Protein Purification

Protein Purification

An overview of protein isolation:

Protein isolation and purification is one the tedious job that takes a very long time if one can’t choose an efficient and suitable technique. There are numerous different types of proteins in a cell and to study about each protein is very difficult one.

Why do it?

Living organisms such as human beings and other higher eukaryotes are complex machines made up of many interacting systems. Protein constitute the majority of the working parts of these systems and these diverse reasons for isolating proteins are

• To gain insight. As with any mechanism, to study the way in which a living system works it is necessary to dismantle the machine and to isolate the component parts so that they may be studied, separately and in their interaction with other parts. The knowledge that is gained in this way may be put to practical use, for example, in the design of medicines, diagnostics, pesticides, or industrial processes. Many proteins may themselves be used as “medicines” to make up for losses or inadequate synthesis. Examples are hormones, such as insulin, which is used in the therapy of diabetes, and blood fractions, such as the so-called Factor VIII, which is used in the therapy of haemophilia. Other proteins may be used in medical diagnostics, an example being the enzymes glucose oxidase and peroxidase, which are used to measure glucose levels in biological fluids, such as blood and urine.

• For use in Industry. Many enzymes are used in industrial processes, especially where the materials being processed are of biological origin. In every case a pure protein is desirable as impurities may either be misleading, dangerous or unproductive, respectively. Protein isolation is, therefore, a very common, almost central, procedure in biochemistry.

• For use in Medicine.

The Background factors:

Properties of proteins that influence the methods used in their study

It must be appreciated that proteins have two properties which determine the overall approach to protein isolation and make this different from the approach used to isolate small natural molecules.

1. Proteins are labile

As molecules go, proteins are relatively large and delicate and their shape is easily changed, a process called denaturation, which leads to loss of their biological activity. This means that only mild procedures can be used and techniques such as boiling and distillation, which are commonly used in organic chemistry, are thus verboten.

2. Proteins are similar to one another.

All proteins are composed of essentially the same amino acids and differ only in the proportions and sequence of their amino acids, and in the 3-D folding of the amino acid chains. Consequently processes with a high discriminating potential are needed to separate proteins. The combined requirement for delicateness yet high discrimination means that, in a word, protein separation techniques have to be very subtle. Subtlety, in fact, is required of both techniques and of experimenters in biochemistry.

Protein fractionation techniques:

Figure 1: Flowchart of Protein purification steps

Protein Extraction

Since proteins are only synthesized by living systems, a protein must be isolated from biological materials or from bioproducts. The main objective of protein isolation is to separate the protein of interest from all non protein and all other unwanted proteins which occur in the same sample. As a general principle, one should aim to achieve the isolation of a protein

1. In as few steps as possible to minimize the lost in each step.

2. In as short a time as possible to prevent protein from denaturation.

Therefore, it is very important to optimize the protocol for specific protein to isolate and purify.

Where to start?

Before starting the extraction and purification process, one should know whether the proteins are extracellular and intracellular. On the basis of these and the background factors of protein, we can correctly apply the best method for extraction and purification which not only reduce the time of extraction but also save energy and money for purification.

For Extracellular proteins:

There is no need to cell disintegration for extracellular proteins. The cell culture is simply filtered and centrifuged. Then the supernatant portion of centrifuge contains extracellular proteins. E.g. Proteases, Antibiotics (proteins).

For Intracellular proteins:

In order to isolate the intracellular proteins, the cells are disrupted with using suitable buffer. As many of proteins are very sensitive in nature the choice of disruption technique should be done accurately. The appropriate method of cell disruption may reduce many steps in protein purification.

Preliminary Fractionation:

Frationation of homogenate: Once a homogenate has been prepared, cytosolic water-soluble proteins can be obtained by the removal of particulate matter, usually through centrifugation. In addition it is often desirable to isolate a specific particulate subsellular fraction if the desired protein is known to reside there. For instance of one wishes to purify a transcription factor localized to the nucleus one would first prepare nuclei and extract them. A low-speed centrifugation separates nuclei from the remainder of the homogenate. The postnuclear supernatant can then be subjected to centrifugation at intermediate speed to sediment mitochondria. Finally the postmitochondrial supernatant can be fractionated in the ultracentrifuge into a ribosomal fraction and a postribosomal supernantant which contains all soluble proteins.

