Active fractions isolated by a preparative fractionation procedure may be subjected to a number of analytical fractionation procedures to determine their purity. Analytical fractionations are distinguished from preparative fractionations by the criteria shown in Table 1.
Table 1. The difference between preparative and analytical fractionations.
Parameters: Analytical / Preparative
Scale: Small / Large
Fate of sample: Destroyed / Preserved
Product Information: Active / fraction
In an analytical fractionation, therefore, a small amount of sample is sacrificed in order to gain information about the state of purity of the material being analysed. Of the many physico-chemical techniques which have contributed to our knowledge of proteins (and nucleic acids), electrophoretic techniques occupy a position of primary importance. Electrophoresis finds its greatest usefulness in the analysis of mixtures and in the determination of purity, although certain forms of electrophoresis may be applied on a preparative scale.
Electrophoresis is a technique used to separate and sometimes purify macromolecules - especially proteins and nucleic acids - that differ in size, charge or conformation. As such, it is one of the most widely-used techniques in biochemistry and molecular biology.
Principles of electrophoresis
Electrophoresis may be defined as the migration of charged ions in an electric field. In metal conductors, electric current flows between electrodes and is carried by ions. The negative electrode - the cathode - donates electrons and the positive electrode - the anode - takes up electrons to complete the circuit. The ions that result from the take up of electrons from the cathode will be negatively charged and will thus migrate towards the positive anode.
Because of their anodic migration, negative ions are called “anions”. Electric current is carried by the movement of electrons, largely along the surface of the metal. In solutions, the Ions which result from the donation of an electron to the electron deficient (i.e. positively charged) anode will themselves be electron deficient, and thus positively charged. These will migrate to the cathode and are thus called cations.
• The force that an ion of charge q will experience when placed in an electric field E is called the electric force (Fef) and is defined as:
Fef = q x E
• This force is opposed by the frictional force,( Ff):
Ff = f V
Where f = frictional coefficient
f = 6hpr for a sphere of radius r (Stoke’s equation)
• In a constant electric field Fef = Ff thus the velocity of a particle moving in an applied electric field, V will also be constant:
V (cm/s) = (q x E) / f = q x E / 6hpr
• Electrophoretic mobility, m, is defined as the velocity/unit electric field
m= V/ E
m= q/f (m=q/6hpr)
The electrophoretic mobility is thus a function of the charge on the protein ion and the medium through which it is traveling. Electrophoretic techniques exploit the fact that different ions have different mobilities in an electric field and so can be separated by electrophoresis.
The flow of electricity in electrophoresis is subject to the same physical laws as other forms of electricity. For example, Ohm's law applies:
I=V/R
Where I = current (amps)
V = potential difference (volts)
R = resistance (ohms).
The unit of electrical charge is the coulomb and the unit of current [the ampere (amp)] may be defined as coulombs sec-1.
The flow- of electricity involves work, which generates heat, and the work (W, in joules) done in transferring a charge of q coulombs between a potential difference of V volts is:-
W=q.V=I.tV
W= I2R.t (Jouleís law of heating)
Which means that I2R.t joules of heat will be developed in the conductor.
The power (in watts) (defined as the rate of work) gives the rate of heating (joules sec-1).
Watts= I2R.t/t= I2R
Heating of the electrophoretic medium has the following effects
1. An increased rate of sample diffusion.
2. Mixing of separated samples due to convection current.
3. Thermal instability, protein denaturation and loss of enzyme activity.
4. Decrease in buffer viscosity and reduction in the resistivity of the medium.
Support Media
GEL ELECTROPHORESIS
Electrophoresis of macromolecules can be carried out in solution. However, the ability to separate molecules is compromised by their diffusion. Greater resolution is achieved if electrophoresis is carried out on semi-solid supports such as polyacrylamide or agarose gels. Gels are formed by cross-linking polymers in aqueous medium. This will form a 3-dimensional meshwork which the molecules must pass through. Polyacrylamide is a common gel for protein electrophoresis whereas agarose is more commonly used for nucleic acids. Agarose gels have a larger pore size than acrylamide gels and are better suited for larger macromolecules. However, either type of gel can be applied to either nucleic acids or proteins depending on the application.
Gels are formed from long polymers in a cross-linked lattice. The space between the polymers are the pores. Higher concentrations of the polymer will result in smaller average pore sizes. Polyacrylamide gels are formed by covalently cross-linking acrylamide monomers with bis-acrylamide with a free radical like persulfate (SO4•). The cross-linking of the acrylamide polymers results in 'pores' of a defined size. The total acrylamide concentration and the ratio of bis-acrylamide to acrylamide will determine the average pore size. The polyacrylamide solution is poured into a mold and polymerized. This mold can be a cylindrical tube, but is usually a 'slab' poured between two glass plates
Since the gel is solid with respect to the mold, all molecules are forced through the gel. Smaller molecules will be able to pass through this lattice more easily resulting in larger molecules having a lower mobility than smaller molecules. In other words, the gel acts like a molecular sieve and retains the larger molecules while letting the smaller ones pass through. (This is opposite of gel filtration where the larger molecules have a higher mobility because they to not enter the gel.) Therefore, the frictional coefficient is related to how easily a protein passes through the pores of the gel and size will be the major determinant of the mobility of molecules in a gel matrix. Protein shape and other factors will still affect mobility, but to a lesser extent. Substituting size for the frictional coefficient results in:
Agarose is a polysaccharide extracted from seaweed. It is typically used at concentrations of 0.5 to 2%. The higher the agarose concentration the "stiffer" the gel. Agarose gels are extremely easy to prepare: you simply mix agarose powder with buffer solution, melt it by heating, and pour the gel. It is also non-toxic.
Agarose gels have a large range of separation, but relatively low resolving power. By varying the concentration of agarose, fragments of DNA from about 200 to 50,000 bp can be separated using standard electrophoretic techniques.
The length of the polymer chains is dictated by the concentration of acrylamide used, which is typically between 3.5 and 20%. Polyacrylamide gels are significantly more annoying to prepare than agarose gels. Because oxygen inhibits the polymerization process, they must be poured between glass plates (or cylinders).
Acrylamide is a potent neurotoxin and should be handled with care! Wear disposable gloves when handling solutions of acrylamide, and a mask when weighing out powder. Polyacrylamide is considered to be non-toxic, but polyacrylamide gels should also be handled with gloves due to the possible presence of free acrylamide.
Polyacrylamide gels have a rather small range of separation, but very high resolving power. In the case of DNA, polyacrylamide is used for separating fragments of less than about 500 bp. However, under appropriate conditions, fragments of DNA differing is length by a single base pair are easily resolved. In contrast to agarose, polyacrylamide gels are used extensively for separating and characterizing mixtures of proteins.
PAGE
1. NONDENATURING (Native) PAGE
Nondenaturing system is used to separate intact proteins, especially oligomeric proteins, by a nondestructive means for later assessment of biological activity. On nondenaturing system, protein separation depends on a combination of differences in molecular size and shape as well as charge. Separation by size is accomplished by varying the pore size of the acrylamide polymer as a function of both the concentration of the acrylamide (range about 3-30%, w/v) and the amount of cross-linker used. In general, the higher the acrylamide concentration, the smaller the protein that remains in gel: this can be counteracted by decreasing the amount of cross-linker used, which in turn increases the degree of gel swelling during standard staining and washing procedures. Separation by charge in nondenaturing gel systems is permitted because the protein separation can be performed at any pH between 3 and 11 to allow for maximal charge differences neighboring protein species. These include the molecular weight of the stability in the range of pH 4 to 9. Once these parameters are known optimal resolution can be obtained by varying certain components within a single gel system. For example the initial choice of the gel system for separation of acidic protein of molecular weight 20,000 might be a system operating at an alkaline pH i.e.9 with an acylamide concentration of 12-15 %.
