Kda gel electrophoresis

Kda gel electrophoresis DEFAULT

Electrophoresis for western blot

Protein sizeGel acrylamide percentage
4–40 kDa20%
12–45 kDa15%
10–70 kDa12.5%
15–100 kDa10%
25–200 kDa8%

Acrylamide is a potent cumulative neurotoxin: wear gloves at all times.

Place gels in the electrophoresis tank as instructed by the manufacturer and bathe in migration buffer.


Positive controls

A positive control lysate can be used to demonstrate that the protocol is efficient and correct and that the antibody recognizes the target protein which may not be present in the experimental samples.

We strongly recommend the use of a positive control lysate when setting up a new experiment; this will give you immediate confidence in the protocol.



Molecular weight markers

A range of molecular weight markers will enable the determination of the protein size (see below) and also allow you to monitor the progress of an electrophoretic run. A range of molecular weight markers are commercially available.

We have the following molecular weight markers:

Molecular Weight Marker (ab48854)
​Prism Protein Ladder (10-175 kDa) (ab115832)
Prism Ultra Protein Ladder (10-180 kDa) (ab116027)
​Prism Ultra Protein Ladder (10-245 kDa) (ab116028)
Prism Ultra Protein Ladder (3.5-245 kDa) (ab116029)



Loading samples and running the gel

Use special gel loading tips or a micro-syringe to load the complete sample into wells. Take care not to touch the bottom of the wells with the tip as this will create a distorted band.

Never overfill wells. This could lead to poor data and poorly resolved bands if samples spill into adjacent wells.

Load 20–40 µg total protein per mini-gel well.

The gels should be submerged in migration buffer normally containing SDS, except in native gel electrophoresis.

A standard migration buffer (also called running buffer) for PAGE is 1x Tris-glycine:

​25 mM Tris base
190 mM glycine
0.1% SDS
Check the pH; it should be around 8.3.

Run the gel for the recommended time as instructed by the manufacturer; this can vary from machine to machine (1 h to overnight depending on the voltage).

When the dye (the migration front) reaches the bottom of the gel, turn the power off. Proteins will slowly elute from the gel at this point, so do not store the gel; proceed immediately to transfer.



Loading controls​

Loading controls are required to ensure that the lanes in your gel have been evenly loaded with sample, especially when a comparison must be made between the expression levels of a protein in different samples. They are also useful to check for even transfer from the gel to the membrane across the whole gel.

Where even loading or transfer have not occurred, the loading control bands can be used to quantify the protein amounts in each lane. For publication-quality work, use of a loading control is absolutely essential.

Visit our loading control guide.

The following table contains information about common loading controls:

Loading controlSample typeMolecular weightCaution
VinculinWhole cell125 kDa
CyclophilinWhole cell24 kDa
GAPDHWhole cell35 kDaSome physiological factors, such as hypoxia and diabetes, increase GAPDH expression in certain cell types.
Cofilin Whole cell
Nuclear
Membrane
Cytoskeleton
19 kDa
Alpha tubulinWhole cell
Cytoskeleton
50 kDaTubulin expression may vary according to resistance to antimicrobial and antimiotic drugs (Sangrajang S et al., 1998; Prasad V et al., 2000).
Beta tubulinWhole cell
Cytoskeleton
50 kDaTubulin expression may vary according to resistance to antimicrobial and antimiotic drugs (Sangrajang S et al., 1998; Prasad V et al., 2000).
ActinWhole cell
Cytoskeleton
42 kDa
Beta actinWhole cell
Cytoskeleton
40 kDaNot suitable for skeletal muscle samples. Changes in cell-growth conditions and interactions with extracellular matrix components may after actin protein synthesis (Farmer et al., 1983).
VDAC1/PorinMitochondrial30 kDa
COX IVMitochondrial20 kDaMany proteins run at the same 16 kDa size as COX IV.
HSP60Mitochondrial
Membrane
60 kDa
Lamin B1Nuclear66 kDaNot suitable for samples where the nuclear envelope is removed.
HDAC1Nuclear55 kDa
YY1Nuclear45 kDa
TBPNuclear35 kDaNot suitable for samples where DNA is removed.
PCNANuclear30 kDa
Cdk4Nuclear
Membrane
34 kDa
Na-K ATPaseMembrane110 kDa
TransferrinSerum75 kDa
Sours: https://www.abcam.com/protocols/electrophoresis-for-western-blot

Explore: Protein electrophoresis products  Download: Protein Electrophoresis Handbook

What is protein electrophoresis?

