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How do SDS-PAGE gels differ in a Bis-Tris system vs. a Tris-Glycine system?

How do SDS-PAGE gels differ in a Bis-Tris system vs. a Tris-Glycine system?


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Protein migrate differently in Bis-Tris and Tris-Glycine gels. I was curious about the actual reasons why. Do certain gel systems result in a tighter resolution than others?


SDS PAGE system rely on the fact that protein is denatured and surrounded by the SDS negatively charged detergent micelle. This eliminates most of the charge and idiosyncratic solubility differences from one protein to another and gives a reasonable separation based only on size of the protein which is related to the size of the SDS micelle around each molecule.

Bis-Tris and Tris-glycine buffers have quite different charge shielding characteristics. Bis (also known as 2-[Bis(2-hydroxyethyl)amino]-2-(hydroxymethyl)-1,3-propanediol) has a tertiary amine with a pKa of 6.46 and a pKb of 7.54. glycine is a zwitterion at any pH between 2.3 and 9.6. This creates a difference in the way that the buffer shields the SDS PAGE micelles from the rest of the electrical field, slowing down (probably glycine slows things down a bit) the time to resolution, but also giving the micelles more time to migrate.

So for SDS PAGE systems, the resolution of the gel has at least as much to do with the size of the gel pores (based on the acrylamide and bis-acrylamide percentages), the amount of protein you are putting into a given volume of loading well, and the size differences of the bands you are tying to separate. (you need to get pretty lucky to separate a 100.54 kda band from a 100.85 kDa band, but 1.5 to 1.8 kDa is easier 10 to 14 kDa is even easier). Also consider adjusting the current or voltage of the power supply. The buffer system is only one consideration in planning your experiment and often not the primary factor of quality.


Background

The pH of the separating gel in “standard” SDS-PAGE (a.k.a. Laemmli buffer system) is roughly 8-9 which is conducive to the deamination and alkylation of proteins, as well as reoxidation of reduced cysteines during electrophoresis. What this means is that your protein will form disulfide crosslinks during the stacking event because the protein migrates into the gel away from the reducing reagent in the sample buffer, and gets focused to a high concentration. Acrylamide gels cast in alkaline buffers are also unstable during long term storage, breaking down to acrylic acid after 1 to 2 months resulting in loss of pore size, poor resolution, and modified proteins.

In this protocol, in-gel cysteine reoxidation is suppressed by casting and running under slightly acidic (

pH 6.5) conditions favoring cysteine protonation. Additionally, a reducing agent, sodium bisulfite, is included in the running buffer and will migrate into the gel and maintain a reducing environment. Another feature of this gel system is that the lower MW proteins near the buffer front do not accelerate towards the end of the run to the same degree as in Laemmli buffers. The result is higher resolution and a band distribution not unlike a gradient gel.

The Stacking and Resolving layers of the gel use the same buffer. This allows gels to be cast and stored for a long time (diffusion doesn't ruin the stacking chemistry). Also, the same tank running buffer is used at both the cathode and anode.


Sample bis-Tris MES-Tris Gels The first gel is of purified proteins. The second gel is of total E. coli lysate with and without an induced protein. Note the broad separation range of the molecular weight markers.


The Chemical Ingredients and What they Do

What exactly does SDS do?
It unfolds proteins. Application of SDS to proteins causes them to lose their higher order structures and become linear. Since SDS is anionic (negatively charged), it binds to all the positive charges on a protein, effectively coating the protein in negative charge.

Why do we want the protein coated in negative charges?
To remove charge as a factor in protein migration through the gel. SDS binds to proteins with high affinity and in high concentrations. This results in all proteins (regardless of size) having a similar net negative charge and a similar charge-to-mass ratio. In this way, when they start moving through a gel, the speed that they move will be dependent on their size, and not their charge.

After getting hit with SDS, is a protein’s size the only thing that affects its migration through the gel?
It is by far the biggest factor. However, SDS can bind differently to different proteins. Hydrophobic proteins may bind more SDS, and proteins with post-translational modifications such as phosphorylation and glycosylation may bind less SDS. These effects are usually negligible, but not always, and should be considered if your protein is running at a different molecular weight than expected.

What is in the running buffer?
Tris, glycine, and SDS, pH 8.3. Tris is the buffer used for most SDS-PAGE. Its pKa of 8.1 makes it an excellent buffer in the 7-9 pH range. This makes it a good choice for most biological systems. SDS in the buffer helps keep the proteins linear. Glycine is an amino acid whose charge state plays a big role in the stacking gel. More on that in a bit.

What is in the sample loading buffer?
Tris-HCl, SDS, glycerol, beta mercaptoethanol (BME), Bromophenol Blue. This is the buffer you mix with your protein samples prior to loading the gel. Again with the Tris buffer and its pKa. The SDS denatures and linearizes the proteins, coating them in negative charge. BME breaks up disulfide bonds in the proteins to help them enter the gel. Glycerol adds density to the sample, helping it drop to the bottom of the loading wells and to keep it from diffusing out of the well while the rest of the gel is loaded. Bromophenol Blue is a dye that helps visualization of the samples in the wells and their movement through the gel. Sample loading buffer is also known as Laemmli Buffer, named after the Swiss professor who invented it around 1970.

What is in the gels?
Tris-HCl, acrylamide, water, SDS, ammonium persulfate, and TEMED. Although the pH values are different, both the stacking and resolving layers of the gel contain these components. Tris and SDS are there for the reasons described above. Ammonium persulfate and TEMED work together to catalyze the polymerization of the acrylamide. The Cl- ions from the Tris-HCl work with the glycine ions in the stacking gel. Again, more to come on that.