It should be noted that biologically active proteins such as enzymes often require specials electrophoresis conditions in order to retain activity after separation. For example the heat lability of the some enzymes requires that electrophoresis be conducted most conveniently in a refrigerated room or if necessary using circulating water of 0-4C in the cooling jacket of the slab gel apparatus. In addition ammonium persulphate an agent commonly used in gel polymerization reaction often interferes with enzymes activity after elution from the gel. For this reason the photo activated polymerizing agent riboflavin can also be used instead of ammonium persulphate in the preparation of the stacking gel. For the various systems if ammonium persulphate affects enzyme activity then preelectrophoresis can be used. Dithioithreitol (40-100 mM) or 2-mercaptoethanol (upto 1 M) may be included in the sample buffer to reduce some disulfide linkages, although detergent denaturing is often necessary for complete disulfide reduction.
2. SDS- PAGE
Acrylamide mixed with bisacrylamide forms a cross-linked polymer network when the polymerizing agent, ammonium persulfate, is added (Figure 1). The ammonium persulfate produces free radicals faster in the presence of TEMED (N, N, N, N’-tetramethylenediamine). The size of the pores created in the gel is inversely related to the amount of acrylamide used. Gels with a low percentage of acrylamide are typically used to resolve large proteins and high percentage gels are used to resolve small proteins.
Figure 2. Polymerization and cross-linking of acrylamide.
However, polyacrylamide of sufficient stability will always be so dense, that friction influences the migration of proteins in the gel. The electrophoretic mobility of a protein is determined mostly by its net charge per unit mass at the given pH, but is inversely proportional to its frictional coefficient in the gel, determined by the proteins size and shape. In the denaturing, reductive variant of PAGE, SDS-PAGE all differences between the proteins in charge per unit mass has been eliminated by the SDS (sodium dodecylsulphate) and the proteins migrate solely according to subunit size. The charged SDS molecules bind all along the polypetide chain, giving the chain equal charge per unit length. Thus, the denatured polypeptide chains are separated electrophoretically only according to their subunit length. The polyacrylamide gels may be cast as i fixed concentration gel (e.g. 10%) or as a gradient gel (e.g. 7-15 %). There is also a choice between continuous and discontinuous buffer systems. If the same buffer ions are present throughout sample, gel and electrophoresis buffer the electrophoresis is referred to as continuous buffer PAGE. In discontinuous buffer PAGE the separation is usually preceded by a stacking of the sample to a narrow band in a low-porosity stacking gel.
The high pore size stacking gel is to concentrate the protein sample into a sharp band before it inters to the main separating gel. This is achieved by utilizing differences in the ionic strength and pH between the electrophoresis buffer and the stacking gel. The band sharpening effect relies on the fact that negatively charged glycine ions (in running buffer) have a lower electrophoretic mobility than do the protein-SDS complex which in turn has lower mobility than the chloride ion of the loaded buffer and the stacking gel. When current is switched on, all the ionic species have to migrate at the same speed otherwise there would be a break in the electrical current. The glycinate ions can only move at the same speed as Cl- ion if they are in the region of the higher field strength. Field strength is inversely proportional to conductivity which is proportional to conc. The result is that three species of interest adjust their concentrations so that [Cl-] > [protein-SDS] > [glycinate ions]. There is only a small quantity of protein-SDS complexes so they concentrate in a very tight bond between glycinate and Cl- boundaries. Once the glycinate reaches the separating gel it becomes more fully ionized in higher pH environment and its mobility increases. Thus the interface between the glycinate and Cl- leaves behind the protein-SDS complexes which are left to electrophoresis at their own rates. The negatively charged protein SDS complexes now continue to move towards the anode and because they have the same charge per unit length they travel into the separating gel under the applied electric field with the same mobility .However as they pass through the separating gel the protein separate owing to the molecular sieving properties of the gel. Detection of the separated protein bands may be done in several ways. Staining for protein with Coomassie Brilliant Blue G-250 is a standard procedure detecting 0.1 to 1µg per band (depending on band dimensions). Coomassie Brilliant Blue G- 250 can also be used and by using dilute staining solution, destaining can be omitted. An increase in detection sensitivity of approximately a factor 100 can be obtained with silver staining.
Determination of Molecular Weight
This is done by SDS-PAGE of proteins known molecular weight along with the proteins to be characterized. A linear relationship exists between the logarithm of the molecular weight of an SDS-denatured polypeptide and its Rf value. The Rf is calculated as the ratio of the distance migrated by the molecule to that migrated by a marker dye-front. A simple way of determining relative molecular weight by electrophoresis (Mr) is to plot a standard curve of distance migrated vs. log10MW for known samples, and read off the logMr of the sample after measuring distance migrated on the same gel.
Isoelectric focusing (IEF)
IEF, also known as electrofocusing, is a technique for separating different molecules by their electric charge differences. It is a type of zone electrophoresis, usually performed in a gel, that takes advantage of the fact that a molecule's charge changes with the pH of its surroundings. A protein which is in a pH region below its pI will be positively charged and so will migrate towards the cathode. However, as it migrates, the charge will decrease until the protein reaches a pH which is its pI. At this point it has no net charge and so migration ceases. Should it overshoot this point, it will enter a region of pH above its pI and so become negatively charged. It will then reverse its direction of migration and now migrate towards the anode. Therefore proteins become focused into sharp stationary bands with each protein positioned at a point in the pH gradient corresponding to its pI. The technique is capable of extremely high resolution with proteins differing by a single charge being fractionated into separate bands.
Molecules to be focused are distributed over a medium that has a pH gradient (usually created by aliphatic ampholytes). An electric current is passed through the medium, creating a "positive" anode and "negative" cathode end. Negatively charged molecules migrate through the pH gradient in the medium toward the "positive" end while positively charged molecules move toward the "negative" end. As a particle moves towards the pole opposite of its charge it moves through the changing pH gradient until it reaches a point in which the pH of that molecules isoelectric point is reached. At this point the molecule no longer has a net electric charge (due to the protonation or deprotonation of the associated functional groups) and as such will not proceed any further within the gel. The gradient is initially established before adding the particles of interest by first subjecting a solution of small molecules such as polyampholytes with varying pI values to electrophoresis.
The method is applied particularly often in the study of proteins, which separate based on their relative content of acidic and basic residues, whose value is represented by the pI. Proteins are introduced into a Immobilized pH gradient gel composed of polyacrylamide, starch, or agarose where a pH gradient has been established. Gels with large pores are usually used in this process to eliminate any "sieving" effects, or artifacts in the pI caused by differing migration rates for proteins of differing sizes. Isoelectric focusing can resolve proteins that differ in pI value by as little as 0.01. Isoelectric focusing is the first step in two-dimensional gel electrophoresis, in which proteins are first separated by their pI and then further separated by molecular weight through SDS-PAGE.
Table 1. The difference between preparative and analytical fractionations.
Parameters: Analytical / Preparative
Scale: Small / Large
Fate of sample: Destroyed / Preserved
Product Information: Active / fraction
In an analytical fractionation, therefore, a small amount of sample is sacrificed in order to gain information about the state of purity of the material being analysed. Of the many physico-chemical techniques which have contributed to our knowledge of proteins (and nucleic acids), electrophoretic techniques occupy a position of primary importance. Electrophoresis finds its greatest usefulness in the analysis of mixtures and in the determination of purity, although certain forms of electrophoresis may be applied on a preparative scale.
Electrophoresis is a technique used to separate and sometimes purify macromolecules - especially proteins and nucleic acids - that differ in size, charge or conformation. As such, it is one of the most widely-used techniques in biochemistry and molecular biology.
Principles of electrophoresis
Electrophoresis may be defined as the migration of charged ions in an electric field. In metal conductors, electric current flows between electrodes and is carried by ions. The negative electrode - the cathode - donates electrons and the positive electrode - the anode - takes up electrons to complete the circuit. The ions that result from the take up of electrons from the cathode will be negatively charged and will thus migrate towards the positive anode.