Protein electrophoresis is a standard laboratory technique by which charged protein molecules are transported through a solvent by an electrical field. Both proteins and nucleic acids may be separated by electrophoresis, which is a simple, rapid, and sensitive analytical tool. Most biological molecules carry a net charge at any pH other than their isoelectric point and will migrate at a rate proportional to their charge density. The mobility of a molecule through an electric field will depend on the following factors: field strength, net charge on the molecule, size and shape of the molecule, ionic strength, and properties of the matrix through which the molecule migrates (e.g., viscosity, pore size). Polyacrylamide and agarose are two support matrices commonly used in electrophoresis. These matrices serve as porous media and behave like a molecular sieve. Agarose has a large pore size and is suitable for separating nucleic acids and large protein complexes. Polyacrylamide has a smaller pore size and is ideal for separating majority of proteins and smaller nucleic acids.

Several forms of polyacrylamide gel electrophoresis (PAGE) exist, and each form can provide different types of information about proteins of interest. Denaturing and reducing sodium dodecyl sulfate PAGE (SDS-PAGE) with a discontinuous buffer system is the most widely used electrophoresis technique and separates proteins primarily by mass. Nondenaturing PAGE, also called native-PAGE, separates proteins according to their mass/charge ratio. Two-dimensional (2D) PAGE separates proteins by native isoelectric point in the first dimension and by mass in the second dimension.

SDS-PAGE separates proteins primarily by mass because the ionic detergent SDS denatures and binds to proteins to make them uniformly negatively charged. Thus, when a current is applied, all SDS-bound proteins in a sample will migrate through the gel toward the positively charged electrode. Proteins with less mass travel more quickly through the gel than those with greater mass because of the sieving effect of the gel matrix. Once separated by electrophoresis, proteins can be detected in a gel with various stains, transferred onto a membrane for detection by western blotting and/or excised and extracted for analysis by mass spectrometry. Protein gel electrophoresis is, therefore, a fundamental step in many kinds of proteomics analysis.

Watch this video summary of protein gel electrophoresis by SDS-PAGE.

What are polyacrylamide gels?

Polyacrylamide is the material of choice for preparing electrophoretic gels to separate proteins by size. Polyacrylamide gels are prepared by mixing acrylamide with bisacrylamide to form a crosslinked polymer network when the polymerizing agent, ammonium persulfate (APS), is added. TEMED (N,N,N’,N'-tetramethylenediamine) catalyzes the polymerization reaction by promoting the production of free radicals by APS. At this stage it becomes polyacrylamide.

Polymerization and crosslinking of acrylamide. The ratio of bisacrylamide (N,N'-methylenediacrylamide) to acrylamide, as well as the total concentration of both components, affects the pore size and rigidity of the final gel matrix. These, in turn, affect the range of protein sizes (molecular weights) that can be resolved.

Example recipe for a traditional polyacrylamide gel: 10% Tris-glycine mini gel for SDS-PAGE:

  • 7.5 mL 40% acrylamide solution
  • 3.9 mL 1% bisacrylamide solution
  • 7.5 mL 1.5 M Tris-HCl, pH 8.7
  • Add water to 30 mL
  • 0.3 mL 10% APS
  • 0.3 mL 10% SDS
  • 0.03 mL TEMED

The size of the pores created in the gel is inversely related to the polyacrylamide percentage (concentration). For instance, a 7% polyacrylamide gel has larger pores than a 12% polyacrylamide gel. Low-percentage gels are used to resolve large proteins, and high-percentage gels are used to resolve small proteins. "Gradient gels" are specially prepared to have a low percentage of polyacrylamide at the top (beginning of sample path) and a high percentage at the bottom (end), enabling a broader range of protein sizes to be separated.

Electrophoresis gels are formulated in buffers that enable electrical current to flow through the matrix. The prepared solution is poured into the thin space between two glass or plastic plates that form a cassette. This process is referred to as casting a gel. Once the gel polymerizes, the cassette is mounted (usually vertically) into an apparatus so that the top and bottom edges are placed in contact with buffer chambers containing a cathode and an anode, respectively. The running buffer contains ions that conduct current through the gel. When proteins are loaded into wells at the top edge and current is applied, the proteins are drawn by the current through the matrix slab and separated by the sieving properties of the gel.

To obtain optimal resolution of proteins, a stacking gel is cast over the top of the resolving gel. The stacking gel has a lower concentration of acrylamide (e.g., 7% for larger pore size), lower pH (e.g., 6.8), and a different ionic content. This allows the proteins in a loaded sample to be concentrated into one tight band during the first few minutes of electrophoresis before entering the resolving portion of a gel. A stacking gel is not necessary when using a gradient gel, as the gradient itself performs this function.

Polyacrylamide gel electrophoresis in progress. Prepared gel cassettes are inserted into a gel tank, in this case the Invitrogen Mini Gel Tank, which holds two mini gels at a time. After wells are loaded with protein samples, the gels submerged in a conducting running buffer, and electrical current is applied, typically for 20 to 40 minutes. Run times vary according to the size and percentage of the gel and gel chemistry.