Knowing what gradient to choose depends on the size of the proteins you are trying to visualize on your gel. Keeping Table 1 in mind, here are some scenarios and the matching gradient gels.

Range of protein sizesLow / High acrylamide percentagesApplication
4 – 250 kDa4% / 20%Discovery work you are looking for everything under the sun
10 – 100 kDa8% / 15%A more targeted approach, but you want to avoid multiple gels
50 – 75 kDa10% / 12.5%You are trying to resolve similarly sized proteins

Protein size determination? - (Feb/26/2008 )

It depends On how monomers are attached if it is a dimer at all. if they are attached by cystine bonds you still can perform SDS-PAGE. Otherwise it depends on what techniques are available to you.
No in PAGE proteins do not migrate according to their mass, completely false! they migrate according to their net charge and tertiary structure shape.

I do not agree that PAGE give any useful information about size since in pI-appropriate native PAGE protein runs in retention to its size

What you mean? I didn't say PAGE, you said PAGE

it is reliable, however, to find the right running conditions, particularly the right buffer system, is tedious work

I do not know the difference between blue and regulare native gel go ask mdfenko

I said SDS-PAGE!
It is a standard method for determining size, so standard that you can find it in almost every biochemistry text book.
As I said PAGE won't give you any information about size. If you set pH to pI how do you suppose the protein is going to move in electric field? and decide do you want to set pH to pI of unknown protein or your marker?
take your words back or show me a reference.

you wrote on 1st May: "If you run a PAGE it won't give you any useful information about size of your protein."

So, you did discus PAGE. If you think in terms of SDS-PAGE, it should be specified.

Running native PAGE with a pH of pI of the protein of interest actually does not make sense but I did not recommend this.

I do not agree that PAGE give any useful information about size since in pI-appropriate native PAGE protein runs in retention to its size

What you mean? I didn't say PAGE, you said PAGE

it is reliable, however, to find the right running conditions, particularly the right buffer system, is tedious work

I do not know the difference between blue and regulare native gel go ask mdfenko

I said SDS-PAGE!
It is a standard method for determining size, so standard that you can find it in almost every biochemistry text book.
As I said PAGE won't give you any information about size. If you set pH to pI how do you suppose the protein is going to move in electric field? and decide do you want to set pH to pI of unknown protein or your marker?
take your words back or show me a reference.

you wrote on 1st May: "If you run a PAGE it won't give you any useful information about size of your protein."

So, you also did discus PAGE in general. If you thought in terms of SDS-PAGE, it should have been specified.

Running native PAGE with a pH of pI of the protein of interest actually does not make sense but I did not recommend this.

Unfortunately BN-PAGE is not that accurate in determining the size of a protein, this is because they migrate depending on how much coomassie they bind. Someone in your building must have an old FPLC laying around with a gel filtration column. Run some standards and that is probably the best method to determine molecular weight. HPLC is probably even more accurate but harder to come by.

I do not agree that PAGE give any useful information about size since in pI-appropriate native PAGE protein runs in retention to its size

What you mean? I didn't say PAGE, you said PAGE

it is reliable, however, to find the right running conditions, particularly the right buffer system, is tedious work

I do not know the difference between blue and regulare native gel go ask mdfenko

I said SDS-PAGE!
It is a standard method for determining size, so standard that you can find it in almost every biochemistry text book.
As I said PAGE won't give you any information about size. If you set pH to pI how do you suppose the protein is going to move in electric field? and decide do you want to set pH to pI of unknown protein or your marker?
take your words back or show me a reference.

you wrote on 1st May: "If you run a PAGE it won't give you any useful information about size of your protein."

So, you did discus PAGE. If you think in terms of SDS-PAGE, it should be specified.

Running native PAGE with a pH of pI of the protein of interest actually does not make sense but I did not recommend this.

amazing explanation, I don't need to say anything readers can read quoted text and judge!

From the homepages of Invitrogen I found information of Invitrogen BN-PAGE system:

The NativePAGE™ Novex® Bis-Tris Gel System is a pre-cast polyacrylamide mini gel system that provides a sensitive and high-resolution method for analysis of native membrane protein complexes, native soluble proteins, molecular mass estimations, and assessment of purity of native proteins. It is based on the blue native polyacrylamide gel electrophoresis technique (BN PAGE) developed by Schagger and von Jagow (1-3).

And description of markers:
The NativeMark™ Unstained Protein Standard is a ready-to-use protein marker to allow for accurate molecular weight estimation of proteins using native gel electrophoresis with Tris-Glycine or NuPAGE® Novex Tris-Acetate Gels.

So from this description I get the idea that it is reliable for estimation of molecular weight. It is based in Coomassie and gradient gel

From the homepages of Invitrogen I found information of Invitrogen BN-PAGE system:

The NativePAGE™ Novex® Bis-Tris Gel System is a pre-cast polyacrylamide mini gel system that provides a sensitive and high-resolution method for analysis of native membrane protein complexes, native soluble proteins, molecular mass estimations, and assessment of purity of native proteins. It is based on the blue native polyacrylamide gel electrophoresis technique (BN PAGE) developed by Schagger and von Jagow (1-3).

And description of markers:
The NativeMark™ Unstained Protein Standard is a ready-to-use protein marker to allow for accurate molecular weight estimation of proteins using native gel electrophoresis with Tris-Glycine or NuPAGE® Novex Tris-Acetate Gels.

