My Scientific Gallery; Upendra Thapa Shrestha

pH Value, indicators, charge relations

2012-02-09T06:34:00.000-08:00

pH Value, indicators, charge relations

pH Value:

The pH value is the negative logarithm to the base 10 of the hydrogen ion concentration.

Derivation:

In pure water, the concentration of hydrogen ions [H₃O⁺] is equal to the concentration of hydroxyl ions [OH^–], viz

[H₃O⁺] = [OH^-] = 10^-7 mol/L

At this equilibrium, the water has a neutral reaction. In practice, the minus sign is dropped, 10^-7 = pH 7.

According to the above equation, if the hydrogen ion concentration in an aqueous system is increased, the hydroxyl ion concentration becomes less, and the system has an acid reaction.

If the hydroxyl ion concentration is increased, the hydrogen ion concentration decreases accordingly, and the solution has an alkaline reaction.

pH = 1 ←	pH = 7	→ pH = 14
acid	neutral	alkaline

Determination of pH value:

1. Potentiometric (electrometric)

With a hydrogen electrode or glass electrode (measuring range pH 0 – pH 14)

2. Colorimetric

With indicators or indicator papers.

Common indicators

Indicator	Effective pH range	Color change	Concentration of solution
Thymol Blue	1.2 – 2.8	red – yellow	0.1% in 20% alcohol
Dimethyl Yellow	2.9 – 4.0	red – yellow	0.1% in 90% alcohol
Bromophenol Blue	3.0 – 4.6	yellow – blue	0.1% in 20% alcohol
Congo Red	3.0 – 5.0	blue – red	1% in water
Methyl Orange	3.2 – 4.4	red – yellow	0.1% in water
Bromocresol Green	3.8 – 5.4	yellow – blue	0.1% in 50% alcohol
Methyl Red	4.8 – 6.0	red – yellow	0.2 % in 90% alcohol
Litmus	5.0 – 8.0	red – blue	0.3 % in 90% alcohol
Bromocresol Purple	5.2 – 6.8	yellow – purple red	0.04 % in 90% alcohol
Bromothymol Blue	6.0 – 7.6	yellow – blue	0.1% in 20% alcohol
Neutral Red	6.8 – 8.0	red – yellowish orange	0.1 in 70% alcohol
Phenol Red	6.6 – 8.0	yellow – red	0.02 % in 90% alcohol
o-Cresol Red	7.0 – 8.8	yellow – purple red	0.1% in 20% alcohol
Tropaeoline 000	7.6 – 8.9	yellowish green – pink	1% in water
Phenolphthalein	8.2 –10.0	colorless – pink	1.0 % in 50% alcohol
Thymol Blue	8.0 – 9.6	yellow – blue	0.1% in 20% alcohol
Thymolphthalein	9.4 –10.6	colorless – blue	0.1% in 90% alcohol
Alizarin Yellow R	10.1 –12.0	yellow – red	0.1% in 50% alcohol
Tropaeoline O	11.3 –13.0	yellow – orange red	0.1% in water

Example: pH colour range of bromocresol green

Color: Yellow = pH 3.5 and lower

Yellow-green = pH 4.0

Green = pH 4.5

Blue-green = pH 5.0

Blue = pH 5.5 and higher

Protein Purification

2012-02-03T06:04:00.000-08:00

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.

2D-Gel Electrophoresis and Western Blotting

2012-01-17T03:51:00.000-08:00

Two Dimensional Polyacrylamide Gel Electrophoresis

Why 2D GE?

• Oldest method for large scale protein separation (since 1975)

• Still most popular method for protein display and quantification

• Permits simultaneous detection, display, purification, identification, quantification

• Robust, increasingly reproducible, simple, cost effective, scalable & parallelizable

• Provides pI, MW, quantity

Steps in 2D GE

• Sample preparation

• Isoelectric focusing (first dimension)

• SDS-PAGE (second dimension)

• Visualization of proteins spots

• Identification of protein spots

1. Sample Preparation

• Sample preparation is key to successful 2D gel experiments

• Must break all non-covelent protein-protein, protein-DNA, protein-lipid interactions, disrupt S-S bonds

• Must prevent proteolysis, accidental phosphorylation, oxidation, cleavage, deamidation

• Must remove substances that might interfere with separation process such as salts, polar detergents (SDS), lipids, polysaccharides, nucleic acids

• Must try to keep proteins soluble during both phases of electrophoresis process

Protein Solubilization

• 8 M Urea (neutral chaotrope)

• 4% CHAPS (zwitterionic detergent)

• 2-20 mM Tris base (for buffering)

• 5-20 mM DTT (to reduce disulfides

• Carrier ampholytes or IPG buffer (up to 2% v/v) to enhance protein solubility and reduce charge-charge interactions

Other Considerations

• Further purification or separation?