Fractionation of proteins in solution

Eventually the desired protein is obtained in aqueous solution together with a host of unrelated proteins. At this moment the protein becomes amenable to a number of manipulations that can, in principle at least, separate it from all other proteins in the sample.

a. Fractionation by precipitation.

To begin the process of fractionation, it is possible to alter the composition of the buffer to force the precipitation of a portion of the proteins. This procedure takes advantage of the differential solubility of proteins under varying conditions as well as the high protein concentration of the solution which permits aggregates of protein to form over a short period of time (minutes to hours). Precipitation of one group of proteins permits their separation from other proteins that remain in solution by low speed centrifugation.

(i) "Salting out" - Ammonium Sulfate Precipitation: high salt concentrations promote precipitations. With increasing ionic strength, proteins begin to interact via hydrophobic patches on their surface, as proteins and salt compete for the residual water. Ammonium sulfate is used as the salt of choice, since it preserves protein activity and promotes precipitation at lower concentrations than other salts. (Phosphate should be better than sulfate on theoretical grounds, but is ruled out because trivalent phosphate occurs only at extremely low pH. Sodium sulfate and many potassium salts are not sufficiently soluble; many multivalent cations are toxic; also potassium phosphate at even fairly low molar concentration is too dense to allow precipitate to settle - economy of the pure salt also plays a role). One adds increasing amounts of ammonium sulfate to the extract, with intermittent centrifugation steps. These stepwise precipitations are referred to as ammonium sulfate "cuts." The amounts of solid ammonium sulfate to be added to a known volume in order to obtain the desired percentage saturation can be looked up in tables. Solid ammonium sulfate should be added slowly, while the solution is stirring to allow a uniform increase in the concentration, and in powdered form, to ensure rapid equilibration.

What are the mechanisms of protein precipitation by Ammonium Sulphate?

Polyvalent anions are more effective at salting out than univalent anions, while polyvalent cations tend to negate the effect of polyvalent anions. The best combination is therefore a polyvalent anion with a univalent cation. Anions can be arranged in a so-called “Hofmeister series”, which describes their relative effectiveness in salting out at equivalent molar concentrations2. In decreasing order of effectiveness, the series is:

citrate > sulfate > phosphate > chloride > nitrate >thiocyanate.

This series also describes a decreasing tendency for the anions to stabilise protein structure. Citrate and sulfate are thus “kosmotropes”, which tend to stabilise protein structure, while thiocyanate and nitrate are “chaotropes” which tend to destabilize protein structure. An ideal salt would, therefore, be citrate or sulfate combined with a univalent cation. Ammonium sulfate is most popular because it meets these criteria, is available in a pure form at low cost and is highly soluble, so that high solution concentrations can be attained. The sulfate ion has been viewed in a number of ways, regarding how it salts out proteins, including, ionic strength effects, kosmotropy, exclusion-crowding, dehydration, and binding to cationic sites, especially when the protein has a net positive charge (denoted ZH+)3. All of these may play a role, depending upon the salt concentration and the pHdependent charge on the protein.

Ionic strength effects. It will be noticed that the Hofmeister series goes from multivalent to univalent ions. This largely reflects the fact that the Hofmeister series is based on molarity, while ionic strength is a factor in salting out. The valency of the ion has an effect on ionic strength as can be illustrated by comparing NaCl with (NH4)2SO4.

Ionic strength is defined as:- 1/2S ci (zi)²

Where, ci = concentration of each type of ion (moles/litre)

zi= charge of each type of ion

Thus in the case of 1 M NaCl, ionic strength is 1 and for 1 M (NH4)2SO4, it is 3.