So from this description I get the idea that it is reliable for estimation of molecular weight. It is based in Coomassie and gradient gel

sarita, buddy,
I went to their website and read it but I really can't accept a quote from a website as a scientific reason they didn't said why and how PAGE is useful in determining molecular mass they just said it and maybe if you ask more informed persons within that company they tell you we meant this and we meant that.
If you refer me to a reference like a renown text book(stryer, voet. ) or a decent article(nature, JBC. ) I will take back my words with honor but the truth is all famous references are crying that PAGE won't give you any useful information about molecular mass and to the contrary of the website they present it with reasons that I indicated in previous posts.

From the homepages of Invitrogen I found information of Invitrogen BN-PAGE system:

The NativePAGE™ Novex® Bis-Tris Gel System is a pre-cast polyacrylamide mini gel system that provides a sensitive and high-resolution method for analysis of native membrane protein complexes, native soluble proteins, molecular mass estimations, and assessment of purity of native proteins. It is based on the blue native polyacrylamide gel electrophoresis technique (BN PAGE) developed by Schagger and von Jagow (1-3).

And description of markers:
The NativeMark™ Unstained Protein Standard is a ready-to-use protein marker to allow for accurate molecular weight estimation of proteins using native gel electrophoresis with Tris-Glycine or NuPAGE® Novex Tris-Acetate Gels.

So from this description I get the idea that it is reliable for estimation of molecular weight. It is based in Coomassie and gradient gel

sarita, buddy,
I went to their website and read it but I really can't accept a quote from a website as a scientific reason they didn't said why and how PAGE is useful in determining molecular mass they just said it and maybe if you ask more informed persons within that company they tell you we meant this and we meant that.
If you refer me to a reference like a renown text book(stryer, voet. ) or a decent article(nature, JBC. ) I will take back my words with honor but the truth is all famous references are crying that PAGE won't give you any useful information about molecular mass and to the contrary of the website they present it with reasons that I indicated in previous posts.

the references you request are given on the web page from invitrogen:

make sure you click on "read all"

From the homepages of Invitrogen I found information of Invitrogen BN-PAGE system:

The NativePAGE™ Novex® Bis-Tris Gel System is a pre-cast polyacrylamide mini gel system that provides a sensitive and high-resolution method for analysis of native membrane protein complexes, native soluble proteins, molecular mass estimations, and assessment of purity of native proteins. It is based on the blue native polyacrylamide gel electrophoresis technique (BN PAGE) developed by Schagger and von Jagow (1-3).

And description of markers:
The NativeMark™ Unstained Protein Standard is a ready-to-use protein marker to allow for accurate molecular weight estimation of proteins using native gel electrophoresis with Tris-Glycine or NuPAGE® Novex Tris-Acetate Gels.

So from this description I get the idea that it is reliable for estimation of molecular weight. It is based in Coomassie and gradient gel

sarita, buddy,
I went to their website and read it but I really can't accept a quote from a website as a scientific reason they didn't said why and how PAGE is useful in determining molecular mass they just said it and maybe if you ask more informed persons within that company they tell you we meant this and we meant that.
If you refer me to a reference like a renown text book(stryer, voet. ) or a decent article(nature, JBC. ) I will take back my words with honor but the truth is all famous references are crying that PAGE won't give you any useful information about molecular mass and to the contrary of the website they present it with reasons that I indicated in previous posts.

the references you request are given on the web page from invitrogen:

make sure you click on "read all"

hello, did you bother to read even abstract of those articles? They are talking about 2d electrophoresis and combining BN-PAGE with SDS-PAGE!

Abstract
Blue native Electrophoresis is a "charge shift" method developed for isolation of native membrane protein complexes from biological membranes that also separates both acidic and basic water-soluble proteins at a fixed pH of 7.5. In combination with a second dimension sodium dodecylsulfate electrophoresis it provides an analytical method for the determination of molecular mass and oligomeric state of nondissociated complexes, of subunit composition, and of degree of purity and for the detection of subcomplexes. The method was applied to analysis of cytochrome bc/bf complexes. By combination of a novel colorless native polyacrylamide gel electrophoresis (CN-PAGE) with blue native BN-PAGE, a two-dimensional native technique was developed that is suitable for preparation of highly pure membrane protein complexes.

As I had predicted if you ask more informed persons of the company they will say we meant this and we meant that, and we meant in combination with SDS-PAGE!
PAGE by itself won't give you any information about molecular mass! carve it in your brain! case closed!

From the homepages of Invitrogen I found information of Invitrogen BN-PAGE system:

The NativePAGE™ Novex® Bis-Tris Gel System is a pre-cast polyacrylamide mini gel system that provides a sensitive and high-resolution method for analysis of native membrane protein complexes, native soluble proteins, molecular mass estimations, and assessment of purity of native proteins. It is based on the blue native polyacrylamide gel electrophoresis technique (BN PAGE) developed by Schagger and von Jagow (1-3).

And description of markers:
The NativeMark™ Unstained Protein Standard is a ready-to-use protein marker to allow for accurate molecular weight estimation of proteins using native gel electrophoresis with Tris-Glycine or NuPAGE® Novex Tris-Acetate Gels.