SDS-PAGE (denaturing) vs. native-PAGE

SDS-PAGE

In SDS-PAGE, the gel is cast in a buffer containing sodium dodecyl sulfate (SDS), an anionic detergent. SDS denatures proteins by wrapping around the polypeptide backbone. By heating the protein sample between 70-100°C in the presence of excess SDS and thiol reagent, disulfide bonds are cleaved, and the protein is fully dissociated into its subunits. Under these conditions most polypeptides bind SDS in a constant weight ratio (1.4 g of SDS:1 g of polypeptide). The intrinsic charges of the polypeptide are insignificant compared to the negative charges provided by the bound detergent so that the SDS-polypeptide complexes have essentially the same negative charge and shape. Consequently, proteins migrate through the gel strictly according to polypeptide size with very little effect from compositional differences. The simplicity and speed of this method, plus the fact that only microgram quantities of protein are required, have made SDS-PAGE the most widely used method for determination of molecular mass in a polypeptide sample. Proteins from almost any source are readily solubilized by SDS so the method is generally applicable.

When a set of proteins of known mass are run alongside samples in the same gel, they provide a reference by which the mass of sample proteins can be determined. These sets of reference proteins are called mass markers or molecular weight markers (MW markers), protein ladders, or size standards, and they are available commercially in several forms.

Native-PAGE

In native-PAGE, proteins are separated according to the net charge, size, and shape of their native structure. Electrophoretic migration occurs because most proteins carry a net negative charge in alkaline running buffers. The higher the negative charge density (more charges per molecule mass), the faster a protein will migrate. At the same time, the frictional force of the gel matrix creates a sieving effect, regulating the movement of proteins according to their size and three-dimensional shape. Small proteins face only a small frictional force, while larger proteins face a larger frictional force. Thus native-PAGE separates proteins based upon both their charge and mass.

Because no denaturants are used in native-PAGE, subunit interactions within a multimeric protein are generally retained and information can be gained about the quaternary structure. In addition, some proteins retain their enzymatic activity (function) following separation by native-PAGE. Thus, this technique may be used for preparation of purified, active proteins.

Following electrophoresis, proteins can be recovered from a native gel by passive diffusion or electro-elution. To maintain the integrity of proteins during electrophoresis, it is important to keep the apparatus cool and minimize denaturation and proteolysis. pH extremes should generally be avoided in native-PAGE, as they may lead to irreversible damage, such as denaturation or aggregation, to proteins of interest.

1D vs. 2D PAGE

1-dimensional polyacrylamide gel electrophoresis

The most common form of protein gel electrophoresis is comparative analysis of multiple samples by one-dimensional (1D) electrophoresis. Gel sizes range from 2 x 3 cm (tiny) to 15 x 18 cm (large format). The most popular size (approx. 8 x 8 cm) is usually referred to as a "mini gel". Medium-sized gels (8 x 13 cm) are called midi gels. Small gels require less time and reagents than their larger counterparts and are suited for rapid protein screening. However, larger gels provide better resolution and are needed for separating similar proteins or a large number of proteins.

Protein samples are added to sample wells at the top of the gel. When the electrical current is applied, the proteins move down through the gel matrix, creating what are called lanes of protein bands. Samples that are loaded in adjacent wells and electrophoresed together are easily compared to each other after staining or other detection strategies. The intensity of staining and thickness of protein bands are indicative of their relative abundance. The positions (height) of bands within their respective lanes indicate their relative sizes (and/or other factors affecting their rate of migration through the gel).

Protein lanes and bands in 1D SDS-PAGE. Depicted here is a protein ladder, purified proteins and E. coli lysate loaded on a 4–20% gradient Novex Tris-Glycine gel; Lanes 1, 5, 10: 5 µL Thermo Scientific PageRuler Unstained Protein Ladder); lanes 2, 6, 9: 5 µL Mark12 Unstained Standard; lane 3: 10 µg E. coli lysate (10 µL sample volume); lane 4: 6 µg BSA (10 µL sample volume); lane 7: 6 µg hIgG (10 µL sample volume); lane 8: 20 µg E. coli lysate (20 µL sample volume). Electrophoresis was performed using the Mini Gel Tank. Sharp, straight bands were observed after staining with SimplyBlue SafeStain. Images were acquired using a flatbed scanner.

2-dimensional polyacrylamide gel electrophoresis

Multiple components of a single sample can be resolved most completely by two-dimensional electrophoresis (2D-PAGE). The first dimension separates proteins according to their native isoelectric point (pI) using a form of electrophoresis called isoelectric focusing (IEF). The second dimension separates by mass using ordinary SDS-PAGE. 2D PAGE provides the highest resolution for protein analysis and is an important technique in proteomic research, where resolution of thousands of proteins on a single gel is sometimes necessary.