Ionic strength effects come into play at low salt concentrations M) and, as the name implies, they are not specific to ammonium sulfate. At low ionic strength, protein solubility is at its minimum at the proteinís pI. At this pH, intramolecular electrostatic forces between oppositely charged side chains are at a maximum, protein conformation is maximally tightened and protein hydration is least. On either side of the pI, titration of ionisable groups leads to a lessening of intramolecular ionic interactions. In consequence, protein structure becomes more relaxed and hydration and solubility are increased. Addition of low concentrations of salt causes a similar weakening of intramolecular ionic bonds, with similar consequences of more relaxed protein structure and greater solubility. The increase in solubility of protein upon addition of modest amounts of salt is known as “salting in”.

Kosmotropy. At concentrations above 0.2 M the sulfate ion acts as a Hofmeister kosmotrope, i.e. it stabilises protein structure, and concomitantly reduces its solubility. The effect of a kosmotrope, in stabilising protein structure, can be described by the reaction:-

Relaxed, open protein structure Û compact, tight structure

(more soluble, less stable) (less soluble, more stable)

Kosmotropes may be described as “pushing” if they act on the left of this reaction and “pulling” if they act on the right, in either case driving the reaction to the right. Sulfate can act as a pulling kosmotrope by virtue of its interaction with protein cationic sites. Consistent with this, the precipitation of proteins is usually promoted at pH values below the pI, where the protein has a maximal number of cationic sites3. Reinforcing its pulling effect is the fact that the sulfate ion is divalent, and so can bind to more than one cationic site at a time, and that it has a tetrahedral structure, with four oxygen atoms that can hydrogen bond to multiple sites on the protein. Sulfate also acts as a pushing kosmotrope by virtue of its extraordinary hydration. By virtue of its hydration, the sulfate ion can act as a dehydrating agent and, in its hydrated form, as an exclusion crowding agent. The sulfate anion has 13 or 14 water molecules in its first hydration layer and possibly more in a second layer4. Consequently, in salting out at, say, 3 M ammonium sulfate, the sulfate anion will have accreted to itself 40 to 45 M out of the total of 55 M H2O in neat water. In salting out, therefore, a large proportion of the water will be involved in hydrating the sulfate ions and increasing their effective radius. The large, hydrated, [SO4.(H2O)n]2- ions crowd and exclude the proteins, pushing them into tighter, more ordered (less soluble) conformations, with lower entropy. The preferential accretion of the water molecules to the sulfate ions excludes the proteins from a proportion of the water (the proportion increasing with the salt concentration), ultimately bringing them to their solubility limit. No other salt has the combination of properties which make ammonium sulfate so effective at salting out. Consequently, when the word salt is used in the context of salting out, it invariably means ammonium sulfate. Similarly, the term “ionic strength” is often used loosely, when what is really meant is the concentration of ammonium sulfate.

(ii) Isoelectric precipitation. Solubility of proteins depends on ionic strength and pH; usually very soluble at physiological ionic strength and pH, as cellular protein concentration can reach 40% of total mass. If the pH approaches the pI of a given protein the net charge on that protein will go to zero, and in the absence of electrostatic repulsion, weak electrostatic attractive forces may lead to aggregation and precipitation; this tendency to interact can be promoted by reducing the ionic strength or by adding organic solvent (see solvent precipitation below!). "The effects of ionic strength of the medium on the interaction between charged macroions can be summarized as follows. At very low ionic strength, the counterion atmosphere is highly expanded and diffuse, and screening is ineffective. Like-charged particles repel strongly; unlike charged particles attract one another strongly. As the ionic strength increases, the counterion atmosphere shrinks and becomes concentrated about the macroion. Screening becomes effective. - The effects of charge and ionic strength on the solubility of polyampholytes such as proteins can be explained in terms of these electrostatic interactions. Consider the behavior of the common milk protein b-lactoglobulin. The isoelectric point of this polyampholyte is about 5.3. Above or below this pH, the molecules all have either negative or positive charges and repel one another, so the protein is very soluble at either acidic or basic pH. At the isoelectric point, the net charge is zero but each molecule still carries surface patches of both positive charge and negative charge. The ionic interactions between them, together with other kinds of intermolecular interactions such as van der Waals forces, make the molecules tend to clump together and precipitate. Therefore, solubility is minimal at the isoelectric point. If the ionic strength is increased, however, the counter on atmosphere shrinks about the charged regions, and the attractive interactions between positive and negative groups are effectively screened. Hence, solubility increases, even at the isoelectric point. This effect of putting proteins into solution by increasing the salt concentration is called salting in."]