So from this description I get the idea that it is reliable for estimation of molecular weight. It is based in Coomassie and gradient gel

sarita, buddy,
I went to their website and read it but I really can't accept a quote from a website as a scientific reason they didn't said why and how PAGE is useful in determining molecular mass they just said it and maybe if you ask more informed persons within that company they tell you we meant this and we meant that.
If you refer me to a reference like a renown text book(stryer, voet. ) or a decent article(nature, JBC. ) I will take back my words with honor but the truth is all famous references are crying that PAGE won't give you any useful information about molecular mass and to the contrary of the website they present it with reasons that I indicated in previous posts.

the references you request are given on the web page from invitrogen:

make sure you click on "read all"

hello, did you bother to read even abstract of those articles? They are talking about 2d electrophoresis and combining BN-PAGE with SDS-PAGE!

Abstract
Blue native Electrophoresis is a "charge shift" method developed for isolation of native membrane protein complexes from biological membranes that also separates both acidic and basic water-soluble proteins at a fixed pH of 7.5. In combination with a second dimension sodium dodecylsulfate electrophoresis it provides an analytical method for the determination of molecular mass and oligomeric state of nondissociated complexes, of subunit composition, and of degree of purity and for the detection of subcomplexes. The method was applied to analysis of cytochrome bc/bf complexes. By combination of a novel colorless native polyacrylamide gel electrophoresis (CN-PAGE) with blue native BN-PAGE, a two-dimensional native technique was developed that is suitable for preparation of highly pure membrane protein complexes.

As I had predicted if you ask more informed persons of the company they will say we meant this and we meant that, and we meant in combination with SDS-PAGE!
PAGE by itself won't give you any information about molecular mass! carve it in your brain! case closed!

now, if you read the paper (1991) (and more specifically, the discussion) you would have seen that the author was claiming to make some estimate of size. the second dimension was to see the subunits of the complexes being studied.

even native gels can give some size estimation (which can be enhanced with a gradient). the gel matrix and porosity will allow smaller proteins to pass through faster than large proteins with the same or similar charge.


Contents

SDS-PAGE is an electrophoresis method that allows protein separation by mass. The medium (also referred to as ′matrix′) is a polyacrylamide-based discontinuous gel. The polyacrylamide-gel is typically sandwiched between two glass plates in a slab gel. Although tube gels (in glass cylinders) were used historically, they were rapidly made obsolete with the invention of the more convenient slab gels. [3] In addition, SDS (sodium dodecyl sulfate) is used. About 1.4 grams of SDS bind to a gram of protein, [4] [5] [6] corresponding to one SDS molecule per two amino acids. SDS acts as a surfactant, masking the proteins' intrinsic charge and conferring them very similar charge-to-mass ratios. The intrinsic charges of the proteins are negligible in comparison to the SDS loading, and the positive charges are also greatly reduced in the basic pH range of a separating gel. Upon application of a constant electric field, the protein migrate towards the anode, each with a different speed, depending on its mass. This simple procedure allows precise protein separation by mass.

SDS tends to form spherical micelles in aqueous solutions above a certain concentration called the critical micellar concentration (CMC). Above the critical micellar concentration of 7 to 10 millimolar in solutions, the SDS simultaneously occurs as single molecules (monomer) and as micelles, below the CMC SDS occurs only as monomers in aqueous solutions. At the critical micellar concentration, a micelle consists of about 62 SDS molecules. [7] However, only SDS monomers bind to proteins via hydrophobic interactions, whereas the SDS micelles are anionic on the outside and do not adsorb any protein. [4] SDS is amphipathic in nature, which allows it to unfold both polar and nonpolar sections of protein structure. [8] In SDS concentrations above 0.1 millimolar, the unfolding of proteins begins, [4] and above 1 mM, most proteins are denatured. [4] Due to the strong denaturing effect of SDS and the subsequent dissociation of protein complexes, quaternary structures can generally not be determined with SDS. Exceptions are proteins that are stabilised by covalent cross-linking e.g. -S-S- linkages and the SDS-resistant protein complexes, which are stable even in the presence of SDS (the latter, however, only at room temperature). To denature the SDS-resistant complexes a high activation energy is required, which is achieved by heating. SDS resistance is based on a metastability of the protein fold. Although the native, fully folded, SDS-resistant protein does not have sufficient stability in the presence of SDS, the chemical equilibrium of denaturation at room temperature occurs slowly. Stable protein complexes are characterised not only by SDS resistance but also by stability against proteases and an increased biological half-life. [9]

Alternatively, polyacrylamide gel electrophoresis can also be performed with the cationic surfactants CTAB in a CTAB-PAGE, [10] [11] [12] or 16-BAC in a BAC-PAGE. [13]

The SDS-PAGE method is composed of gel preparation, sample preparation, electrophoresis, protein staining or western blotting and analysis of the generated banding pattern.