To perform IEF, a pH gradient is established in a tube or strip gel using a specially formulated buffer system or ampholyte mixture. Ready-made IEF strip gels (called immobilized pH gradient strips or IPG strips) and required instruments are available from certain manufacturers. During IEF, proteins migrate within the strip to become focused at the pH points at which their net charges are zero. These are their respective isoelectric points.

The IEF strip is then laid sideways across the top of an ordinary 1D gel, allowing the proteins to be separated in the second dimension according to size.

Example 2-D electrophoresis data. In the first dimension, one or more samples are resolved by isoelectric focusing (IEF) in strip gels. IEF is usually performed using precast immobilized pH-gradient (IPG) strips on a specialized horizontal electrophoresis platform. For the second dimension, a gel containing the pI-resolved sample is laid across to top of a slab gel so that the sample can then be further resolved by SDS-PAGE.

Comparison of different gel chemistry systems

Three basic types of buffers are required: the gel casting buffer, the sample buffer, and the running buffer that fills the electrode reservoirs. Electrophoresis may be performed using continuous or discontinuous buffer systems. A continuous buffer system, which utilizes only one buffer in the gel, sample, and gel chamber reservoirs, is most often used for nucleic acid analysis and rarely used for protein gel electrophoresis. Proteins separated using a continuous buffer system tend to be diffuse and poorly resolved. Conversely, discontinuous buffer systems utilize a different gel buffer and running buffer. These systems also use two gel layers of different pore sizes and different buffer compositions (the stacking and separating gels). Electrophoresis using a discontinuous buffer system results in concentration of the sample and higher resolution. The various commonly used discontinuous gel buffer systems as summarized below.

Tris-Glycine

The most widely used gel system for separating a broad range of proteins is the Laemmli system. The classical Laemmli system, consisting of Tris-glycine gels and Tris-glycine running buffer, can be used for both SDS-PAGE and native PAGE. This system is used widely because reagents for casting Tris-glycine gels are relatively inexpensive and readily available. Gels using this chemistry can be made in a variety gel formats and percentages.

The formulation of this discontinuous buffer system creates a stacking effect to produce sharp protein bands at the beginning of the electrophoretic run. A boundary is formed between chloride, the leading ion, and glycinate, the trailing ion. Tris buffer provides the common cations. As proteins migrate into the resolving gel, they are separated according to size. Tris-glycine gels are used in conjunction with Laemmli sample buffer, and Tris/glycine/SDS running buffer is used for denaturing SDS-PAGE. Native PAGE is performed using native sample and running buffers without denaturants or SDS. The pH and ionic strength of the buffer used for running the gel (Tris, pH 8.3) are different from those of the buffers used in the stacking gel (Tris, pH 6.8) and the resolving gel (Tris, pH 8.8). The highly alkaline operating pH of the Laemmli system may cause band distortion, loss of resolution, or artifact bands.

Disadvantages of using the Laemmli system:

  • Hydrolysis of polyacrylamide at the high pH of the resolving gel, resulting in a short shelf life of 8 weeks
  • Chemical alterations such as deamination and alkylation of proteins due to the high pH of the resolving gel
  • Reoxidation of reduced disulfides from cysteine-containing proteins
  • Cleavage of Asp-Pro bonds of proteins when heated at 100°C in Laemmli sample buffer, pH 5.2

Bis-Tris

In contrast to conventional Tris-glycine gels, Bis-Tris HCI–buffered gels run closer to neutral pH, thus offering enhanced stability and greatly extended shelf-life over Tris-glycine gels (up to 16 months at room temperature). The neutral pH provides reduced protein degradation and is good for applications where high sensitivity is required such as analysis of posttranslational modifications, mass spectrometry, or sequencing.

For Bis-Tris gels, chloride serves as the leading ion and MES or MOPS act as the trailing ion. Bis-Tris buffer forms the common cation. Markedly different protein migration patterns are produced depending on whether a Bis-Tris gel is run with MES or MOPS denaturing running buffer: MES buffer is used for smaller proteins, and MOPS buffer is used for mid-sized proteins.

Due to differences in ionic composition and pH, gel patterns obtained with Bis-Tris gels cannot be compared to those obtained with Tris-glycine gels. To prevent protein reoxidation, Bis-Tris gels must be run with alternative reducing agents such as sodium bisulfite. Reducing agents frequently used with Tris-glycine gels, such as beta-mercaptoethanol and dithiothreitol (DTT), do not undergo ionization at low pH levels and are not able to migrate with proteins in a Bis-Tris gel.

Tris-Acetate

Tris-acetate gel chemistry enables the optimal separation of high molecular weight proteins. Tris-acetate gels use a discontinuous buffer system involving three ions- acetate, tricine and tris. Acetate serves as a leading ion due to its high affinity to the anode relative to other anions in the system. Tricine serves as the trailing ion.Tris-acetate gels can be used with both SDS-PAGE and native PAGE running buffers. Compared with Tris-glycine gels, Tris-acetate gels have a lower pH, which enhances the stability of these gels and minimizes protein modifications, resulting in sharper bands.