• Subcellular fractionation

• Chromatographic separation

• Affinity purification

• Optimizing electrophoresis parameters

• IEF pH gradient, Acrylamide %, loading

• Limits of detection

• ng? (Coomasie stain) pg or fg? (Western)

2. Isoelectric Focusing

• Separation of basis of pI, not Mw

• Requires very high voltages (5000V)

• Requires a long period of time (10h)

• Presence of a pH gradient is critical

• Degree of resolution determined by slope of pH gradient and electric field strength

• Uses ampholytes to establish pH gradient

IEF Principles

Ampholytes vs. IPG

• Ampholytes are small, soluble, organic molecules with high buffering capacity near their pI (not characterized)

• Used to create pH gradients via user

• Gradients not stable

• Batch-to-batch variation is problematic

• An immobilized pH gradient (IPG) is made by covalently integrating acrylamido buffer molecules into acrylamide matrix at time of gel casting

• Stable gradients

• Pre-made (at factory)

• Simplified handling

Narrow-Range IPG Strips

3. SDS-PAGE

• Separation of basis of MW, not pI

• Requires modest voltages (200V)

• Requires a shorter period of time (2h)

• Presence of SDS is critical to disrupting structure and making mobility ~ 1/MW

• Degree of resolution determined by %acrylamide & electric field strength

• After IEF, the IPG strip is soaked in an equilibration buffer (50 mM Tris, pH 8.8, 2% SDS, 6M Urea, 30% glycerol, DTT, tracking dye)

• IPG strip is then placed on top of pre-cast SDS-PAGE gel and electric current applied

• This is equivalent to pipetting samples into SDS-PAGE wells (an infinite #)

4. Visualization of proteins spots

• Coomassie Stain (100 ng to 10 mg protein)

• Silver Stain (1 ng to 1 mg protein)

• Fluorescent (Sypro Ruby) Stain (1 ng & up)

5. Identification of protein spots

• Confirmed by Western, Northern or Southern

• Confirmed by amino acid composition

• Confirmed by amino acid sequencing

• Confirmed by Mw & pI

Advantages and Disadvantages of 2D GE

Western blot

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.

· method of detecting specific proteins in a given sample of tissue homogenate or extract.

· 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

· Other related techniques include using antibodies to detect proteins in tissues and cells by immunostaining and enzyme-linked immunosorbent assay (ELISA).

Steps in a Western blot

1. Tissue preparation

· Samples may be taken from whole tissue or from cell culture.

· 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.

· Sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis

· It is also possible to use a two-dimensional (2-D) gel

3. Transfer

· Tranfer the proteins from the gel onto a membrane made of nitrocellulose or polyvinylidene fluoride (PVDF by capillary action,

· 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.

· "Blotting" process, the proteins are exposed on a thin surface layer for detection

· The uniformity and overall effectiveness of transfer of protein from the gel to the membrane can be checked by staining the membrane with Coomassie

4. Blocking

· To prevent interactions between the membrane and the antibody used for detection of the target protein. Blocking of non-specific binding is achieved.

· typically Bovine serum albumin (BSA) or non-fat dry milk are used

5. Detection

Two methods are used

5.1 Two step

Primary antibody

· Antibodies are generated when a host species or immune cell culture is exposed to the protein of interest (

· The antibody solution and the membrane can be sealed and incubated together for anywhere from 30 minutes to overnight.

Secondary antibody

· Exposed to Secondary antibody, directed at a species-specific portion of the primary antibody like "anti-mouse," "anti-goat," etc.

· The secondary antibody is usually linked to biotin or to a reporter enzyme such as alkaline phosphatase or horseradish peroxidase.

· 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.

5.2 One step

· 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.

6. Analysis

Diferent methods

6.1 Colorimetric detection

· This converts the soluble dye into an insoluble form of a different color that precipitates next to the enzyme and thereby stains the membrane(such as peroxidase).

6.2 Chemiluminescence

· 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.

6.3 Radioactive detection

· 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

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

Electrophoresis

2011-12-27T06:44:00.000-08:00

Electrophoresis

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.

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 (F_ef) and is defined as:

F_ef = q x E

• This force is opposed by the frictional force,( F_f):

F_f = f V

Where f = frictional coefficient

f = 6hpr for a sphere of radius r (Stoke’s equation)

• In a constant electric field F_ef = F_f 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

An increased rate of sample diffusion.

Mixing of separated samples due to convection current.

Thermal instability, protein denaturation and loss of enzyme activity.

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.

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.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.

Centrifugation, a separating technique

2011-12-14T06:49:00.000-08:00

Centrifugation

Centrifugation is a basic separation technique. A centrifuge is a device for separating particles in an applied centrifugal field in a solution.

There are two different forces act on an object moving in a circular motion.

Centrifugal force:

Force directed outward from the center. E. g. While turning a bus in twist way, the passengers strike on the bus wall is due to centrifugal force.

Centripetal force:

The force exerted towards the center is now as centripetal force. E. g. the force act on passengers by the turning car

Now, suppose a particle is exerted to sediment by centrifugal force, then

The rate or velocity at which it sediments is proportional to the force applied

Sedimentation is more rapid when the force applied is greater than the gravitational force of the Earth

Basis of separation is to exert a larger force than does the Earth’s gravitational force.

