniques used by molecular biologists. To name only a few applications, deoxyribonucleic acid (DNA) electrophoresis is used to map the order of restriction fragments within chromosomes , to analyze DNA variation within a population by restriction fragment length polymorphisms (RFLPs), and to determine the nucleotide sequence of a piece of DNA.

Electrophoresis refers to the migration of a charged molecule through a restrictive matrix , or gel, drawn by an electrical force. As the force drags the molecule through the gel, it encounters resistance from the strands of the gel, retarding its rate of migration. In gel electrophoresis, larger molecules migrate more slowly than smaller ones, and so the distance of migration within a gel can be used to determine a molecule's size.

Although it is possible to separate whole chromosomes using specialized electrophoresis techniques, DNA that is to be analyzed by electrophoresis is usually cut into smaller pieces using restriction enzymes . Fragments of DNA prepared by treatment with restriction enzymes are commonly separated from one another, and their sizes determined, using a gel of agarose electrophoresis, a complex carbohydrate . DNA is negatively charged due to the phosphodiester bonds that join the individual nucleotide building blocks. DNA will therefore electrophorese toward the positive electrode when placed in an electrical field. To visualize the results after electrophoresis, the gel is soaked in a solution that causes DNA to fluoresce when exposed to ultraviolet light.

Treatment of the DNA sample with multiple restriction enzymes in various combinations enables the researcher to generate a restriction map of the original DNA fragment, which identifies the sites at the DNA where the restriction enzymes are.

Many research questions require a detailed analysis of one specific DNA fragment in a complex mixture. In such cases, a radioactive DNA probe can be used to identify the fragment based on its nucleotide sequence. The method, known as hybridization, is based on the rules of complementary base pairing (A bonds to T, G bonds to C). A probe is designed whose sequence is complementary to the piece of DNA to be detected. The gel-separated DNA is first transferred to a nylon membrane using a technique called a Southern blot.

During the blotting procedure, the strands within the DNA double helix are separated from each other, or denatured, by treatment with a base. Because double-stranded DNA is more stable than single-stranded, during the hybridization the single-stranded probe will locate and bind to the single-stranded gel-separated fragment with complementary sequence. Being fluorescent or radioactive, the position of the probe can be determined using photographic methods. The target sequence can then be removed by cutting at the piece of the gel that contains it.

The most common technique for determining DNA sequence is the Sanger method, which generates fragments that differ in length by a single nucleotide. High-resolution polyacrylamide gel electrophoresis is then used to separate the fragments and to allow the sequence to be determined.

Electrophoresis of ribonucleic acid (RNA) is an integral procedure in many studies of gene expression . RNA is isolated, separated by electrophoresis, and then the gel-separated RNA fragments are transferred to a nylon membrane using a technique called a Northern blot. Hybridization with a single-stranded DNA probe is then used to determine the position of a specific RNA fragment.

DNA and RNA are relatively simple in terms of structure and composition. Proteins , however, are composed of twenty different amino acids in various combinations, and proteins vary significantly in their three-dimensional structure. The composition of amino acids will affect the charge on the protein, which ultimately will affect its electrophoretic behavior. The shape of a protein similarly will affect its rate of migration. As a result, a specialized technique, SDS-polyacrylamide gel electrophoresis (SDS-PAGE), is usually used to analyze proteins. In this method, protein samples are heated and then treated with the detergent sodium dodecyl sulfate (SDS). Proteins treated in this way are unfolded, linear, and uniformly coated by negatively charged detergent molecules. The rate of migration of treated proteins is inversely proportional to the logarithm of molecular weight. Following electrophoresis, the protein in the gel can be stained to visualize all the proteins in a sample, or the proteins in the gel can be transferred to a nylon membrane (Western blot) and specific ones detected with the use of enzyme -linked antibodies.

Regardless of the macromolecule being studied, gel electrophoresis is a crucial technique to the molecular biologist. Many scientific questions can be answered using electrophoresis, and as a result an active molecular biology research lab will have several benches that are devoted to the required specialized reagents and equipment.


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