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In SDS-PAGE, why are the sample buffer and running buffer different in concentration?

In SDS-PAGE, why are the sample buffer and running buffer different in concentration?


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I'm currently trying to understand the concepts of SDS-PAGE. Every protocol I've read says to use 5X for sample buffer and 1X for running buffer, but I don't really understand for what reason? What are the negative consequences if I don't?


The reason why the sample buffer is more concentrated (typically 2x or 5x depending on your protein concentration) is its dilution when you mix it with the sample. You mix 4 parts of your sample and 1 part of 5x sample buffer, so that the final concentration of this buffer is 1x.

If you use 1x sample buffer together with the same amount of sample, the final concentration of the sample buffer would be 0.5x, if you do the same 4+1, it would only be 0.2x in the end.

This dilution is too strong and the buffer will not fulfill its purpose, the complete denaturation of all proteins in the sample, so that their distribution on the gel depends only on their molecular weight and not also on their tertiary/quarternary structure.

Using a 5x buffer allows you to add a lot of sample when diluting it to 1x without changing the concentration of the protein too much. It also allows to load larger amounts of sample compared to 2x buffer, interesting when the protein concentration is low.

The running buffer is not further diluted by adding sample, so that it has to be diluted to 1x in order to have the right ionic strength. Since these are often used in big quantities in the lab, it is more convenient to prepare concentrated buffers in smaller quantities and dilute them before usage than preparing big tanks of buffer.


Why BOIL.. samples for sdspage - sample + sample buffer + 'BOIL' 2 mins (Jul/31/2009 )

the standard protocols for sdspage involve addition of sample buffer to the protein sample and then boiling the samples for 2 mins.. before loading on to the gel.
is there some specific reason for boiling the samples.
is this specific for reducing samples? or is applicable to both nonreducing and reducing conditions.

why and for how long should the samples be boiled. waht decides the time?

(try not boiling - you'll see what i mean)

High temperature denatures protein and promotes denaturation by SDS, it also breaks disulfide bonds and promotes (accelerates) reaction with reducing agenst (eg. DTT, bME).

K.B. on Jul 31 2009, 07ᛝ AM said:

i tried no boiling n boiling for different times.. 2 min 5 min n so on..
when i donot boil my samples i donot see a low molecular weight band.. and when i boil.. it increases with increase in time for boiling.

also, higher the amount of sds in the sample buffer.. lesser the low molecular weight band intensity is..

any ideas.. ?
how do i know whether this band truely exists or not?

Boiling for too long may result in protein degradation. This may explain the band.

As for the sds.. I don't know!

alicee! on Aug 2 2009, 10ᛛ PM said:

K.B. on Jul 31 2009, 07ᛝ AM said:

i tried no boiling n boiling for different times.. 2 min 5 min n so on..
when i donot boil my samples i donot see a low molecular weight band.. and when i boil.. it increases with increase in time for boiling.

also, higher the amount of sds in the sample buffer.. lesser the low molecular weight band intensity is..

any ideas.. ?
how do i know whether this band truely exists or not?

1. This is a matter of optimization (i.e. you try out many conditions - no boiling, some boiling, more boiling, less SDS, more SDS etc). Some proteins, depending upon their physical properties and chemical make-up, work best with boiling, some don't (especially the membrane proteins).

2. As a rule of thumb (not always applicable), the higher the MW, the more boiling time, more SDS concentration is required.

1. If it is a specific product and a specific band coming up with only your specific antibody and not with IgG,

2. If it is the expected size,

3. If it vanishes with well-controlled competetion experiments,

4. and lastly, if nothing else is sure, if you sequence the protein,

cellcounter on Aug 4 2009, 07ᚺ AM said:

alicee! on Aug 2 2009, 10ᛛ PM said:

K.B. on Jul 31 2009, 07ᛝ AM said:

i tried no boiling n boiling for different times.. 2 min 5 min n so on..
when i donot boil my samples i donot see a low molecular weight band.. and when i boil.. it increases with increase in time for boiling.

also, higher the amount of sds in the sample buffer.. lesser the low molecular weight band intensity is..

any ideas.. ?
how do i know whether this band truely exists or not?