Because of their anodic migration, negative ions are called “anions”. Electric current is carried by the movement of electrons, largely along the surface of the metal. In solutions, the Ions which result from the donation of an electron to the electron deficient (i.e. positively charged) anode will themselves be electron deficient, and thus positively charged. These will migrate to the cathode and are thus called cations.
• The force that an ion of charge q will experience when placed in an electric field E is called the electric force (Fef) and is defined as:
Fef = q x E
• This force is opposed by the frictional force,( Ff):
Ff = f V
Where f = frictional coefficient
f = 6hpr for a sphere of radius r (Stoke’s equation)
• In a constant electric field Fef = Ff thus the velocity of a particle moving in an applied electric field, V will also be constant:
V (cm/s) = (q x E) / f = q x E / 6hpr
• Electrophoretic mobility, m, is defined as the velocity/unit electric field
m= V/ E
m= q/f (m=q/6hpr)
The electrophoretic mobility is thus a function of the charge on the protein ion and the medium through which it is traveling. Electrophoretic techniques exploit the fact that different ions have different mobilities in an electric field and so can be separated by electrophoresis.
The flow of electricity in electrophoresis is subject to the same physical laws as other forms of electricity. For example, Ohm's law applies:
I=V/R
Where I = current (amps)
V = potential difference (volts)
R = resistance (ohms).
The unit of electrical charge is the coulomb and the unit of current [the ampere (amp)] may be defined as coulombs sec-1.
The flow- of electricity involves work, which generates heat, and the work (W, in joules) done in transferring a charge of q coulombs between a potential difference of V volts is:-
W=q.V=I.tV
W= I2R.t (Jouleís law of heating)
Which means that I2R.t joules of heat will be developed in the conductor.
The power (in watts) (defined as the rate of work) gives the rate of heating (joules sec-1).
Watts= I2R.t/t= I2R
Heating of the electrophoretic medium has the following effects
1. An increased rate of sample diffusion.
2. Mixing of separated samples due to convection current.
3. Thermal instability, protein denaturation and loss of enzyme activity.
4. Decrease in buffer viscosity and reduction in the resistivity of the medium.
Support Media
GEL ELECTROPHORESIS
Electrophoresis of macromolecules can be carried out in solution. However, the ability to separate molecules is compromised by their diffusion. Greater resolution is achieved if electrophoresis is carried out on semi-solid supports such as polyacrylamide or agarose gels. Gels are formed by cross-linking polymers in aqueous medium. This will form a 3-dimensional meshwork which the molecules must pass through. Polyacrylamide is a common gel for protein electrophoresis whereas agarose is more commonly used for nucleic acids. Agarose gels have a larger pore size than acrylamide gels and are better suited for larger macromolecules. However, either type of gel can be applied to either nucleic acids or proteins depending on the application.
Gels are formed from long polymers in a cross-linked lattice. The space between the polymers are the pores. Higher concentrations of the polymer will result in smaller average pore sizes. Polyacrylamide gels are formed by covalently cross-linking acrylamide monomers with bis-acrylamide with a free radical like persulfate (SO4•). The cross-linking of the acrylamide polymers results in 'pores' of a defined size. The total acrylamide concentration and the ratio of bis-acrylamide to acrylamide will determine the average pore size. The polyacrylamide solution is poured into a mold and polymerized. This mold can be a cylindrical tube, but is usually a 'slab' poured between two glass plates
Since the gel is solid with respect to the mold, all molecules are forced through the gel. Smaller molecules will be able to pass through this lattice more easily resulting in larger molecules having a lower mobility than smaller molecules. In other words, the gel acts like a molecular sieve and retains the larger molecules while letting the smaller ones pass through. (This is opposite of gel filtration where the larger molecules have a higher mobility because they to not enter the gel.) Therefore, the frictional coefficient is related to how easily a protein passes through the pores of the gel and size will be the major determinant of the mobility of molecules in a gel matrix. Protein shape and other factors will still affect mobility, but to a lesser extent. Substituting size for the frictional coefficient results in:
Agarose is a polysaccharide extracted from seaweed. It is typically used at concentrations of 0.5 to 2%. The higher the agarose concentration the "stiffer" the gel. Agarose gels are extremely easy to prepare: you simply mix agarose powder with buffer solution, melt it by heating, and pour the gel. It is also non-toxic.
Agarose gels have a large range of separation, but relatively low resolving power. By varying the concentration of agarose, fragments of DNA from about 200 to 50,000 bp can be separated using standard electrophoretic techniques.
The length of the polymer chains is dictated by the concentration of acrylamide used, which is typically between 3.5 and 20%. Polyacrylamide gels are significantly more annoying to prepare than agarose gels. Because oxygen inhibits the polymerization process, they must be poured between glass plates (or cylinders).
Acrylamide is a potent neurotoxin and should be handled with care! Wear disposable gloves when handling solutions of acrylamide, and a mask when weighing out powder. Polyacrylamide is considered to be non-toxic, but polyacrylamide gels should also be handled with gloves due to the possible presence of free acrylamide.
Polyacrylamide gels have a rather small range of separation, but very high resolving power. In the case of DNA, polyacrylamide is used for separating fragments of less than about 500 bp. However, under appropriate conditions, fragments of DNA differing is length by a single base pair are easily resolved. In contrast to agarose, polyacrylamide gels are used extensively for separating and characterizing mixtures of proteins.
PAGE
1. NONDENATURING (Native) PAGE
Nondenaturing system is used to separate intact proteins, especially oligomeric proteins, by a nondestructive means for later assessment of biological activity. On nondenaturing system, protein separation depends on a combination of differences in molecular size and shape as well as charge. Separation by size is accomplished by varying the pore size of the acrylamide polymer as a function of both the concentration of the acrylamide (range about 3-30%, w/v) and the amount of cross-linker used. In general, the higher the acrylamide concentration, the smaller the protein that remains in gel: this can be counteracted by decreasing the amount of cross-linker used, which in turn increases the degree of gel swelling during standard staining and washing procedures. Separation by charge in nondenaturing gel systems is permitted because the protein separation can be performed at any pH between 3 and 11 to allow for maximal charge differences neighboring protein species. These include the molecular weight of the stability in the range of pH 4 to 9. Once these parameters are known optimal resolution can be obtained by varying certain components within a single gel system. For example the initial choice of the gel system for separation of acidic protein of molecular weight 20,000 might be a system operating at an alkaline pH i.e.9 with an acylamide concentration of 12-15 %.
It should be noted that biologically active proteins such as enzymes often require specials electrophoresis conditions in order to retain activity after separation. For example the heat lability of the some enzymes requires that electrophoresis be conducted most conveniently in a refrigerated room or if necessary using circulating water of 0-4C in the cooling jacket of the slab gel apparatus. In addition ammonium persulphate an agent commonly used in gel polymerization reaction often interferes with enzymes activity after elution from the gel. For this reason the photo activated polymerizing agent riboflavin can also be used instead of ammonium persulphate in the preparation of the stacking gel. For the various systems if ammonium persulphate affects enzyme activity then preelectrophoresis can be used. Dithioithreitol (40-100 mM) or 2-mercaptoethanol (upto 1 M) may be included in the sample buffer to reduce some disulfide linkages, although detergent denaturing is often necessary for complete disulfide reduction.
2. SDS- PAGE
Acrylamide mixed with bisacrylamide forms a cross-linked polymer network when the polymerizing agent, ammonium persulfate, is added (Figure 1). The ammonium persulfate produces free radicals faster in the presence of TEMED (N, N, N, N’-tetramethylenediamine). The size of the pores created in the gel is inversely related to the amount of acrylamide used. Gels with a low percentage of acrylamide are typically used to resolve large proteins and high percentage gels are used to resolve small proteins.