(iii) Solvent precipitation: used to be popular in the early days of protein isolation, especially in the fractionation of plasma proteins (ethanol; acetone). Mechanism: reduction of water activity (i.e. availability of water for protein solvation); most likely to occur at pI which suggests that the interaction between proteins may be similar to that in isoelectric precipitation. Order of precipitation largely determined by the size of the protein. Disadvantages: flammable (acetone); easily denatures proteins when conducted at temperatures above 0^oC. A variant of the procedure is PEG precipitation (at 20% and MW of 4000 or greater).

(iv) Selective denaturation of contaminant proteins in isolated cases can be brought about by heat, extremes of pH, and organic solvents. (examples are isolation of calmodulin from brain extracts by boiling??) but note that heat treatment may result in chemical modification of the desired protein and is therefore not recommended

Dialysis and ultrafiltration: Fractionation by means of semipermeable membranes is generally used for buffer changes but can also provide low-resolution protein fractionation. In dialysis the protein sample is enclosed in a bag consisting of a semipermeable membrane (made of cellulose) and exposed to a large volume of a desired buffer. The low-molecular weight compounds (buffering agents, salts) pass freely through the membrane pores wherease the protein is retained. This procedure lends itself to desalting steps and buffer exchanges. Ultrafiltration is similar except that the pores can be larger and therefore allow smaller proteins to pass through. In other words such membranes can be used for protein fractionation. Since dialysis speed is a function of molecular mass, ultrafiltration employs pressure to force the sample through. Membranes with various molecular weight cutoffs are available (from less than 10 to more than 100 kDa). Obviously this method is not very efficient, as only 2 fractions, a filtrate and a "retained fraction" are obtained.

Chromatographic and electrophoretic procedures

1. Chromatography - any of a number of methods in which solutes (in our case proteins) are fractionated by partitioning between a mobile or buffer and an immobile or matrix, phase. The most widely used technique involves a matrix packed in a column through which the protein sample is passed.

a. Gel filtration.

Gel filtration (also size exclusion or gel permeation chromatography) separates molecules, including proteins, on the basis of their size and shape. Large proteins do not enter the pores of the chromatographic matrix, but pass through, flowing in the interstitial space between the matrix beads; this space is also known as the void volume, V₀. For most resins, V₀ is approximately 1/3 of V_T, where V_T is the total packed volume of the resin (V_T = lr²p; where, r = radius of the column and l = length of the resin bed). If a molecule is smaller than the pores in the resin, it will diffuse into the beads. In the space outside the beads there is bulk flow only, while inside the beads solutes move by diffusion only. Therefore, large proteins confined to the interstitial space will migrate more rapidly through a resin than small molecules which diffuse into and back out of the resin and consequently are partly trapped and fall behind. Since pores vary in size, proteins of intermediate size will penetrate to varying degrees into the beads and thus are separated from each other on the basis of their size. Note:Proteins of equal mass but different shapes will differ in their apparent size. An elongated protein will appear larger, i.e. it will have a larger tumbling radius or radius of gyration and move more slowly than a spherical protein of the same mass.

A given protein will elute at a reproducible and characteristic position from a particular resin with a specific buffer. This property is usually reported as the (V_E - V₀)/(V_T - V₀) and corresponds to the R_f (= ‘relation to front’, i.e. the front of the solvent) value that is used for thin layer chromatography. A protein that elutes in the void volume will have an Rf = 0, whereas a protein that elutes at V_T will have a Rf = 1. Since, upon gel filtration, the sample is diluted by a factor of two to three or more, it is necessary to load concentrated protein samples. Under ideal conditions, the sample volume will be less than 2% of V_T. In practice, it is often difficult to concentrate the sample sufficientlly, and a sample volume that is up to 5% of the V_T can be used to give suboptimal results. The protein concentration can be as high as 50 mg/ml, and care must be taken not to disturb the packed resin with such high-density samples.