1. This is a matter of optimization (i.e. you try out many conditions - no boiling, some boiling, more boiling, less SDS, more SDS etc). Some proteins, depending upon their physical properties and chemical make-up, work best with boiling, some don't (especially the membrane proteins).
yes but different conditions do show different results.. it is difficult to say which one of these is a true result.
meaning.. no boiling.. vs. boiling 2 mins. 5 mins.. n up to 10 mins.. the lower molecular band increases in intensitiy. it is difficult to say if this is already present in the sample (some how bound/associated with the principal protein) and is dissociated in presence of the sample buffer and the boiling treatment.
same is with the sds.. higher sds to protein ratio.. lower the low MW band intensity. it is difficult to say whihc is the true result.
it would be inappropriate to not report the impurity if it is present. also the true quantity of the impurity needs to be known but the procedural problems are not answering these questions.

2. As a rule of thumb (not always applicable), the higher the MW, the more boiling time, more SDS concentration is required.
mine is near 19 kDa. I see an extra band near 14-15 kda.

1. If it is a specific product and a specific band coming up with only your specific antibody and not with IgG,
Yes it is product specific, confirmed with an immunodetection on a western blot.

2. If it is the expected size,

3. If it vanishes with well-controlled competetion experiments,

4. and lastly, if nothing else is sure, if you sequence the protein,

size of the protein: 19kda
low Mw band obtained: 14 kDa.
mostly with intact N terminal sequence. may be truncated at C terminal. Am unable to sequence this.
Not able to answer if this a true band or generated due to sample treatment.
If I knew the site of cleavage, how would that help confirm how much of it is present.

Any protein sample in case of reducing SDS PAGE, sample has to boiled with SSB for atleast 5 min.The boiling will enhance the action of SDS,therefore the tertiary structure of proteins becomes primary and break in disulfide bond by reducing agent in SSB will lead form single band for separation based on charge and molecular weight in gel.

How about spinning down after the boil, how long do you spin for and at what temperature and RPM? Are the benefits to spinning longer?

Bill Nye on Sep 1 2009, 04ᚻ PM said:

just to spin down any ppt formed during sample preparation. I usually spin at full speed (13,000rpm) for 5min and load the supernatant.


Polyacrylamide Gel Electrophoresis

Polyacrylamide gel electrophoresis (PAGE) is probably the most common analytical technique used to separate and characterize proteins. A solution of acrylamide and bisacrylamide is polymerized. Acrylamide alone forms linear polymers. The bisacrylamide introduces crosslinks between polyacrylamide chains. The 'pore size' is determined by the ratio of acrylamide to bisacrylamide, and by the concentration of acrylamide. A high ratio of bisacrylamide to acrylamide and a high acrylamide concentration cause low electrophoretic mobility. Polymerization of acrylamide and bisacrylamide monomers is induced by ammonium persulfate (APS), which spontaneously decomposes to form free radicals. TEMED, a free radical stabilizer, is generally included to promote polymerization.

The gels are usually prepared with the top portion of the gel under the sample wells made less dense than the remainder of the gel below that is intentionally made denser. The top portion is referred to as the &ldquostacking gel&rdquo and the lower portion is termed the &ldquorunning gel&rdquo or &ldquoseparating gel&rdquo. The purpose of the stacking gel is to concentrate all of the different sized proteins into a compact horizontal zone by sandwiching them between a gradient of glycine molecules above and chloride ions below. This way most of the proteins will enter the denser resolving gel simultaneously before they begin to migrate downwards at different rates based on their size. This way, the bands are much clearer and better separated for visualization and analysis. Without the stacking gel, the proteins will produce a long smear through the resolving gel instead of tight distinct bands for us to analyze.

Figure 1. PAGE gel. A protein first runs through the stacking gel, where the samples spread out. Once a protein reaches the separating gel, the proteins pack together in tight bands. As they move through the resolving gel they separate by size.