Gel production Edit

When using different buffers in the gel (discontinuous gel electrophoresis), the gels are made up to one day prior to electrophoresis, so that the diffusion does not lead to a mixing of the buffers. The gel is produced by radical polymerisation in a mold consisting of two sealed glass plates with spacers between the glass plates. In a typical mini-gel setting, the spacers have a thickness of 0.75 mm or 1.5 mm, which determines the loading capacity of the gel. For pouring the gel solution, the plates are usually clamped in a stand which temporarily seals the otherwise open underside of the glass plates with the two spacers. For the gel solution, acrylamide is mixed as gel-former (usually 4% V/V in the stacking gel and 10-12 % in the separating gel), methylenebisacrylamide as a cross-linker, stacking or separating gel buffer, water and SDS. By adding the catalyst TEMED and the radical initiator ammonium persulfate (APS) the polymerisation is started. The solution is then poured between the glass plates without creating bubbles. Depending on the amount of catalyst and radical starter and depending on the temperature, the polymerisation lasts between a quarter of an hour and several hours. The lower gel (separating gel) is poured first and covered with a few drops of a barely water-soluble alcohol (usually buffer-saturated butanol or isopropanol), which eliminates bubbles from the meniscus and protects the gel solution of the radical scavenger oxygen. After the polymerisation of the separating gel, the alcohol is discarded and the residual alcohol is removed with filter paper. After addition of APS and TEMED to the stacking gel solution, it is poured on top of the solid separation gel. Afterwards, a suitable sample comb is inserted between the glass plates without creating bubbles. The sample comb is carefully pulled out after polymerisation, leaving pockets for the sample application. For later use of proteins for protein sequencing, the gels are often prepared the day before electrophoresis to reduce reactions of unpolymerised acrylamide with cysteines in proteins.

By using a gradient mixer, gradient gels with a gradient of acrylamide (usually from 4 to 12%) can be cast, which have a larger separation range of the molecular masses. [14] Commercial gel systems (so-called pre-cast gels) usually use the buffer substance Bis-tris methane with a pH value between 6.4 and 7.2 both in the stacking gel and in the separating gel. [15] [16] These gels are delivered cast and ready-to-use. Since they use only one buffer (continuous gel electrophoresis) and have a nearly neutral pH, they can be stored for several weeks. The more neutral pH slows the hydrolysis and thus the decomposition of the polyacrylamide. Furthermore, there are fewer acrylamide-modified cysteines in the proteins. [15] Due to the constant pH in collecting and separating gel there is no stacking effect. Proteins in BisTris gels can not be stained with ruthenium complexes. [17] This gel system has a comparatively large separation range, which can be varied by using MES or MOPS in the running buffer. [15]

Sample preparation Edit

During sample preparation, the sample buffer, and thus SDS, is added in excess to the proteins, and the sample is then heated to 95 °C for five minutes, or alternatively 70 °C for ten minutes. Heating disrupts the secondary and tertiary structures of the protein by disrupting hydrogen bonds and stretching the molecules. Optionally, disulfide bridges can be cleaved by reduction. For this purpose, reducing thiols such as β-mercaptoethanol (β-ME, 5% by volume), dithiothreitol (DTT, 10 millimolar) or dithioerythritol (DTE, 10 millimolar) are added to the sample buffer. After cooling to room temperature, each sample is pipetted into its own well in the gel, which was previously immersed in electrophoresis buffer in the electrophoresis apparatus.

In addition to the samples, a molecular-weight size marker is usually loaded onto the gel. This consists of proteins of known sizes and thereby allows the estimation (with an error of ± 10%) of the sizes of the proteins in the actual samples, which migrate in parallel in different tracks of the gel. [18] The size marker is often pipetted into the first or last pocket of a gel.

Electrophoresis Edit

For separation, the denatured samples are loaded onto a gel of polyacrylamide, which is placed in an electrophoresis buffer with suitable electrolytes. Thereafter, a voltage (usually around 100 V, 10-20 V per cm gel length) is applied, which causes a migration of negatively charged molecules through the gel in the direction of the positively charged anode. The gel acts like a sieve. Small proteins migrate relatively easily through the mesh of the gel, while larger proteins are more likely to be retained and thereby migrate more slowly through the gel, thereby allowing proteins to be separated by molecular size. The electrophoresis lasts between half an hour to several hours depending on the voltage and length of gel used.

The fastest-migrating proteins (with a molecular weight of less than 5 kDa) form the buffer front together with the anionic components of the electrophoresis buffer, which also migrate through the gel. The area of the buffer front is made visible by adding the comparatively small, anionic dye bromophenol blue to the sample buffer. Due to the relatively small molecule size of bromophenol blue, it migrates faster than proteins. By optical control of the migrating colored band, the electrophoresis can be stopped before the dye and also the samples have completely migrated through the gel and leave it.

The most commonly used method is the discontinuous SDS-PAGE. In this method, the proteins migrate first into a collecting gel with neutral pH, in which they are concentrated and then they migrate into a separating gel with basic pH, in which the actual separation takes place. Stacking and separating gels differ by different pore size (4-6 % T and 10-20 % T), ionic strength and pH values (pH 6.8 or pH 8.8). The electrolyte most frequently used is an SDS-containing Tris-glycine-chloride buffer system. At neutral pH, glycine predominantly forms the zwitterionic form, at high pH the glycines lose positive charges and become predominantly anionic. In the collection gel, the smaller, negatively charged chloride ions migrate in front of the proteins (as leading ions) and the slightly larger, negatively and partially positively charged glycinate ions migrate behind the proteins (as initial trailing ions), whereas in the comparatively basic separating gel both ions migrate in front of the proteins. The pH gradient between the stacking and separation gel buffers leads to a stacking effect at the border of the stacking gel to the separation gel, since the glycinate partially loses its slowing positive charges as the pH increases and then, as the former trailing ion, overtakes the proteins and becomes a leading ion, which causes the bands of the different proteins (visible after a staining) to become narrower and sharper - the stacking effect. For the separation of smaller proteins and peptides, the TRIS-Tricine buffer system of Schägger and von Jagow is used due to the higher spread of the proteins in the range of 0.5 to 50 kDa. [19]