Gel filtration resins are composed of a variety of materials, including dextran, agarose, and polyacrylamide and are available in various pore sizes. Since resolution increases with bead uniformity and decreases with bead size, small homogeneous beads are preferred. Such beads tend to require higher pressures for chromatography than that achieved with conventional chromatography systems. Because the beads used in gel filtration are porous, they can be sensitive to even moderate pressure. Typically, the resins with larger pore sizes (which are more fragile) should be used at lower hydrostatic pressures than the more sturdy resins with smaller pores. Many resins are cross-linked (e.g. crosslinked dextran and agarose) to increase their strength.

b. Adsorption chromatography

In gel permeation chromatography proteins do not bind significantly to the matrix. Sorting does not occur as a result of differential binding but of differential exclusion from pores in the beads. On the other hand many sorbents are available that bind proteins with different affinities. At equilibrium the behavior of a specific protein can be described by a partition coefficient a, the fraction of the protein that is adsorbed. a therefore can have values between 0 (no binding whatever) and 1.0 (complete adsorption without any desorption). Differences in partition coefficients can be utilized for fractionation by allowing proteins to choose between a stationary, adsorbed and a mobile unadsorbed phase. In this case the completely unadsorbed protein will move with the buffer front; its mobility compared to the buffer front, R_f, will be unity. In contrast, the completely absorbed protein will not move at all, and R_f will be 0. In other words: a + R_f = 1.0

Binding of protein to the matrix is most simply achieved by mixing them. Such socalled batch adsorption is especially popular with hydroxyapatite, but can also be performed with ion exchange resins. One of the great advantages of adsorptive methods is the ability to use high volumes of sample, obviating concentration steps. This saves time and generally leads to higher overall yields in protein preparations. For example, one can load ion exchange resins in batch mode by stirring a large-volume sample with the resin. Subsequently, the resin is poured into a column for gradient elution, or batch eluted on a sintered glass funnel. Thus, one can move rapidly through large volume steps at the early stages of the preparation.

(i) Ion exchange chromatography: In this type of chromatography, the charged groups on the surface of a protein bind to an insoluble matrix with opposite charge. More precisely, the ionized protein displaces the counterions (e.g., chloride or sodium) of the matrix functional group, and will itself be displaced by an increasing concentration of ions in the elution buffer. Alternately, a pH gradient may be employed so that the net charge on the adsorbed protein decreases. Under specific starting conditions of buffer, pH, and ionic strength, the net charge on the protein of interest can be manipulated to interact with the matrix. Thus, the most important parameters to consider in an ion exchange separation are the choice of ion exchange matrix and the initial conditions.

The Ion Exchange Matrix: Ion exchangers consist of an insoluble containing charged groups. Anion exchangers contain basic groups that are positively charged below their pK's (e.g. amino functions) and cation exchangers contain acidic groups that are negatively charged above their pK's (e.g. carboxylate). For example, carboxymethyl is a weakly acidic cationic exchanger, while sulfopropyl is a strong cation exchanger, since the pK for the acidic proton is lower for sulfate than for acetate, i.e. the sulfopropyl function, i.e. the sulfopropyl function remains chraged over a greater part of the pH range than the carboxymethyl. A cationic protein will bind to both types of cation exchangers, but it will usually require higher concentration of counter ions to desorb the protein from the "stronger" ion exchanger, sulfopropyl. Similarly, DEAE is a weakly basic anion exchanger, while the quaternary ammonium ions are strong anion exchangers. A special case of ion exchange chromatography is metal chelation chromatography. Here resins contain carboxylate functions in close proximity which are capable of making coordination complexes with metal ions such as Ni²⁺. The metal cations are then capable of interacting strongly with strings of imidazols, i.e. histidine residues in proteins. The strong coordination bond can be broken by an excess of free imidazol. This type of chromatography is especially useful for the isolation of proteins under denaturing conditions, provided a string of histidines is present. Such a His tag is easily engineered into fusion proteins.