SDS-PAGE

Sodium dodecyl sulfate (SDS) is an amphipathic detergent. It has an anionic head group and a lipophilic tail. It binds non-covalently to proteins, where roughly one SDS molecule is attracted to every two amino acids. SDS causes proteins to denature and disassociate from each other (excluding covalent cross-linking) and essentially unravel into linear molecules. It also confers negative charge. In the presence of SDS, the intrinsic charge of a protein is masked. During SDS-PAGE, all proteins migrate towards the anode (the positively charged electrode). SDS-treated proteins have very similar charge-to-mass ratios, and similar shapes. During PAGE, the rate of migration of SDS-treated proteins is effectively determined by their unfolded length, which is related to their molecular weight.

Figure 2: A protein surrounded by the SDS molecules.


Amounts to load

Polyacrylamide has a limited capacity for protein. Overloading results in precipitation and aggregation of proteins, producing streaks and smears. Underloading results in complete disappointment, as one may detect only the most abundant of polypeptides, if that. The objectives of sample preparation are to put the proteins into a denaturing buffer, rendering them suitable for electrophoresis, and to adjust the concentrations of sample so that an appropriate amount of protein can be loaded onto a gel.

We get the best results if we load 10 µl of a 2 mg/ml final concentration of denatured protein per sample well. While some of the more concentrated proteins will be overloaded, we will detect bands that represent the less common ones. A typical mini-gel well holds 10 µl easily, and perhaps 20 µl or more if the well dividers are in good shape.

We will dilute all samples to a predetermined concentration and volume before mixing with the denaturing buffer. Efficient laboratory personnel divide responsibilities, so that while gels are polymerizing they are preparing the samples themselves, to volumes that are at least double the minimum needed to fill the sample wells. Such people start their work prepared with calculations of the volumes of sample, water, and 2x concentrated sample buffer they need in order to prepare each of their samples for electrophoresis.

To completely denature the samples we heat them in a steaming water bath for at least 10 minutes. Standards for molecular weight determination are prepared the same way. They are expensive, and although the suppliers give instructions for mixing, it is usually necessary to test them and to make adjustments before relying on them for internal calibration of an important gel. A "dirty" sample (containing a lot of particulate matter) should be centrifuged just before loading. However, samples containing soluble proteins only and samples from a typical blood fractionation are so "clean," that centrifugation is not necessary.


  • 62.5 mM Tris-HCl pH 6.8
  • 2.5 % SDS
  • 0.002 % Bromophenol Blue
  • 0.7135 M (5%) β-mercaptoethanol
  • 10 % glycerol

To make 10 ml of 10x stock

  • In 70 % glycerol / 30 % water, dissolve the following:
    • 0.606 g Tris-base
    • 2.5 g SDS
    • Adjust the pH using pH indicator strips to 6.8
    • Add 2 mg of Bromophenol Blue and make sure the powder is completely dissolved
    • Adjust the final volume to 10 ml with 70 % glycerol / 30 % water before storing at -20°C.
    • Add the appropriate volume of a β-mercaptoethanol 100% stock to your samples just before denaturing them at 95°C.

    It’s All About the Glycine

    So what’s up with glycine?
    A lot. It is the key to the discontinuous buffer system. It is the ionic state of glycine that really allows the stacking buffer to do its thing. Glycine is an amino acid with the chemical formula NH2-CH2-COOH. The charge of its ion is dependent on the pH of the solution that it is in. In acidic environments, a greater percentage of glycine molecules become positively charged. At a neutral pH of around 7, the ion is uncharged (a zwitterion), having both a positive charge and a negative charge. At higher pHs, glycine becomes more negatively charged.

    What does glycine’s charge have to do with the stacking layer?
    Everything. Glycine is in the running buffer, which is typically at a pH of 8.3. At this pH, glycine is predominately negatively charged, forming glycinate anions. When an electric field is applied, glycinate anions hit the pH 6.8 stacking buffer, and change to become mostly neutrally charged glycine zwitterions. That means they move slowly through the stacking layer toward the anode due to their lack of charge.

    By contrast, the Cl- ions (from the Tris-HCl in the gel) move at a faster rate towards the anode. When the Cl- and glycine zwitterions hit the loading wells with your protein samples, they create a narrow but steep voltage gradient in between the highly mobile Cl- ion front (leading ions) and the slower moving, more neutral glycine zwitterion front (trailing ions). The electromobilities of the proteins in your sample are somewhere in between these two extremes, and so your proteins are concentrated into this zone and herded through the stacking gel between the Cl- and glycine zwitterion fronts.