Tris Tricine

The Tris-Tricine gel system is a modification of the Tris-glycine gel system and is optimized to resolve low molecular weight proteins in the range of 2–20 kDa. As a result of reformulating the Laemmli running buffer and using Tricine in place of glycine, SDS-polypeptides form behind the leading ion front rather than running with the SDS front, thus allowing for their separation into discrete bands.

Zymogram

Zymogram gels are Tris-glycine gels containing gelatin or casein and are used to characterize proteases that utilize them as substrates. Samples are run under denaturing conditions, but due to the absence of reducing agents, proteins undergo renaturation. Proteolytic proteins present in the sample consume the substrate, generating clear bands against a background stained blue.

Gel buffer system selection

The choice of whether to use one chemistry or another depends on the abundance of the protein separating, the size of the protein and the downstream application. For separation of a broad range of proteins two chemistries: Bis-Tris and Tris-glycine are well suited. Bis-Tris gel chemistry provides greater sensitivity for protein detection compared to Tris-glycine gel chemistry. Choose Bis-Tris gel chemistry when you have a low abundance of protein or when the downstream application requires high protein integrity, such as posttranslational modification analysis, mass spectrometry, or sequencing.

 Bis-TrisTris-glycineTris-acetateTricine
Protein sample typeBroad range MW (6-400 kDa)Broad range MW (6-400 kDa)High range MW (40-500 kDa)Low range MW (2.5-40 kDa)
Chemistry benefitsNeutral pH for high-sensitivity applications and reduced protein degradationTraditional Laemmli-styleAnalysis of high molecular weight proteins; neutral pHAnalysis of low molecular weight proteins
Recommended forWestern blotting, mass spectrometry, posttranslationally modified proteins, dilute samples, and low-abundance proteinsWestern blotting, in-gel staining, samples containing detergents and high salt, native- PAGE applicationsHigh molecular weight proteins, western blotting, mass spectrometry, posttranslationally modified proteins, native-PAGE applicationsLow molecular weight proteins, western blotting, in-gel staining

Sample buffers and running buffer formulations

Protein samples prepared for SDS-PAGE analysis are denatured by heating in the presence of a sample buffer containing 1% SDS with or without a reducing agent such as 20mM DTT, 2-mercaptoethanol (BME) or Tris(2-carboxyethyl)phosphine (TCEP). The protein sample is mixed with the sample buffer and heated for 3 to 5 minutes (according to the specific protocol) then cooled to room temperature before it is pipetted into the sample well of a gel. Loading buffers also contain glycerol so that they are heavier than water and sink neatly to the bottom of the buffer-submerged well when added to a gel.

If a suitable, negatively charged, low-molecular weight dye is also included in the sample buffer, it will migrate at the buffer-front, enabling one to monitor the progress of electrophoresis. The most common tracking dyes for sample loading buffers are bromophenol blue, phenol red and Coomassie blue. The table below summarizes common sample buffers and running buffers used in the different gel buffer systems.