(ii) Hydrophobic Interaction Chromatography: In hydrophobic interaction chromatography (HIC) advantage is taken of hydrophobic patches on the protein surface that interact with non-polar materials under non-denaturing conditions. As in ammonium sulfate fractionation, high ionic strength promotes interactions between surface hydrophobic patches on the proteins. In HIC one raises the ionic strength to just below the point of precipitation in the presence of an immobilized hydrophobic matrix such as phenyl or octyl agarose. Under these conditions, proteins can be made to bind rather than to precipitate (note that plain agarose or Sephadex will do, as hydrophobic interactions with the polysaccharide matrix in the absence of alkyl or aryl substituents are possible). A descending salt gradient then causes desorption of proteins in the order of hydrophobic affinity between matrix and protein surface. At very low ionic strength, there may still be proteins retained by the column, and these must be removed by the addition of nonpolar or chaotropic components to the matrix. Solvent modifiers added to cause desorption of strongly bound proteins can lead to denaturation of proteins. Under those conditions, HIC becomes similar to reverse phase systems using organic solvents (Notice again the similarity to protein precipitation methods, this time solvent precipitation, and to reverse phase HPLC, which is widely used in the fracionation of peptides. Matrix:Immobilized aromatic and aliphatic compounds are used such as phenyl- and octylagarose. Also stress that HIC is possible even with unsubstituted sepharose. Loading Conditions:Protein solutions should be supplemented with a salt to promote HIC. Ammonium sulfate is a popular salt for HIC, since concentrations of 1 M (ca 25% saturation) are usually sufficient for initial conditions. Sodium or potassium chloride can be used, but often concentrations of 2 M or higher are needed to force interactions with the HIC column.

(iii) Hydroxyapatite chromatography: Chromatography of proteins on hudroxyapatite (HTP - microcrystalline precipitates of calcium phosphate Ca₅(PO₄)₃OH) which affords fractionations that often are not attainable with any other method (e.g. separation of isozymes or of antibodies that only differ in their light chains etc.). Depends on specific interaction with calcium and phosphate; elution is accomplished with an ascending gradient of phosphate. This is sometimes convenient as fairly high sodium chloride concentrations that are frequently encountered in an ion exchange column eluate do not interfere with adsorption of the sample onto the HTP column.

Monitoring the Purification Process

After each step in purification process, the protein sample should be estimated and its purity should be monitored. Through this process we can know the lost of protein and its purity level. It also helps in further purifying the protein. This monitoring process detects the efficient purification level to stop the process. There are many methods of protein purification monitoring and estimation.

For Protein Determination:

1. UVMethod:

Quantitation of the amount of protein in a solution is possible in a simple spectrom­eter. Absorption of radiation in the UV by proteins depends on the Tyr and Trp content (and to a very small extent on the amount of Phe and disulfide bonds). There­fore the A₂₈₀ varies greatly between different proteins (for a 1 mg/mL solution, from 0 up to 4 [for some tyrosine-rich wool proteins], although most values are in the range 0.5–1.5. The advantages of this method are that it is simple, and the sample is recoverable. The method has some disadvantages, including interference from other chromophores, and the specific absorption value for a given protein must be deter­mined. The extinction of nucleic acid in the 280-nm region may be as much as 10 times that of protein at their same wavelength, and hence, a few percent of nucleic acid can greatly influence the absorption.

2. Biuret Method:

In alkaline solution, proteins reduce cupric (Cu2+) ions to cuprous (Cu1+) ions which react with peptide bonds to give a blue colored complex. This reaction is called the biuret reaction and is named after the compound biuret, which is the simplest compound that yields the characteristic color. Because the reaction is with peptide bonds, there is little variation in the color intensity given by different proteins. The biuret method can be used for the measurement of protein concentration in the presence of polyethylene glycol, a common protein precipitant. A disadvantage of the biuret method is that it is relatively insensitive, so that large amounts of protein are required for the assay. A more sensitive variant of the method, the micro-biuret assay, has been devisedí, which overcomes this limitation to some extent. Another limitation is that amino buffers, such as Tris, which are commonly used in the pH range ca. 8-10, can interfere with the reaction.

3. The Lowry Method:

The most accurate method of determining protein concentration is probably acid hydrolysis followed by amino acid analysis. Most other methods are sensitive to the amino acid composition of the protein, and absolute concentrations cannot be obtained. The procedure of Lowry et al. is no exception, but its sensitivity is moderately constant from protein to protein, and it has been so widely used that Lowry protein estimations are a completely acceptable alternative to a rigorous absolute determina­tion in almost all circumstances in which protein mixtures or crude extracts are involved.