Figure 2. Polymerization and cross-linking of acrylamide.
However, polyacrylamide of sufficient stability will always be so dense, that friction influences the migration of proteins in the gel. The electrophoretic mobility of a protein is determined mostly by its net charge per unit mass at the given pH, but is inversely proportional to its frictional coefficient in the gel, determined by the proteins size and shape. In the denaturing, reductive variant of PAGE, SDS-PAGE all differences between the proteins in charge per unit mass has been eliminated by the SDS (sodium dodecylsulphate) and the proteins migrate solely according to subunit size. The charged SDS molecules bind all along the polypetide chain, giving the chain equal charge per unit length. Thus, the denatured polypeptide chains are separated electrophoretically only according to their subunit length. The polyacrylamide gels may be cast as i fixed concentration gel (e.g. 10%) or as a gradient gel (e.g. 7-15 %). There is also a choice between continuous and discontinuous buffer systems. If the same buffer ions are present throughout sample, gel and electrophoresis buffer the electrophoresis is referred to as continuous buffer PAGE. In discontinuous buffer PAGE the separation is usually preceded by a stacking of the sample to a narrow band in a low-porosity stacking gel.
The high pore size stacking gel is to concentrate the protein sample into a sharp band before it inters to the main separating gel. This is achieved by utilizing differences in the ionic strength and pH between the electrophoresis buffer and the stacking gel. The band sharpening effect relies on the fact that negatively charged glycine ions (in running buffer) have a lower electrophoretic mobility than do the protein-SDS complex which in turn has lower mobility than the chloride ion of the loaded buffer and the stacking gel. When current is switched on, all the ionic species have to migrate at the same speed otherwise there would be a break in the electrical current. The glycinate ions can only move at the same speed as Cl- ion if they are in the region of the higher field strength. Field strength is inversely proportional to conductivity which is proportional to conc. The result is that three species of interest adjust their concentrations so that [Cl-] > [protein-SDS] > [glycinate ions]. There is only a small quantity of protein-SDS complexes so they concentrate in a very tight bond between glycinate and Cl- boundaries. Once the glycinate reaches the separating gel it becomes more fully ionized in higher pH environment and its mobility increases. Thus the interface between the glycinate and Cl- leaves behind the protein-SDS complexes which are left to electrophoresis at their own rates. The negatively charged protein SDS complexes now continue to move towards the anode and because they have the same charge per unit length they travel into the separating gel under the applied electric field with the same mobility .However as they pass through the separating gel the protein separate owing to the molecular sieving properties of the gel. Detection of the separated protein bands may be done in several ways. Staining for protein with Coomassie Brilliant Blue G-250 is a standard procedure detecting 0.1 to 1µg per band (depending on band dimensions). Coomassie Brilliant Blue G- 250 can also be used and by using dilute staining solution, destaining can be omitted. An increase in detection sensitivity of approximately a factor 100 can be obtained with silver staining.
Determination of Molecular Weight
This is done by SDS-PAGE of proteins known molecular weight along with the proteins to be characterized. A linear relationship exists between the logarithm of the molecular weight of an SDS-denatured polypeptide and its Rf value. The Rf is calculated as the ratio of the distance migrated by the molecule to that migrated by a marker dye-front. A simple way of determining relative molecular weight by electrophoresis (Mr) is to plot a standard curve of distance migrated vs. log10MW for known samples, and read off the logMr of the sample after measuring distance migrated on the same gel.
Isoelectric focusing (IEF)
IEF, also known as electrofocusing, is a technique for separating different molecules by their electric charge differences. It is a type of zone electrophoresis, usually performed in a gel, that takes advantage of the fact that a molecule's charge changes with the pH of its surroundings. A protein which is in a pH region below its pI will be positively charged and so will migrate towards the cathode. However, as it migrates, the charge will decrease until the protein reaches a pH which is its pI. At this point it has no net charge and so migration ceases. Should it overshoot this point, it will enter a region of pH above its pI and so become negatively charged. It will then reverse its direction of migration and now migrate towards the anode. Therefore proteins become focused into sharp stationary bands with each protein positioned at a point in the pH gradient corresponding to its pI. The technique is capable of extremely high resolution with proteins differing by a single charge being fractionated into separate bands.
Molecules to be focused are distributed over a medium that has a pH gradient (usually created by aliphatic ampholytes). An electric current is passed through the medium, creating a "positive" anode and "negative" cathode end. Negatively charged molecules migrate through the pH gradient in the medium toward the "positive" end while positively charged molecules move toward the "negative" end. As a particle moves towards the pole opposite of its charge it moves through the changing pH gradient until it reaches a point in which the pH of that molecules isoelectric point is reached. At this point the molecule no longer has a net electric charge (due to the protonation or deprotonation of the associated functional groups) and as such will not proceed any further within the gel. The gradient is initially established before adding the particles of interest by first subjecting a solution of small molecules such as polyampholytes with varying pI values to electrophoresis.
The method is applied particularly often in the study of proteins, which separate based on their relative content of acidic and basic residues, whose value is represented by the pI. Proteins are introduced into a Immobilized pH gradient gel composed of polyacrylamide, starch, or agarose where a pH gradient has been established. Gels with large pores are usually used in this process to eliminate any "sieving" effects, or artifacts in the pI caused by differing migration rates for proteins of differing sizes. Isoelectric focusing can resolve proteins that differ in pI value by as little as 0.01. Isoelectric focusing is the first step in two-dimensional gel electrophoresis, in which proteins are first separated by their pI and then further separated by molecular weight through SDS-PAGE.
Two-dimensional gel electrophoresis
Two-dimensional gel electrophoresis, abbreviated as 2-DE or 2-D electrophoresis, is a form of gel electrophoresis commonly used to analyze proteins. Mixtures of proteins are separated by two properties in two dimensions on 2D gels.
Basis for separation
2-D electrophoresis begins with 1-D electrophoresis but then separates the molecules by a second property in a direction 90 degrees from the first. In 1-D electrophoresis, proteins (or other molecules) are separated in one dimension, so that all the proteins/molecules will lie along a lane but be separated from each other by a property (e.g. isoelectric point). The result is that the molecules are spread out across a 2-D gel. Because it is unlikely that two molecules will be similar in two distinct properties, molecules are more effectively separated in 2-D electrophoresis than in 1-D electrophoresis.
The two dimensions that proteins are separated into using this technique can be isoelectric point, protein complex mass in the native state, and protein mass.
To separate the proteins by isoelectric point is called isoelectric focusing (IEF). Thereby, a gradient of pH is applied to a gel and an electric potential is applied across the gel, making one end more positive than the other. At all pHs other than their isoelectric point, proteins will be charged. If they are positively charged, they will be pulled towards the more negative end of the gel and if they are negatively charged they will be pulled to the more positive end of the gel. The proteins applied in the first dimension will move along the gel and will accumulate at their isoelectric point; that is, the point at which the overall charge on the protein is 0 (a neutral charge).
For the analysis of the functioning of proteins in a cell, the knowledge of their cooperation is essential. Most often proteins act together in complexes to be fully functional. The analysis of this sub organelle organisation of the cell requires techniques conserving the native state of the protein complexes. In native polyacrylamide gel electrophoresis (native PAGE), proteins remain in their native state and are separated in the electric field following their mass and the mass of their complexes respectively. To obtain a separation by size and not by net charge, as in IEF, an additional charge is transferred to the proteins by the use of coomassie or lithium dodecyl sulfate (LDS). After completion of the first dimension the complexes are destroyed by applying the denaturing SDS-PAGE in the second dimension, where the proteins of which the complexes are composed of are separated by their mass.