Buffer formulations for discontinuous PAGE

Gel chemistrySample bufferRunning bufferSelection criteria
SDS-PAGE
Tris-glycineTris-glycine SDS sample buffer: Tris HCl (63 mM), glycerol (10%), SDS (2%), bromophenol blue (0.0025%), pH 6.8Tris-glycine SDS: Tris base (25 mM), glycine (192 mM), SDS (0.1%), pH 8.3Ease of preparation; relatively inexpensive, separation of broad range of molecular weight proteins
Bis-TrisLDS sample buffer: Tris base (141 mM), Tris HCl (106 mM), LDS (2%), EDTA (0.51 mM), SERVA Blue G-250 (0.22 mM), phenol red (0.175 mM), pH 8.5MES SDS: MES (50 mM), Tris base (50 mM), SDS (0.1%), EDTA (1 mM), pH 7.3
MOPS SDS: MOPS (50 mM), Tris base (50 mM), SDS (0.1%), EDTA (1 mM), pH 7.7
Relatively long shelf life; room temperature storage; neutral pH minimizes protein modifications, separation of broad range of molecular weight proteins
Tris-AcetateLDS sample buffer: Tris base (141 mM), Tris HCl (106 mM), LDS (2%), EDTA (0.51 mM), SERVA Blue G-250 (0.22 mM), phenol red (0.175 mM), pH 8.5Tris-acetate SDS: Tris base (50 mM), Tricine (50 mM), SDS (0.1%), pH 8.24Superior separation of protein complexes and high MW proteins; relatively long shelf life
Tris-TricineTricine SDS sample buffer: Tris HCl (450 mM), glycerol (12%), SDS (4%), Coomassie Blue G (0.00075%), phenol red (0.0025%), pH 8.45Tricine-SDS: Tris base (100 mM), tricine (100 mM), SDS (0.1%), pH 8.3Ideal for separating peptides and low molecular weight proteins
Native-PAGE
Tris-glycineNative sample buffer: Tris HCl (100 mM), glycerol (10%), bromophenol blue (0.00025%), pH 8.6Tris-Glycine Native buffer: Tris base (25 mM), glycine (192 mM), pH 8.3Retention of native protein structure
Tris-acetateNative sample buffer: Tris HCl (100 mM), glycerol (10%), bromophenol blue (0.00025%), pH 8.6Tris-Glycine Native buffer: Tris base (25 mM), glycine (192 mM), pH 8.3Superior separation of protein complexes and high MW proteins
IEF
IEFIEF Sample Buffer pH 3-7: Lysine (40 mM), glycerol (15%)
IEF Sample Buffer pH 3-10: Arginine (20 mM), Lysine (20 mM), glycerol (15%)
IEF cathode buffer pH 3-7: Lysine (40 mM)
IEF cathode buffer pH 3-10: Arginine (20 mM), lysine (20 mM)
IEF anode buffer: phosphoric acid 85% (7 mM)
Use to separate proteins according to isoelectric point (pI) rather than molecular weight
Protease detection
ZymogramTris-glycine SDS: Tris HCl (63 mM), glycerol (10%), SDS (2%), bromophenol blue (0.0025%), pH 6.8Tris-glycine SDS: Tris base (25 mM), glycine (192 mM), SDS (0.1%), pH 8.3Gelatin or casein gels provide substrates used to detect proteases

Gel electrophoresis running conditions

Gel TypeVoltageExpected currentRun time
Tris-glycineDenaturing: 125 volts constant
Native: 20-125 volts constant
Denaturing: 30-40 mA (start), 8-12 mA (end)
Native: 6-12 mA (start), 3-6 mA (end)
Denaturing: 90 min
Native: 1-12 hr
Bis-Tris200 volts constantNon-reducing: 100-125 mA (start), 60-70 mA (end) Reducing: 110-125 mA (start), 70-80 mA (end)35-50 min
Tris-AcetateDenaturing: 150 volts constant
Native: 20-150 volts constant
Denaturing and Native:
40-55 mA (start), 25-40 mA (end)
Denaturing: 60 min
Native: 1-12 hr
Tricine125 volts constant80 mA (start), 40 mA (end)90 min
IEF100 volts for 1hr, 200 volts for 1hr, 500 volts for 30 min5 mA (start), 6 mA (end)2.5 hr
Zymogram125 volts constant30-40 mA (start), 8-12 mA (end)90 min

Precast gels vs. handcast gels

Traditionally, researchers casted their own gels using standard recipes that are widely available in protein methods literature. More laboratories are moving to the convenience and consistency afforded by commercially available, ready-to-use precast gels. Precast gels are available in a variety of percentages, including difficult-to-pour gradient gels that provide excellent resolution and that separate proteins over the widest possible range of molecular weights. Precast gels are also available in the different buffer formulations (e.g., Tris-glycine, Bis-Tris, Tris-acetate, Tricine), which are designed to optimize shelf life, run time, and/or protein resolution.

For researchers who require unique gel formulations not available as precast gels, a wide range of reagents and equipment are available for pouring gels. However, technological innovations in buffers and gel polymerization methods enable manufacturers to produce gels with greater uniformity and longer shelf life than individual researchers can prepare on their own with traditional equipment and methods. In addition, precast polyacrylamide gels eliminate the need to work with the acrylamide monomer, which is a known neurotoxin and suspected carcinogen.

Precast vs. handcast protein gels for SDS-PAGE. Polyacrylamide gels can be purchased precast and ready- to- use (left) or prepared from reagents in the lab using a gel-casting system (right). Pictured here are the Novex Tris-Glycine Mini Gels, WedgeWell format (left) and the SureCast Gel Handcast System.

Protein gel electrophoresis chambers

To perform protein gel electrophoresis, the polyacrylamide gel and buffer must be placed in an electrophoresis chamber that is connected to a power source, and which is designed to conduct current through the buffer solution. When current is applied, the smaller molecules migrate more rapidly and the larger molecules migrate more slowly through the gel matrix. Multiple gel chamber designs exist. The choice of equipment is usually based on these factors: the dimensions of the gel cassette, with some tank designs accommodating more cassette sizes than others; the nature of the protein target, and corresponding gel resolution requirements; and whether a precast or handcast gel, and vertical or horizontal electrophoresis system, has been selected.

Mini gel tank for protein gel electrophoresis. This gel tank holds up to two mini gels and is compatible with the Invitrogen SureCast Gel Handcast System, and with all Invitrogen precast gels. The unique tank design enables side-by-side gel loading and enhanced viewing during use.