Gel staining Edit

At the end of the electrophoretic separation, all proteins are sorted by size and can then be analyzed by other methods, e. g. protein staining such as Coomassie staining (most common and easy to use), [20] [21] silver staining (highest sensitivity), [22] [23] [24] [25] [26] [27] stains all staining, Amido black 10B staining, [21] Fast green FCF staining, [21] fluorescent stains such as epicocconone stain [28] and SYPRO orange stain, [29] and immunological detection such as the Western Blot. [30] [31] The fluorescent dyes have a comparatively higher linearity between protein quantity and color intensity of about three orders of magnitude above the detection limit, i. e. the amount of protein can be estimated by color intensity. When using the fluorescent protein dye trichloroethanol, a subsequent protein staining is omitted if it was added to the gel solution and the gel was irradiated with UV light after electrophoresis. [32] [33]

In Coomassie Staining, Gel is Fixed in a 50% ethanol 10% glacial acetic acid solution for 1 hr. Then the solution is changed for fresh one and after 1 to 12 hrs Gel is changed to a Staining solution (50% methanol, 10% Glacial acetic Acid, 0.1% Coomassie Brilliant Blue) followed by destaining changing several times a destaining solution of 40% methanol, 10% glacial acetic acid.

Analysis Edit

Protein staining in the gel creates a documentable banding pattern of the various proteins. Glycoproteins have differential levels of glycosylations and adsorb SDS more unevenly at the glycosylations, resulting in broader and blurred bands. [34] Membrane proteins, because of their transmembrane domain, are often composed of the more hydrophobic amino acids, have lower solubility in aqueous solutions, tend to bind lipids, and tend to precipitate in aqueous solutions due to hydrophobic effects when sufficient amounts of detergent are not present. This precipitation manifests itself for membrane proteins in a SDS-PAGE in "tailing" above the band of the transmembrane protein. In this case, more SDS can be used (by using more or more concentrated sample buffer) and the amount of protein in the sample application can be reduced. An overloading of the gel with a soluble protein creates a semicircular band of this protein (e. g. in the marker lane of the image at 66 kDa), allowing other proteins with similar molecular weights to be covered. A low contrast (as in the marker lane of the image) between bands within a lane indicates either the presence of many proteins (low purity) or, if using purified proteins and a low contrast occurs only below one band, it indicates a proteolytic degradation of the protein, which first causes degradation bands, and after further degradation produces a homogeneous color ("smear") below a band. [35] The documentation of the banding pattern is usually done by photographing or scanning. For a subsequent recovery of the molecules in individual bands, a gel extraction can be performed.

Archiving Edit

After protein staining and documentation of the banding pattern, the polyacrylamide gel can be dried for archival storage. Proteins can be extracted from it at a later date. The gel is either placed in a drying frame (with or without the use of heat) or in a vacuum dryer. The drying frame consists of two parts, one of which serves as a base for a wet cellophane film to which the gel and a one percent glycerol solution are added. Then a second wet cellophane film is applied bubble-free, the second frame part is put on top and the frame is sealed with clips. The removal of the air bubbles avoids a fragmentation of the gel during drying. The water evaporates through the cellophane film. In contrast to the drying frame, a vacuum dryer generates a vacuum and heats the gel to about 50 °C.

For a more accurate determination of the molecular weight, the relative migration distances of the individual protein bands are measured in the separating gel. [36] [37] The measurements are usually performed in triplicate for increased accuracy. The relative mobility (called Rf value or Rm value) is the quotient of the distance of the band of the protein and the distance of the buffer front. The distances of the bands and the buffer front are each measured from the beginning of the separation gel. The distance of the buffer front roughly corresponds to the distance of the bromophenol blue contained in the sample buffer. The relative distances of the proteins of the size marker are plotted semi-logarithmically against their known molecular weights. By comparison with the linear part of the generated graph or by a regression analysis, the molecular weight of an unknown protein can be determined by its relative mobility. Bands of proteins with glycosylations can be blurred. [34] Proteins with many basic amino acids (e. g. histones) [38] can lead to an overestimation of the molecular weight or even not migrate into the gel at all, because they move slower in the electrophoresis due to the positive charges or even to the opposite direction. Accordingly, many acidic amino acids can lead to accelerated migration of a protein and an underestimation of its molecular mass. [39]

The SDS-PAGE in combination with a protein stain is widely used in biochemistry for the quick and exact separation and subsequent analysis of proteins. It has comparatively low instrument and reagent costs and is an easy-to-use method. Because of its low scalability, it is mostly used for analytical purposes and less for preparative purposes, especially when larger amounts of a protein are to be isolated.

Additionally, SDS-PAGE is used in combination with the western blot for the determination of the presence of a specific protein in a mixture of proteins - or for the analysis of post-translational modifications. Post-translational modifications of proteins can lead to a different relative mobility (i.e. a band shift) or to a change in the binding of a detection antibody used in the western blot (i.e. a band disappears or appears).

In mass spectrometry of proteins, SDS-PAGE is a widely used method for sample preparation prior to spectrometry, mostly using in-gel digestion. In regards to determining the molecular mass of a protein, the SDS-PAGE is a bit more exact than an analytical ultracentrifugation, but less exact than a mass spectrometry or - ignoring post-translational modifications - a calculation of the protein molecular mass from the DNA sequence.