Before separating the proteins by mass, they are treated with sodium dodecyl sulfate (SDS) along with other reagents (SDS-PAGE in 1-D). This denatures the proteins (that is, it unfolds them into long, straight molecules) and binds a number of SDS molecules roughly proportional to the protein's length. Because a protein's length (when unfolded) is roughly proportional to its mass, this is equivalent to saying that it attaches a number of SDS molecules roughly proportional to the protein's mass. Since the SDS molecules are negatively charged, the result of this is that all of the proteins will have approximately the same mass-to-charge ratio as each other. In addition, proteins will not migrate when they have no charge (a result of the isoelectric focusing step) therefore the coating of the protein in SDS (negatively charged) allows migration of the proteins in the second dimension (NB SDS is not compatible for use in the first dimension as it is charged and a nonionic or zwitterionic detergent needs to be used). In the second dimension, an electric potential is again applied, but at a 90 degree angle from the first field. The proteins will be attracted to the more positive side of the gel proportionally to their mass-to-charge ratio. As previously explained, this ratio will be nearly the same for all proteins. The proteins' progress will be slowed by frictional forces. The gel therefore acts like a molecular sieve when the current is applied, separating the proteins on the basis of their molecular weight with larger proteins being retained higher in the gel and smaller proteins being able to pass through the sieve and reach lower regions of the gel.
The result of this is a gel with proteins spread out on its surface. These proteins can then be detected by a variety of means, but the most commonly used stains are silver and coomassie staining. In this case, a silver colloid is applied to the gel. The silver binds to cysteine groups within the protein. The silver is darkened by exposure to ultra-violet light. The darkness of the silver can be related to the amount of silver and therefore the amount of protein at a given location on the gel. This measurement can only give approximate amounts, but is adequate for most purposes.
Molecules other than proteins can be separated by 2D electrophoresis. In supercoiling assays, coiled DNA is separated in the first dimension and denatured by a DNA intercalator (such as ethidium bromide or the less carcinogenic chloroquine) in the second. This is comparable to the combination of native PAGE /SDS-PAGE in protein separation.
In summary 2D provides resolution according to two traits, whereof one is most often molecular charge. The investigated molecule needs not be protein.
In quantitative proteomics, these tools primarily analyze bio-markers by quantifying individual proteins, and showing the separation between one or more protein "spots" on a scanned image of a 2-DE gel. Additionally, these tools match spots between gels of similar samples to show, for example, proteomic differences between early and advanced stages of an illness. Software packages include Delta2D, PDQuest and Progenesis - among others. While this technology is widely utilized, the intelligence has not been perfected. For example, while PDQuest and Progenesis tend to agree on the quantification and analysis of well-defined well-separated protein spots, they deliver different results and analysis tendencies with less-defined less-separated spots.
Challenges for automatic software-based analysis include:
• incompletely separated (overlapping) spots (less-defined and/or separated)
• weak spots / noise (e.g., "ghost spots")
• running differences between gels (e.g., protein migrates to different positions on different gels)
• unmatched/undetected spots, leading to missing values[2]
• differences in software algorithms and therefore analysis tendencies
New approaches taken by software packages, such as Delta2D and Progenesis, go a long way in compensating for running differences between gels and solving the missing values problem to allow for robust statistical analysis of 2-DE gels. Generated picking lists can be used for the automated in-gel digestion of protein spots, and subsequent identification of the proteins by mass spectrometry. Although 2-DE automated image analysis technology has not been perfected, manual visual analysis is no longer realistic for objective and statistically valid experiment designs.
Two-dimensional gel electrophoresis, abbreviated as 2-DE or 2-D electrophoresis, is a form of gel electrophoresis commonly used to analyze proteins. Mixtures of proteins are separated by two properties in two dimensions on 2D gels.
Basis for separation
2-D electrophoresis begins with 1-D electrophoresis but then separates the molecules by a second property in a direction 90 degrees from the first. In 1-D electrophoresis, proteins (or other molecules) are separated in one dimension, so that all the proteins/molecules will lie along a lane but be separated from each other by a property (e.g. isoelectric point). The result is that the molecules are spread out across a 2-D gel. Because it is unlikely that two molecules will be similar in two distinct properties, molecules are more effectively separated in 2-D electrophoresis than in 1-D electrophoresis.
The two dimensions that proteins are separated into using this technique can be isoelectric point, protein complex mass in the native state, and protein mass.
To separate the proteins by isoelectric point is called isoelectric focusing (IEF). Thereby, a gradient of pH is applied to a gel and an electric potential is applied across the gel, making one end more positive than the other. At all pHs other than their isoelectric point, proteins will be charged. If they are positively charged, they will be pulled towards the more negative end of the gel and if they are negatively charged they will be pulled to the more positive end of the gel. The proteins applied in the first dimension will move along the gel and will accumulate at their isoelectric point; that is, the point at which the overall charge on the protein is 0 (a neutral charge).
For the analysis of the functioning of proteins in a cell, the knowledge of their cooperation is essential. Most often proteins act together in complexes to be fully functional. The analysis of this sub organelle organisation of the cell requires techniques conserving the native state of the protein complexes. In native polyacrylamide gel electrophoresis (native PAGE), proteins remain in their native state and are separated in the electric field following their mass and the mass of their complexes respectively. To obtain a separation by size and not by net charge, as in IEF, an additional charge is transferred to the proteins by the use of coomassie or lithium dodecyl sulfate (LDS). After completion of the first dimension the complexes are destroyed by applying the denaturing SDS-PAGE in the second dimension, where the proteins of which the complexes are composed of are separated by their mass.
Before separating the proteins by mass, they are treated with sodium dodecyl sulfate (SDS) along with other reagents (SDS-PAGE in 1-D). This denatures the proteins (that is, it unfolds them into long, straight molecules) and binds a number of SDS molecules roughly proportional to the protein's length. Because a protein's length (when unfolded) is roughly proportional to its mass, this is equivalent to saying that it attaches a number of SDS molecules roughly proportional to the protein's mass. Since the SDS molecules are negatively charged, the result of this is that all of the proteins will have approximately the same mass-to-charge ratio as each other. In addition, proteins will not migrate when they have no charge (a result of the isoelectric focusing step) therefore the coating of the protein in SDS (negatively charged) allows migration of the proteins in the second dimension (NB SDS is not compatible for use in the first dimension as it is charged and a nonionic or zwitterionic detergent needs to be used). In the second dimension, an electric potential is again applied, but at a 90 degree angle from the first field. The proteins will be attracted to the more positive side of the gel proportionally to their mass-to-charge ratio. As previously explained, this ratio will be nearly the same for all proteins. The proteins' progress will be slowed by frictional forces. The gel therefore acts like a molecular sieve when the current is applied, separating the proteins on the basis of their molecular weight with larger proteins being retained higher in the gel and smaller proteins being able to pass through the sieve and reach lower regions of the gel.
The result of this is a gel with proteins spread out on its surface. These proteins can then be detected by a variety of means, but the most commonly used stains are silver and coomassie staining. In this case, a silver colloid is applied to the gel. The silver binds to cysteine groups within the protein. The silver is darkened by exposure to ultra-violet light. The darkness of the silver can be related to the amount of silver and therefore the amount of protein at a given location on the gel. This measurement can only give approximate amounts, but is adequate for most purposes.
Molecules other than proteins can be separated by 2D electrophoresis. In supercoiling assays, coiled DNA is separated in the first dimension and denatured by a DNA intercalator (such as ethidium bromide or the less carcinogenic chloroquine) in the second. This is comparable to the combination of native PAGE /SDS-PAGE in protein separation.
In summary 2D provides resolution according to two traits, whereof one is most often molecular charge. The investigated molecule needs not be protein.
In quantitative proteomics, these tools primarily analyze bio-markers by quantifying individual proteins, and showing the separation between one or more protein "spots" on a scanned image of a 2-DE gel. Additionally, these tools match spots between gels of similar samples to show, for example, proteomic differences between early and advanced stages of an illness. Software packages include Delta2D, PDQuest and Progenesis - among others. While this technology is widely utilized, the intelligence has not been perfected. For example, while PDQuest and Progenesis tend to agree on the quantification and analysis of well-defined well-separated protein spots, they deliver different results and analysis tendencies with less-defined less-separated spots.