Protein ladders and standards

To assess the molecular masses (sizes) of proteins in a gel, a prepared mixture containing several proteins of known molecular masses is run alongside the test sample in one or more lanes of the gel. Such sets of known proteins are called protein molecular weight (or mass) markers or protein ladders. A standard curve can be constructed from the distances migrated by each marker protein. The distance migrated by the unknown protein is then plotted, and the molecular weight is extrapolated from the standard curve.

Several kinds of ready-to-use protein molecular weight (MW) markers are available that are either unlabeled or prestained for different modes of detection. These are pre-reduced and, therefore, primarily suited for SDS-PAGE rather than native PAGE. MW markers can also be made detectable via specialized labels, such as a fluorescent tag, and by other methods.

Accurate calibration of molecular weight standards in different buffer systems

Generally, protein mobility in SDS gels is a function of the length of the protein in its fully denatured state. By constructing a standard curve with protein standards of known molecular weights, the molecular weight of a sample protein can be calculated based upon its relative mobility. However, the same molecular weight standard may have slightly different mobility and therefore, different apparent molecular weight when run in different SDS-PAGE buffer systems.

The effects of secondary structure

When using SDS-PAGE for molecular weight calibration, slight deviations from the true molecular weight of a protein (definitively calculated from the known amino acid sequence) can occur mostly because of the retention of varying degrees of secondary structure in the protein, even in the presence of SDS. This phenomenon is more prevalent in proteins with highly organized secondary structures (such as collagens, histones, or highly hydrophobic membrane proteins) and in peptides, where the effect of local secondary structure becomes magnified relative to the total size of the peptide.

The pH factor

It has also been observed that slight differences in protein mobilities occur when the same proteins are run in different SDS-PAGE buffer systems. Each SDS-PAGE buffer system has a different pH, which affects the charge of a protein and its binding capacity for SDS. The degree of change in protein mobility is usually small in natural proteins but is more pronounced with atypical or chemically modified proteins, such as pre-stained standards. Apparent molecular weight values for pre-stained standards will vary between gel systems- it is important to use the apparent molecular weights that matches your gel for the most accurate calibration of your sample proteins.

Migration patterns of PageRuler Plus Prestained Protein Ladder in different electrophoretic conditions. The apparent molecular weight of each protein (kDa) varies between the different buffering systems due to the chemical modification of the proteins. Apparent size was determined by calibration of each protein against an unstained protein ladder in specific electrophoresis conditions.

Recommended reading

  • Coligan, J.E., et al., Eds. (2002). Electrophoresis, In Current Protocols in Protein Science, pp. 10.0.1-10.4.36. John Wiley and Sons, Inc. New York.
  • Bollag, D.M., Rozycki, M.D. and Edelstein, S.J. (2002). Protein Methods, 2nd ed. Wiley-Liss, Inc. New York.
  • Hames, B.D. and Rickwood, D. Eds. (1990) Gel Electrophoresis of Proteins: a Practical Approach, 2nd ed. Oxford University Press, New York.

Additional resources

For Research Use Only. Not for use in diagnostic procedures.

Sours: https://www.thermofisher.com/us/en/home/life-science/protein-biology/protein-biology-learning-center/protein-biology-resource-library/pierce-protein-methods/overview-electrophoresis.html
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  2. Powerpoint bible background
  3. 5 halo trim
  4. Working for spherion

Introduction to SDS-PAGE

This material is accompanied by a presentation on protein structure and principles behind denaturing samples and discontinuous gel electrophoresis.

The separation of macromolecules in an electric field is called electrophoresis. A very common method for separating proteins by electrophoresis uses a discontinuous polyacrylamide gel as a support medium and sodium dodecyl sulfate (SDS) to denature the proteins. The method is called sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE). The most commonly used system is also called the Laemmli method after U.K. Laemmli, who was the first to publish a paper employing SDS-PAGE in a scientific study.

SDS (also called lauryl sulfate) is an anionic detergent, meaning that when dissolved its molecules have a net negative charge within a wide pH range. A polypeptide chain binds amounts of SDS in proportion to its relative molecuar mass. The negative charges on SDS destroy most of the complex structure of proteins, and are strongly attracted toward an anode (positively-charged electrode) in an electric field.

Polyacrylamide gels restrain larger molecules from migrating as fast as smaller molecules. Because the charge-to-mass ratio is nearly the same among SDS-denatured polypeptides, the final separation of proteins is dependent almost entirely on the differences in relative molecular mass of polypeptides. In a gel of uniform density the relative migration distance of a protein (Rf, the f as a subscript) is negatively proportional to the log of its mass. If proteins of known mass are run simultaneously with the unknowns, the relationship between Rf and mass can be plotted, and the masses of unknown proteins estimated.