In medical diagnostics, SDS-PAGE is used as part of the HIV test and to evaluate proteinuria. In the HIV test, HIV proteins are separated by SDS-PAGE and subsequently detected by Western Blot with HIV-specific antibodies of the patient, if they are present in his blood serum. SDS-PAGE for proteinuria evaluates the levels of various serum proteins in the urine, e.g. Albumin, Alpha-2-macroglobulin and IgG.

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.

While being one of the more precise and low-cost protein separation and analysis methods, the SDS-PAGE denatures proteins. Where non-denaturing conditions are necessary, proteins are separated by a native PAGE or different chromatographic methods with subsequent photometric quantification, for example affinity chromatography (or even tandem affinity purification), size exclusion chromatography, ion exchange chromatography. [40] Proteins can also be separated by size in a tangential flow filtration [41] or an ultrafiltration. [42] Single proteins can be isolated from a mixture by affinity chromatography or by a pull-down assay. Some historically early and cost effective but crude separation methods usually based upon a series of extractions and precipitations using kosmotropic molecules, for example the ammonium sulfate precipitation and the polyethyleneglycol precipitation.

In 1948, Arne Tiselius was awarded the Nobel Prize in Chemistry for the discovery of the principle of electrophoresis as the migration of charged and dissolved atoms or molecules in an electric field. [43] The use of a solid matrix (initially paper discs) in a zone electrophoresis improved the separation. The discontinuous electrophoresis of 1964 by L. Ornstein and B. J. Davis made it possible to improve the separation by the stacking effect. [44] The use of cross-linked polyacrylamide hydrogels, in contrast to the previously used paper discs or starch gels, provided a higher stability of the gel and no microbial decomposition. The denaturing effect of SDS in continuous polyacrylamide gels and the consequent improvement in resolution was first described in 1965 by David F. Summers in the working group of James E. Darnell to separate poliovirus proteins. [45] The current variant of the SDS-PAGE was described in 1970 by Ulrich K. Laemmli and initially used to characterise the proteins in the head of bacteriophage T4. [1] This Laemmli paper is widely cited for the invention of modern SDS-PAGE, but the technique was actually invented by Jake Maizel, who was doing a sabbatical in the MRC laboratory when Laemmli joined the lab as a postdoctoral fellow. Maizel shared his prior technology with Laemmli and together they made further improvements. Laemmli and Maizel had planned to follow up with a Methods paper but this never materialized. Maizel recounts the history of development of SDS-PAGE in brief commentary. [46]


Abstract

When monoclonal antibodies (mAbs) are analysed by non-reducing sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE), method-induced artifacts are a frequent phenomenon. Previous studies suggested that incomplete denaturation and disulfide-bond scrambling are two main causes of artifact bands. Thus, in practice samples are normally heated and treated with alkylating agent iodoacetamide (IAM) before loading to promote denaturation and block free sulfhydryl groups, respectively. In this work, we further studied the major cause of artifact bands on non-reducing SDS-PAGE and ways of eliminating artifacts with two purified mAbs. In both cases, it was found that artifact bands on non-gradient Tris-glycine gels are mainly caused by incomplete denaturation under typical gel conditions. In general, heating minimizes artifact bands due to incomplete denaturation but it also generates some extra bands. Combining heating with IAM treatment achieved slightly better results than heating alone. As an alternative to heating, treating the samples with 8 M urea also allows close to complete denaturation of samples and thus minimizes artifact bands. In addition, we learned that untreated samples (samples that are not heated or treated with urea) may look different on Bis-Tris gel depending on gel composition (non-gradient vs. gradient) and the buffer used (MES vs. MOPS). In certain case, the apparent lack of artifact bands on gradient Bis-Tris gel may be in fact due to insufficient resolution. In conclusion, this study further confirmed that full-denaturation of sample is critical for minimizing/avoiding artifact bands on non-reducing SDS-PAGE.


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.


Antibody-purity analysis is critical to successful development of monoclonal antibody (MAb) biopharmaceuticals. Their manufacture involves processes of protein purification, formulation, and stability evaluation. All those processes need highly accurate and reproducible analytical results to support decisions made by product developers and manufacturers.

A common technology for antibody-purity analysis is sodium-dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE). In this technique, a polypeptide chain binds SDS proportionally to its relative molecular mass. The detergent nature of SDS denatures proteins by disrupting their noncovalent bonds, thereby simplifying the molecular structure. Negatively charged SDS also acts to coat proteins consistently, allowing for electrically driven separation toward an anode (a positively charged electrode). With an SDS–protein constant-weight binding ratio of 1:1.4, intrinsic polypeptide charge becomes negligible. So the final separation of such proteins depends entirely on differences in the relative molecular mass of their denatured polypeptides.

PRODUCT FOCUS: MONOCLONAL ANTIBODIES

PROCESS FOCUS: MANUFACTURING

WHO SHOULD READ: ANALYTICAL, PRODUCT DEVELOPMENT

KEYWORDS: ELECTROPHORESIS, IGGS, DETERGENTS, GLYCOSYLATION

Because of its automated, quantitative nature, capillary electrophoresis (CE) technology is commonly used for antibody-purity analysis as well. In this technique, an antibody sample is mixed with a replaceable SDS-gel buffer and then electrophoresed through an SDS-gel filled capillary. To accomplish that, samples are injected into the capillary inlets capillary using high voltage. Protein migration through the separation matrix occurs in an anodic direction, and quantitative detection occurs near the distal end of the capillary using a UV absorbance detection system.