Challenges for automatic software-based analysis include:
• incompletely separated (overlapping) spots (less-defined and/or separated)
• weak spots / noise (e.g., "ghost spots")
• running differences between gels (e.g., protein migrates to different positions on different gels)
• unmatched/undetected spots, leading to missing values[2]
• differences in software algorithms and therefore analysis tendencies
New approaches taken by software packages, such as Delta2D and Progenesis, go a long way in compensating for running differences between gels and solving the missing values problem to allow for robust statistical analysis of 2-DE gels. Generated picking lists can be used for the automated in-gel digestion of protein spots, and subsequent identification of the proteins by mass spectrometry. Although 2-DE automated image analysis technology has not been perfected, manual visual analysis is no longer realistic for objective and statistically valid experiment designs.
Western blot
The western blot (alternately, immunoblot) is a method of detecting specific proteins in a given sample of tissue homogenate or extract. It uses gel electrophoresis to separate native or denatured proteins by the length of the polypeptide (denaturing conditions) or by the 3-D structure of the protein (native/ non-denaturing conditions). The proteins are then transferred to a membrane (typically nitrocellulose or PVDF), where they are probed (detected) using antibodies specific to the target protein. There are now many reagent companies that specialize in providing antibodies (both monoclonal and polyclonal antibodies) against many thousands of different proteins. Commercial antibodies can be expensive, though the unbound antibody can be reused between experiments. This method is used in the fields of molecular biology, biochemistry, immunogenetics and other molecular biology disciplines.
Other related techniques include using antibodies to detect proteins in tissues and cells by immunostaining and enzyme-linked immunosorbent assay (ELISA).
• The method originated from the laboratory of George Stark at Stanford. The name western blot was given to the technique by W. Neal Burnette[1] and is a play on the name Southern blot, a technique for DNA detection developed earlier by Edwin Southern. Detection of RNA is termed northern blotting.
Steps in a Western blot
1. Tissue preparation
Samples may be taken from whole tissue or from cell culture. In most cases, solid tissues are first broken down mechanically using a blender (for larger sample volumes), using a homogenizer (smaller volumes), or by sonication. Cells may also be broken open by one of the above mechanical methods. However, it should be noted that bacteria, virus or environmental samples can be the source of protein and thus Western blotting is not restricted to cellular studies only.
Assorted detergents, salts, and buffers may be employed to encourage lysis of cells and to solubilize proteins. Protease and phosphatase inhibitors are often added to prevent the digestion of the sample by its own enzymes.
A combination of biochemical and mechanical techniques – including various types of filtration and centrifugation – can be used to separate different cell compartments and organelles.
2. Gel electrophoresis
The proteins of the sample are separated using gel electrophoresis. Separation of proteins may be by isoelectric point (pI), molecular weight, electric charge, or a combination of these factors. The nature of the separation depends on the treatment of the sample and the nature of the gel.
By far the most common type of gel electrophoresis employs polyacrylamide gels and buffers loaded with sodium dodecyl sulfate (SDS). SDS-PAGE (SDS polyacrylamide gel electrophoresis) maintains polypeptides in a denatured state once they have been treated with strong reducing agents to remove secondary and tertiary structure (e.g. S-S disulfide bonds to SH and SH) and thus allows separation of proteins by their molecular weight. Sampled proteins become covered in the negatively charged SDS and move to the positively charged electrode through the acrylamide mesh of the gel. Smaller proteins migrate faster through this mesh and the proteins are thus separated according to size (usually measured in kilo Daltons, kD). The concentration of acrylamide determines the resolution of the gel - the greater the acrylamide concentration the better the resolution of lower molecular weight proteins. The lower the acrylamide concentration the better the resolution of higher molecular weight proteins. Proteins travel only in one dimension along the gel for most blots.
Samples are loaded into wells in the gel. One lane is usually reserved for a marker or ladder, a commercially available mixture of proteins having defined molecular weights, typically stained so as to form visible, coloured bands. An example of a ladder is the GE Full Range Molecular weight ladder (Figure 1). When voltage is applied along the gel, proteins migrate into it at different speeds. These different rates of advancement (different electrophoretic mobilities) separate into bands within each lane.
It is also possible to use a two-dimensional (2-D) gel which spreads the proteins from a single sample out in two dimensions. Proteins are separated according to isoelectric point (pH at which they have neutral net charge) in the first dimension, and according to their molecular weight in the second dimension.
3. Transfer
In order to make the proteins accessible to antibody detection, they are moved from within the gel onto a membrane made of nitrocellulose or polyvinylidene fluoride (PVDF). The membrane is placed on top of the gel, and a stack of tissue papers placed on top of that. The entire stack is placed in a buffer solution which moves up the paper by capillary action, bringing the proteins with it. Another method for transferring the proteins is called electroblotting and uses an electric current to pull proteins from the gel into the PVDF or nitrocellulose membrane. The proteins move from within the gel onto the membrane while maintaining the organization they had within the gel. As a result of this "blotting" process, the proteins are exposed on a thin surface layer for detection (see below). Both varieties of membrane are chosen for their non-specific protein binding properties (i.e. binds all proteins equally well). Protein binding is based upon hydrophobic interactions, as well as charged interactions between the membrane and protein. Nitrocellulose membranes are cheaper than PVDF, but are far more fragile and do not stand up well to repeated probings.
The uniformity and overall effectiveness of transfer of protein from the gel to the membrane can be checked by staining the membrane with Coomassie or Ponceau S dyes. Coomassie is the more sensitive of the two, although Ponceau S's water solubility makes it easier to subsequently destain and probe the membrane as described below.
4. Blocking
Since the membrane has been chosen for its ability to bind protein, and both antibodies and the target are proteins, steps must be taken to prevent interactions between the membrane and the antibody used for detection of the target protein. Blocking of non-specific binding is achieved by placing the membrane in a dilute solution of protein - typically Bovine serum albumin (BSA) or non-fat dry milk (both are inexpensive), with a minute percentage of detergent such as Tween 20. The protein in the dilute solution attaches to the membrane in all places where the target proteins have not attached. Thus, when the antibody is added, there is no room on the membrane for it to attach other than on the binding sites of the specific target protein. This reduces "noise" in the final product of the Western blot, leading to clearer results, and eliminates false positives.
5. Detection
During the detection process the membrane is "probed" for the protein of interest with a modified antibody which is linked to a reporter enzyme, which when exposed to an appropriate substrate drives a colourimetric reaction and produces a colour. For a variety of reasons, this traditionally takes place in a two-step process, although there are now one-step detection methods available for certain applications.
5.1 Two step
Primary antibody
Antibodies are generated when a host species or immune cell culture is exposed to the protein of interest (or a part thereof). Normally, this is part of the immune response, whereas here they are harvested and used as sensitive and specific detection tools that bind the protein directly.
After blocking, a dilute solution of primary antibody (generally between 0.5 and 5 micrograms/ml) is incubated with the membrane under gentle agitation. Typically, the solution is composed of buffered saline solution with a small percentage of detergent, and sometimes with powdered milk or BSA. The antibody solution and the membrane can be sealed and incubated together for anywhere from 30 minutes to overnight. It can also be incubated at different temperatures, with warmer temperatures being associated with more binding, both specific (to the target protein, the "signal") and non-specific ("noise").