Protein separation by SDS-PAGE can be used to estimate relative molecular mass, to determine the relative abundance of major proteins in a sample, and to determine the distribution of proteins among fractions. The purity of protein samples can be assessed and the progress of a fractionation or purification procedure can be followed. Different staining methods can be used to detect rare proteins and to learn something about their biochemical properties. Specialized techniques such as Western blotting, two-dimensional electrophoresis, and peptide mapping can be used to detect extremely scarce gene products, to find similarities among them, and to detect and separate isoenzymes of proteins.

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.

Polyacrylamide gels for SDS-PAGE

Many systems for protein electrophoresis have been developed, and apparatus used for SDS-PAGE varies widely. The methodology used on these pages employs the Laemmli method. Reference to the Laemmli method in a materials and methods section eliminates the need to describe the buffers, casting of gels, apparatus, etc. Unless the paper employs some modification to the method, the only details of SDS-PAGE that should be reported in a methods section are percent total acrylamide (%T) in a gel, relative percentage and type of crosslinker (%C), and perhaps a reference to the gel dimensions. We use a "mini-gel" system, with 3 1/4" x 4" gel cassettes.

SDS-PAGE can be conducted on pre-cast gels, saving the trouble and hazard of working with acrylamide. The following description applies to shop-made casting and running apparatus that are much cheaper than commercially available equipment. In addition to cost effectiveness, an advantage of making one's own gels the first time is a deeper understanding of the process.

Regardless of the system, preparation requires casting two different layers of acrylamide between glass plates. The lower layer (separating, or resolving, gel) is responsible for actually separating polypeptides by size. The upper layer (stacking gel) includes the sample wells. It is designed to sweep up proteins in a sample between two moving boundaries so that they are compressed (stacked) into micrometer thin layers when they reach the separating gel.

Sours: https://www.ruf.rice.edu/~bioslabs/studies/sds-page/gellab2.html
SDS-PAGE and Native Gel Electrophoresis: denaturing vs non-denaturing gels

SDS-PAGE

biochemical technique

Proteins of the erythrocytemembrane separated by SDS-PAGE according to their molecular masses

SDS-PAGE (sodium dodecyl sulphate–polyacrylamide gel electrophoresis), is a discontinuous electrophoretic system developed by Ulrich K. Laemmli which is commonly used as a method to separate proteins with molecular masses between 5 and 250 kDa.[1][2] The combined use of sodium dodecyl sulfate (SDS, also known as sodium lauryl sulfate) and polyacrylamide gel allows to eliminate the influence of structure and charge, and proteins are separated solely on the basis of differences in their molecular weight.

Properties[edit]

Unfolding of a protein with SDS
Unfolding of a protein with heat

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]

Procedure[edit]

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]

Sample combs with different numbers of pockets, each prong leaves a pocket in the gel when pulled out
Polymerised separating and stacking gel before removing the sample comb (white) between the spacers (black), in the stacking gel are small amounts of bromophenol blue for improved visibility, the separating gel is unstained

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]

Disulfide reduction by DTT

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]

Electrophoresis chamber after a few minutes of electrophoresis. In the first pocket a size marker was applied with bromophenol blue, in the other pockets, the samples were added bromocresol green

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-chloridebuffer 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]

Coomassie-stained 10% Tris/Tricine gel. In the left lane, a molecular weight size marker was used to estimate the size (from top to bottom: 66, 45, 35, 24, 18 and 9 kDa). In the remaining lanes purified yeast proteins were separated.

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]

Two SDS gels after completed separation of the samples and staining in a drying frame

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.

Molecular mass determination[edit]

The proteins of the size marker (black X) show an approximately straight line in the representation of log M over Rf. The molecular weight of the unknown protein (red X) can be determined on the y-axis.

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]

Applications[edit]

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.

Variants[edit]

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.

Alternatives[edit]

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 photometricquantification, 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.

History[edit]

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]

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External links[edit]

Sours: https://en.wikipedia.org/wiki/SDS-PAGE

Electrophoresis kda gel

Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa

A discontinuous sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) system for the separation of proteins in the range from 1 to 100 kDa is described. Tricine, used as the trailing ion, allows a resolution of small proteins at lower acrylamide concentrations than in glycine-SDS-PAGE systems. A superior resolution of proteins, especially in the range between 5 and 20 kDa, is achieved without the necessity to use urea. Proteins above 30 kDa are already destacked within the sample gel. Thus a smooth passage of these proteins from sample to separating gel is warranted and overloading effects are reduced. This is of special importance when large amounts of protein are to be loaded onto preparative gels. The omission of glycine and urea prevents disturbances which might occur in the course of subsequent amino acid sequencing.

Sours: https://pubmed.ncbi.nlm.nih.gov/2449095/
The principle of SDS PAGE-a full and clear explanation of the technique and how does it work

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