    What happens to glycine zwitterion in the resolving layer?
    It gets real negative, real fast. When the Cl- and glycine zwitterion fronts hit the resolving layer at a pH of 8.8, the glycine ions gain a lot of negative charges. They are no longer predominately neutral and take off towards the positively charged anode as glycinate anions. Unaffected by polyacrylamide, they speed past the protein layer, depositing the proteins in a tight band at the top of the resolving layer.


    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.


    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.


    What is the difference between constant current, constant voltage, and constant power?

    Because the conditions of the gel, buffer, and sample can change during the electrophoresis steps, most modern power packs offer a variety of options for maintaining constant voltage, constant current (amps), and constant power (watts). Before going to some recommended settings, here’s a quick refresher on the basics of electric circuits:

    Voltage (V) — the difference in electrical potentials between two charges — is the primary parameter for defining the speed that your protein will move through a gel during SDS-PAGE. If you think about electricity like a water tower, voltage is the water pressure created by placing the water at some height. The higher the voltage, the higher the electric “pressure” and the faster your proteins will run.

    Current (I) refers to the flow of electric charge past a point in a circuit. Using the same water analogy as above, the current is the rate that water flows through the pipe.

    Power (P) — generally defined as work done per unit of time — is simply equal to the voltage multiplied by the current, which can be written as such:

    One additional parameter to consider is the resistance (R), which (as the name implies) is a measure of how difficult it is for the charge to pass through a conductor. In Western blotting, resistance is the measure of how efficiently the ions in your SDS-PAGE buffer allow charge to “flow” through the gel.

    Resistance is related to voltage and current by Ohm’s law:


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      SDS-PAGE
      Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) in the presence of a reducing agent (2-mercaptoethanol) is a technique for the separation of polypeptide subunits according to their molecular weight. The protocol involves denaturing the protein sample by heating it in the presence of SDS and a reducing agent. SDS will bind to the protein causing it to unfold, whereas the reducing agent will reduce the intramolecular and intermolecular disulfide bonds. The binding of SDS by the protein confers a net negative charge and the denatured polypeptide will migrate through a gel of known percent acrylamide in the presence of an applied electric field towards the positive electrode (anode). After the electrophoresis is complete, the gel is stained with Coomassie" Blue R-250 to visualize the polypeptide bands. The molecular weight of the polypeptide is inversely proportional to its mobility. The molecular weight of the polypeptide subunit can be estimated directly from a semilog graph of the molecular weight of standard proteins versus their mobility or from a plot of the log of molecular weight versus mobility. Separation of proteins by SDS-PAGE is an excellent technique for producing individually "purified" proteins.

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      Blotting is a technique for the electrophoretic transfer of DNA, RNA or protein to a suitable membrane. The method most commonly used for the electrotransfer of proteins to nitrocellulose is that reported by Towbin et al. (1979). In order to take advantage of this technique for the purpose of amino acid analysis or N-terminal sequencing, the proteins must be transferred to a membrane that is stable to the chemicals used in these analytical procedures. For protein sequencing and amino acid analysis the proteins are transferred to a chemically stable membrane, polyvinylidene difluoride (PVDF). In our laboratory we use the "wet" transfer technique, rather than the "dry" transfer technique. Proteins are first separated by SDS-PAGE, the gel is removed from the electrophoresis cassette (do not stain the gel before blotting) and equilibrated in transfer buffer without methanol. The PVDF membrane is "activated" by dipping it in methanol it is then placed in transfer buffer containing methanol. The gel-PVDF sandwich is placed in a specially designed holder that in turn is placed in the buffer-containing electrophoresis unit. At the pH of the buffer (pH 8.3) most proteins are negatively charged and will migrate to the anode (positive electrode). In case one suspects the protein has a pI greater than 8.3, a PVDF membrane can be placed at the cathode-side of the gel as well. Alternatively, the pH of the transfer buffer can be adjusted to a higher pH. After transfer, the membrane is stained with Coomassie Blue R-250 and destained to locate the protein bands. Sections containing the proteins bands can then be excised for amino acid analysis and N-terminal protein sequencing.

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