Secondary antibody
After rinsing the membrane to remove unbound primary antibody, the membrane is exposed to another antibody, directed at a species-specific portion of the primary antibody. This is known as a secondary antibody, and due to its targeting properties, tends to be referred to as "anti-mouse," "anti-goat," etc. Antibodies come from animal sources (or animal sourced hybridoma cultures); an anti-mouse secondary will bind to just about any mouse-sourced primary antibody. This allows some cost savings by allowing an entire lab to share a single source of mass-produced antibody, and provides far more consistent results. The secondary antibody is usually linked to biotin or to a reporter enzyme such as alkaline phosphatase or horseradish peroxidase. This means that several secondary antibodies will bind to one primary antibody and enhances the signal.
Most commonly, a horseradish peroxidase-linked secondary is used in conjunction with a chemiluminescent agent, and the reaction product produces luminescence in proportion to the amount of protein. A sensitive sheet of photographic film is placed against the membrane, and exposure to the light from the reaction creates an image of the antibodies bound to the blot.
As with the ELISPOT and ELISA procedures, the enzyme can be provided with a substrate molecule that will be converted by the enzyme to a colored reaction product that will be visible on the membrane (see the figure below with blue bands).
A third alternative is to use a radioactive label rather than an enzyme coupled to the secondary antibody, such as labeling an antibody-binding protein like Staphylococcus Protein A with a radioactive isotope of iodine. Since other methods are safer, quicker and cheaper this method is now rarely used.
5.2 One step
Historically, the probing process was performed in two steps because of the relative ease of producing primary and secondary antibodies in separate processes. This gives researchers and corporations huge advantages in terms of flexibility, and adds an amplification step to the detection process. Given the advent of high-throughput protein analysis and lower limits of detection, however, there has been interest in developing one-step probing systems that would allow the process to occur faster and with less consumables. This requires a probe antibody which both recognizes the protein of interest and contains a detectable label, probes which are often available for known protein tags. The primary probe is incubated with the membrane in a manner similar to that for the primary antibody in a two-step process, and then is ready for direct detection after a series of wash steps.
6. Analysis
After the unbound probes are washed away, the Western blot is ready for detection of the probes that are labeled and bound to the protein of interest. In practical terms, not all Westerns reveal protein only at one band in a membrane. Size approximations are taken by comparing the stained bands to that of the marker or ladder loaded during electrophoresis. The process is repeated for a structural protein, such as actin or tubulin, that should not change between samples. The amount of target protein is indexed to the structural protein to control between groups. This practice ensures correction for the amount of total protein on the membrane in case of errors or incomplete transfers.
6.1 Colorimetric detection
The colorimetric detection method depends on incubation of the Western blot with a substrate that reacts with the reporter enzyme (such as peroxidase) that is bound to the secondary antibody. This converts the soluble dye into an insoluble form of a different color that precipitates next to the enzyme and thereby stains the membrane. Development of the blot is then stopped by washing away the soluble dye. Protein levels are evaluated through densitometry (how intense the stain is) or spectrophotometry.
The western blot (alternately, immunoblot) is a method of detecting specific proteins in a given sample of tissue homogenate or extract. It uses gel electrophoresis to separate native or denatured proteins by the length of the polypeptide (denaturing conditions) or by the 3-D structure of the protein (native/ non-denaturing conditions). The proteins are then transferred to a membrane (typically nitrocellulose or PVDF), where they are probed (detected) using antibodies specific to the target protein. There are now many reagent companies that specialize in providing antibodies (both monoclonal and polyclonal antibodies) against many thousands of different proteins. Commercial antibodies can be expensive, though the unbound antibody can be reused between experiments. This method is used in the fields of molecular biology, biochemistry, immunogenetics and other molecular biology disciplines.
Other related techniques include using antibodies to detect proteins in tissues and cells by immunostaining and enzyme-linked immunosorbent assay (ELISA).
• The method originated from the laboratory of George Stark at Stanford. The name western blot was given to the technique by W. Neal Burnette[1] and is a play on the name Southern blot, a technique for DNA detection developed earlier by Edwin Southern. Detection of RNA is termed northern blotting.
Steps in a Western blot
1. Tissue preparation
Samples may be taken from whole tissue or from cell culture. In most cases, solid tissues are first broken down mechanically using a blender (for larger sample volumes), using a homogenizer (smaller volumes), or by sonication. Cells may also be broken open by one of the above mechanical methods. However, it should be noted that bacteria, virus or environmental samples can be the source of protein and thus Western blotting is not restricted to cellular studies only.
Assorted detergents, salts, and buffers may be employed to encourage lysis of cells and to solubilize proteins. Protease and phosphatase inhibitors are often added to prevent the digestion of the sample by its own enzymes.
A combination of biochemical and mechanical techniques – including various types of filtration and centrifugation – can be used to separate different cell compartments and organelles.
2. Gel electrophoresis
The proteins of the sample are separated using gel electrophoresis. Separation of proteins may be by isoelectric point (pI), molecular weight, electric charge, or a combination of these factors. The nature of the separation depends on the treatment of the sample and the nature of the gel.
By far the most common type of gel electrophoresis employs polyacrylamide gels and buffers loaded with sodium dodecyl sulfate (SDS). SDS-PAGE (SDS polyacrylamide gel electrophoresis) maintains polypeptides in a denatured state once they have been treated with strong reducing agents to remove secondary and tertiary structure (e.g. S-S disulfide bonds to SH and SH) and thus allows separation of proteins by their molecular weight. Sampled proteins become covered in the negatively charged SDS and move to the positively charged electrode through the acrylamide mesh of the gel. Smaller proteins migrate faster through this mesh and the proteins are thus separated according to size (usually measured in kilo Daltons, kD). The concentration of acrylamide determines the resolution of the gel - the greater the acrylamide concentration the better the resolution of lower molecular weight proteins. The lower the acrylamide concentration the better the resolution of higher molecular weight proteins. Proteins travel only in one dimension along the gel for most blots.
Samples are loaded into wells in the gel. One lane is usually reserved for a marker or ladder, a commercially available mixture of proteins having defined molecular weights, typically stained so as to form visible, coloured bands. An example of a ladder is the GE Full Range Molecular weight ladder (Figure 1). When voltage is applied along the gel, proteins migrate into it at different speeds. These different rates of advancement (different electrophoretic mobilities) separate into bands within each lane.
It is also possible to use a two-dimensional (2-D) gel which spreads the proteins from a single sample out in two dimensions. Proteins are separated according to isoelectric point (pH at which they have neutral net charge) in the first dimension, and according to their molecular weight in the second dimension.
3. Transfer
In order to make the proteins accessible to antibody detection, they are moved from within the gel onto a membrane made of nitrocellulose or polyvinylidene fluoride (PVDF). The membrane is placed on top of the gel, and a stack of tissue papers placed on top of that. The entire stack is placed in a buffer solution which moves up the paper by capillary action, bringing the proteins with it. Another method for transferring the proteins is called electroblotting and uses an electric current to pull proteins from the gel into the PVDF or nitrocellulose membrane. The proteins move from within the gel onto the membrane while maintaining the organization they had within the gel. As a result of this "blotting" process, the proteins are exposed on a thin surface layer for detection (see below). Both varieties of membrane are chosen for their non-specific protein binding properties (i.e. binds all proteins equally well). Protein binding is based upon hydrophobic interactions, as well as charged interactions between the membrane and protein. Nitrocellulose membranes are cheaper than PVDF, but are far more fragile and do not stand up well to repeated probings.
The uniformity and overall effectiveness of transfer of protein from the gel to the membrane can be checked by staining the membrane with Coomassie or Ponceau S dyes. Coomassie is the more sensitive of the two, although Ponceau S's water solubility makes it easier to subsequently destain and probe the membrane as described below.
4. Blocking
Since the membrane has been chosen for its ability to bind protein, and both antibodies and the target are proteins, steps must be taken to prevent interactions between the membrane and the antibody used for detection of the target protein. Blocking of non-specific binding is achieved by placing the membrane in a dilute solution of protein - typically Bovine serum albumin (BSA) or non-fat dry milk (both are inexpensive), with a minute percentage of detergent such as Tween 20. The protein in the dilute solution attaches to the membrane in all places where the target proteins have not attached. Thus, when the antibody is added, there is no room on the membrane for it to attach other than on the binding sites of the specific target protein. This reduces "noise" in the final product of the Western blot, leading to clearer results, and eliminates false positives.