It is well known that CE provides high-resolution, quantitative analysis relative to SDS-PAGE. However, direct comparison of the two technologies is difficult unless standardized samples are used. Here we describe our direct comparison of CE and SDS-PAGE based on evaluating the same sample in both a normal and a heat-stressed state by both methods.

Experimental

SDS-PAGE: We used an Invitrogen NuPAGE Mini-Gel electrophoresis system (Life Technologies) with 4–12% Bis-Tris gel and GelCode Blue stain to analyze a human IgG antibody sample and the same sample again after heat stress for 14 days at 45 °C. Samples were diluted to 0.2 mg/mL with water and further diluted to 0.15 mg/mL with 4× Invitrogen NuPAGE LDS sample buffer (Life Technologies). We conducted gel preparation, sample loading, and analysis all according to the gel-manufacturer&aposs suggested procedure (included in the box). Using Alpha View integration software (ProteinSimple), we imaged the resulting gel to quantify percent area for each band and assess molecular weight.

CE-SDS: We used a PA 800 plus capillary electrophoresis system (Beckman Coulter) to analyze the same samples as above, performing sample preparation and system operation as suggested by the system manuals. Antibody samples were diluted to 1.0 mg/mL with SDS sample buffer, and nonreduced samples were heated at 70 °C for three minutes before injection into a bare, fused-silica capillary at 5 kV for 20 seconds. The machine separated proteins in an electric field of 500 V/cm for 35 minutes. No gel staining or destaining was necessary, and UV detection at 220 nm recorded the amount of protein passing through the capillary window. Beckman Coulter 32 Karat software determined sample quantitations and migration times.

Results and Discussion

Figure 1 compares the SDS-PAGE and CE-SDS analyses. In the gel image, lanes 1 and 12 are molecular-weight markers, lanes 2–6 are normal IgG samples, and lanes 7–11 are heat-stressed IgG samples. Normal IgG samples show a single major band at 150 kDa and a minor band at 130 kDa. Heat-stressed IgG samples show a major band at 150 kDa and four minor bands at 300, 130, 90, and 25 kDa. By comparison, CE-SDS results for both IgG and degraded IgG easily show high-resolution separation allowing for easy quantitation of degradation species attributable to a high signal-to-noise ratio.

Using an IgG standard, the PA 800 plus 32 Karat software can make peak assignments for CE-SDS electropherograms (Figure 2). As shown in the bottom electropherogram, the major impurities of normal IgG are a single light chain (LC) and a combination of two heavy chains and one light chain (2H 1L). As shown in the top electropherogram, the major impurities of degraded IgG are LC, two heavy chains (2H), 2H 1L, and nonglycosylated IgG.

Figure 3 shows band assignments based on molecular-weight markers for the SDS-PAGE separation compared with CE results. These data illustrate that signal-to-noise ratios for impurities seen from heat-stressed IgG are much lower in the scan from the gel than corresponding peaks in the CE-SDS electropherogram. It was difficult to perform autointegration for the SDS-PAGE impurity bands using the Alpha View software.

Figure 3 indicates that the CE-SDS also can detect nonglycosylated IgG easily, whereas SDS-PAGE cannot detect that species. This is a significant advantage of CE-SDS because glycosylated and nonglycosylated IgG often are functionally different, so these species must be separated from one another for accurate analysis. Other automated separations techniques — including chromatography methods — often are unable to effect such separations.

Assay reproducibility is a key attribute of analytical separations for quality control (QC) purposes, which are necessary for both interlab and intralab comparability documentation. Figure 4 shows reproducibility among four consecutive analyses of degraded IgG, which illustrates good overall reproducibility across the various fragments for the CE method.

A Superior Technology for IgGs

We found CE-SDS technology to be a much higher-resolving analytical separation option than SDS-PAGE for separation of a normal and heat-stressed IgG samples in purity determinations. We observed a significant difference in peak resolution and signal-to-noise ratio between the two methods. CE-SDS could detect nonglycosylated IgG, which was not resolved by SDS-PAGE. Because glycosylation is very important to IgG function, that separation capability of CE-SDS is significant enough to qualify the technique as a replacement for SDS-PAGE in this application, given the difficulty of quantitating nonglycosylated IgG by other methods.


Mass spectrometry of in-gel digests reveals differences in amino acid sequences of high-molecular-weight glutenin subunits in spelt and emmer compared to common wheat

High-molecular-weight glutenin subunits (HMW-GS) play an important role for the baking quality of wheat. The ancient wheats emmer and spelt differ in their HMW-GS pattern compared to modern common wheat and this might be one reason for their comparatively poor baking quality. The aim of this study was to elucidate similarities and differences in the amino acid sequences of two 1Bx HMW-GS of common wheat, spelt and emmer. First, the sodium dodecyl polyacrylamide gel electrophoresis (SDS-PAGE) system was optimized to separate common wheat, spelt and emmer Bx6 and Bx7 from other HMW-GS (e.g., 1Ax and 1By) in high concentrations. The in-gel digests of the Bx6 and Bx7 bands were analyzed by untargeted LC-MS/MS experiments revealing different UniProtKB accessions in spelt and emmer compared to common wheat. The HMW-GS Bx6 and Bx7, respectively, of emmer and spelt showed differences in the amino acid sequences compared to those of common wheat. The identities of the peptide variations were confirmed by targeted LC-MS/MS. These peptides can be used to differentiate between Bx6 and Bx7 of spelt and emmer and Bx6 and Bx7 of common wheat. The findings should help to increase the reliability and curation status of wheat protein databases and to understand the effects of protein structure on the functional properties.

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