The method is based on both the Biuret reaction, in which the peptide bonds of proteins react with copper under alkaline conditions to produce Cu⁺, which reacts with the Folin reagent, and the Folin–Ciocalteau reaction, which is poorly understood but in essence phosphomolybdotungstate is reduced to heteropolymolybdenum blue by the copper-catalyzed oxidation of aromatic amino acids. The reactions result in a strong blue color, which depends partly on the tyrosine and tryptophan content. The method is sensitive down to about 0.01 mg of protein/mL, and is best used on solutions with concentrations in the range 0.01–1.0 mg/mL of protein.

4. The Bicinchonic Acid Method:

The bicinchoninic acid (BCA) assay, first described by Smith et al. is similar to the Lowry assay, since it also depends on the conversion of Cu²⁺ to Cu⁺ under alkaline conditions. The Cu⁺ is then detected by reaction with BCA. The two assays are of similar sensitivity, but since BCA is stable under alkali conditions, this assay has the advantage that it can be carried out as a one-step process compared to the two steps needed in the Lowry assay. The reaction results in the development of an intense purple color with an absorbance maximum at 562 nm. Since the production of Cu⁺ in this assay is a function of protein concentration and incubation time, the protein content of unknown samples may be determined spectrophotometrically by compari­son with known protein standards. A further advantage of the BCA assay is that it is generally more tolerant to the presence of compounds that interfere with the Lowry assay. In particular it is not affected by a range of detergents and denaturing agents such as urea and guanidinium chloride, although it is more sensitive to the presence of reducing sugars. The sensitivity for standard assay is 0.1–1.0 mg protein/mL and that of microassay is 0.5–10 µg protein/mL

5. The Bradford Method (Dye Binding Method):

A rapid and accurate method for the estimation of protein concentration is essential in many fields of protein study. An assay originally described by Bradford has become the preferred method for quantifying protein in many laboratories. This tech­nique is simpler, faster, and more sensitive than the Lowry method. Moreover, when compared with the Lowry method, it is subject to less interference by common rea­gents and nonprotein components of biological samples.

The Bradford assay relies on the binding of the dye Coomassie Blue G250 to pro­tein. Detailed studies indicate that the free dye can exist in four different ionic forms for which the pK_a values are 1.15, 1.82, and 12.4. Of the three charged forms of the dye that predominate in the acidic assay reagent solution, the more cationic red and green forms have absorbance maxima at 470 nm and 650 nm, respectively. In contrast, the more anionic blue form of the dye, which binds to protein, has an absorbance maxi­mum at 590 nm. Thus, the quantity of protein can be estimated by determining the amount of dye in the blue ionic form. This is usually achieved by measuring the absor­bance of the solution at 595 nm.

The dye appears to bind most readily to arginyl and lysyl residues of proteins. This specificity can lead to variation in the response of the assay to different proteins, which is the main drawback of the method. The original Bradford assay shows large variation in response between different proteins. Several modifica­tions to the method have been developed to overcome this problem. How­ever, these changes generally result in a less robust assay that is often more susceptible to interference by other chemicals. Consequently, the original method devised by Bradford remains the most convenient and widely used formulation. The standard assay, which is suitable for measuring between 10 and 100 µg of protein, and the microassay, which detects between 1 and 10 µg of protein are commonly used. The latter, although more sensitive, is also more prone to interference from other com­pounds because of the greater amount of sample relative to dye reagent in this form of the assay.

For Monitoring Purification

1. Spectrophotometric Method:

A single peak on spectrophotometric analysis shows the purity of protein sample.

2. Enzymatic Method:

For enzymes, the increase in purity will increase the activity of protein. The enzymatic activity can be determined by using specific substrate at specific temperature and pH optimal for the enzyme.

3. Electrophoresis Method:

A single and pure protein reveals a single distinct band upon electrophoresis. Therefore, it is also a best method for monitoring the protein purification.