5. Detection
During the detection process the membrane is "probed" for the protein of interest with a modified antibody which is linked to a reporter enzyme, which when exposed to an appropriate substrate drives a colourimetric reaction and produces a colour. For a variety of reasons, this traditionally takes place in a two-step process, although there are now one-step detection methods available for certain applications.
5.1 Two step
Primary antibody
Antibodies are generated when a host species or immune cell culture is exposed to the protein of interest (or a part thereof). Normally, this is part of the immune response, whereas here they are harvested and used as sensitive and specific detection tools that bind the protein directly.
After blocking, a dilute solution of primary antibody (generally between 0.5 and 5 micrograms/ml) is incubated with the membrane under gentle agitation. Typically, the solution is composed of buffered saline solution with a small percentage of detergent, and sometimes with powdered milk or BSA. The antibody solution and the membrane can be sealed and incubated together for anywhere from 30 minutes to overnight. It can also be incubated at different temperatures, with warmer temperatures being associated with more binding, both specific (to the target protein, the "signal") and non-specific ("noise").
Secondary antibody
After rinsing the membrane to remove unbound primary antibody, the membrane is exposed to another antibody, directed at a species-specific portion of the primary antibody. This is known as a secondary antibody, and due to its targeting properties, tends to be referred to as "anti-mouse," "anti-goat," etc. Antibodies come from animal sources (or animal sourced hybridoma cultures); an anti-mouse secondary will bind to just about any mouse-sourced primary antibody. This allows some cost savings by allowing an entire lab to share a single source of mass-produced antibody, and provides far more consistent results. The secondary antibody is usually linked to biotin or to a reporter enzyme such as alkaline phosphatase or horseradish peroxidase. This means that several secondary antibodies will bind to one primary antibody and enhances the signal.
Most commonly, a horseradish peroxidase-linked secondary is used in conjunction with a chemiluminescent agent, and the reaction product produces luminescence in proportion to the amount of protein. A sensitive sheet of photographic film is placed against the membrane, and exposure to the light from the reaction creates an image of the antibodies bound to the blot.
As with the ELISPOT and ELISA procedures, the enzyme can be provided with a substrate molecule that will be converted by the enzyme to a colored reaction product that will be visible on the membrane (see the figure below with blue bands).
A third alternative is to use a radioactive label rather than an enzyme coupled to the secondary antibody, such as labeling an antibody-binding protein like Staphylococcus Protein A with a radioactive isotope of iodine. Since other methods are safer, quicker and cheaper this method is now rarely used.
5.2 One step
Historically, the probing process was performed in two steps because of the relative ease of producing primary and secondary antibodies in separate processes. This gives researchers and corporations huge advantages in terms of flexibility, and adds an amplification step to the detection process. Given the advent of high-throughput protein analysis and lower limits of detection, however, there has been interest in developing one-step probing systems that would allow the process to occur faster and with less consumables. This requires a probe antibody which both recognizes the protein of interest and contains a detectable label, probes which are often available for known protein tags. The primary probe is incubated with the membrane in a manner similar to that for the primary antibody in a two-step process, and then is ready for direct detection after a series of wash steps.
6. Analysis
After the unbound probes are washed away, the Western blot is ready for detection of the probes that are labeled and bound to the protein of interest. In practical terms, not all Westerns reveal protein only at one band in a membrane. Size approximations are taken by comparing the stained bands to that of the marker or ladder loaded during electrophoresis. The process is repeated for a structural protein, such as actin or tubulin, that should not change between samples. The amount of target protein is indexed to the structural protein to control between groups. This practice ensures correction for the amount of total protein on the membrane in case of errors or incomplete transfers.
6.1 Colorimetric detection
The colorimetric detection method depends on incubation of the Western blot with a substrate that reacts with the reporter enzyme (such as peroxidase) that is bound to the secondary antibody. This converts the soluble dye into an insoluble form of a different color that precipitates next to the enzyme and thereby stains the membrane. Development of the blot is then stopped by washing away the soluble dye. Protein levels are evaluated through densitometry (how intense the stain is) or spectrophotometry.
6.2 Chemiluminescence
Chemiluminescent detection methods depend on incubation of the Western blot with a substrate that will luminesce when exposed to the reporter on the secondary antibody. The light is then detected by photographic film, and more recently by CCD cameras which captures a digital image of the Western blot. The image is analysed by densitometry, which evaluates the relative amount of protein staining and quantifies the results in terms of optical density. Newer software allows further data analysis such as molecular weight analysis if appropriate standards are used. So-called "enhanced chemiluminescent" (ECL) detection is considered to be among the most sensitive detection methods for blotting analysis.
6.3 Radioactive detection
Radioactive labels do not require enzyme substrates, but rather allow the placement of medical X-ray film directly against the Western blot which develops as it is exposed to the label and creates dark regions which correspond to the protein bands of interest (see image to the right). The importance of radioactive detections methods is declining[citation needed], because it is very expensive, health and safety risks are high and ECL provides a useful alternative.
6.4 Fluorescent detection
The fluorescently labeled probe is excited by light and the emission of the excitation is then detected by a photosensor such as CCD camera equipped with appropriate emission filters which captures a digital image of the Western blot and allows further data analysis such as molecular weight analysis and a quantitative Western blot analysis. Fluorescence is considered to be among the most sensitive detection methods for blotting analysis.
6.5 Secondary probing
One major difference between nitrocellulose and PVDF membranes relates to the ability of each to support "stripping" antibodies off and reusing the membrane for subsequent antibody probes. While there are well-established protocols available for stripping nitrocellulose membranes, the sturdier PVDF allows for easier stripping, and for more reuse before background noise limits experiments. Another difference is that, unlike nitrocellulose, PVDF must be soaked in 95% ethanol, isopropanol or methanol before use. PVDF membranes also tend to be thicker and more resistant to damage during use.
Chemiluminescent detection methods depend on incubation of the Western blot with a substrate that will luminesce when exposed to the reporter on the secondary antibody. The light is then detected by photographic film, and more recently by CCD cameras which captures a digital image of the Western blot. The image is analysed by densitometry, which evaluates the relative amount of protein staining and quantifies the results in terms of optical density. Newer software allows further data analysis such as molecular weight analysis if appropriate standards are used. So-called "enhanced chemiluminescent" (ECL) detection is considered to be among the most sensitive detection methods for blotting analysis.
6.3 Radioactive detection
Radioactive labels do not require enzyme substrates, but rather allow the placement of medical X-ray film directly against the Western blot which develops as it is exposed to the label and creates dark regions which correspond to the protein bands of interest (see image to the right). The importance of radioactive detections methods is declining[citation needed], because it is very expensive, health and safety risks are high and ECL provides a useful alternative.
6.4 Fluorescent detection
The fluorescently labeled probe is excited by light and the emission of the excitation is then detected by a photosensor such as CCD camera equipped with appropriate emission filters which captures a digital image of the Western blot and allows further data analysis such as molecular weight analysis and a quantitative Western blot analysis. Fluorescence is considered to be among the most sensitive detection methods for blotting analysis.
6.5 Secondary probing
One major difference between nitrocellulose and PVDF membranes relates to the ability of each to support "stripping" antibodies off and reusing the membrane for subsequent antibody probes. While there are well-established protocols available for stripping nitrocellulose membranes, the sturdier PVDF allows for easier stripping, and for more reuse before background noise limits experiments. Another difference is that, unlike nitrocellulose, PVDF must be soaked in 95% ethanol, isopropanol or methanol before use. PVDF membranes also tend to be thicker and more resistant to damage during use.
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