Basic Principle of Sedimentation

The particles to be separated are suspended in a specific liquid media, held in tubes or bottles which are located in rotor in centrifuge machine, positioned centrally to the drive shaft. These particles are differing in size, shape and density.

As we have already mentioned that,

The rate of sedimentation is dependent upon the applied centrifugal field (G)

G = W²R

Where:

W-Angular velocity of revolving particle (Remember: one revolution of the rotoe is equal to 2 radians)

R- Radial distance from axis of rotation

In terms of revolution per minute, we have W= 2p rev min^-1/ 60

Therefore:

G = W²R

= 4p² (rev min^-1)² R/ 3600

It is expressed as a multiple of the earth’s gravitational field (g=981 cm s^-2)

Hence

RCF, Relative Centrifugal Field=G / g

= 4p² (rev min^-1)² R/ 3600 x 981

= 1.119 x 10^-5(rev min^-1)² R

= x g unit (number times g)

It means, RCF is the ratio of the weight of the particle in the applied centrifugal field to the weight of the same particle when acted by gravity alone. Therefore the rotor speed, radial dimensions and time of the rotor must be quoted during the centrifugation.

However:

This is not the only case in Biochemical experiments as biological samples are always found in dissolved or suspended form in a solution. Thus, the rate of sedimentation not only depends on the centrifugal field but also on

1. Mass of particle

2. Density of particle

3. Density and viscosity of the medium used

4. The extent to which its shape deviates from spherical

Now according to Newton’s Second law of Motion, the centrifugal force exerted on particle is

F = M. a

= M. W²R

Where:

M: mass of particle

a: acceleration while in angular motion= W²R

Increasing the sharpness of a turn, w r decreases. Since r is linear w has greater effect on the particle.

It causes the molecules to sediment down the centrifuge tube. They start to move downward to sediment; however they encounter opposing force, a frictional resistance in their movement.

Frictional force = f. dr/dt

= 6phR_p (dr/dt)

Where:

f- Frictional force

dr/dt- rate of sedimentation expressed as the change in radius with time (velocity v)

h-Viscosity coefficient of medium

R_p- radius of sedimenting particle

The sedimenting molecule must also displace the solvent into which it sediments and give rise to a buoyant force

Buoyant force= mass x a

= V. d_m W²R

Where:

V- Specific volume of the molecule

d_m– Density of the medium

While sedimenting, the velocity of the particle increases until equals the frictional force resisting its motion through the medium. This is an equilibrium state when the particles stop to move or sediment.

Centrifugal force = Frictional force + Buoyant force

M. W²R = 6phR_p (dr/dt) + V. d_m W²R

4/3p R_p³d_pW²R = 6phR_p (dr/dt) + 4/3p R_p³d_mW²R

4/3p R_p³(d_p-d_m) W²R = 6phR_p (dr/dt)

dr/dt = 2/9h R_p²(d_p-d_m) W²R

v = 2/9h R_p²(d_p-d_m) W²R

dr/dt = v, is the velocity of the sedimenting particle

Mass = Deansity x Volume

d_p = Density of particle

d_m= Density of medium

From above equation, it seems clear that velocity is proportional to its size, to the differences in density between the particle and medium and to the applied centrifugal field. It is zero when the density of the particle and medium are equal. It decreases when the viscosity of the medium increases.

Since the R_pis in square form, the size of particle has greater influence on velocity.

For a particle, h, R_p^,d_p^,d_m and W all are constants

t = 9h/ [2W²R_p²(d_p-d_m) ] In R_b/R_t

Where

t = the sedimentation time in seconds

R_t = radial distance from the axis of rotation to liquid meniscus

R_b= radial distance from the axis of rotation to bottom of tube

It is now clear that a mixture of heterogeneous approximately spherical particles can be separated by centrifugation on the basis of their densities, their sizes and etc.

t µ 1 / R_p²

It means, higher the size particles, faster is the sedimentation of it and smaller the size slower is the sedimentation (takes longer time).

Centrifuges and their uses

1. Low Speed Centrifuge

· Least expensive and simplest in many design

· Maximum rotor speed of 4000-6000rpm (3000-7000 X g)

a) Small bench centrifuges

· To collect small amounts of materials (250mm³⁾ that is rapidly sediment (1-2 min)

· No special cooling system

· Ambient air flows around the rotor to cool the system

· Use to rapid sedimentation of blood samples

b) Large capacity refrigerated centrifuges

· Refrigerated rotor chambers for cooling the sample

· Large volumes 10, 50 and 100 cm³ processing depending upon the rotors and tubes

· Maximum capacity of 1.25 dm³

· Rotors are mounted on a rigid suspension

· Erythrocytes, coarse or bulky precipitates, yeast cells, nuclei and chloroplasts

2. Microcentrifuge

· Maximum rotor speed of 12000rpm with RCF of 10000g

· Have total capacity of 1.5ml over very short time (0.05-5 min)

· Use to sediment large particles like cell ppt

3. High speed refrigerated centrifuge

· Maximum rotor speed of 25000rpm with RCF of 60000g

· Have total capacity of 1.25 dm³

· Interchangeable fixed angle and swinging buckets rotors

· Use to collect microorganisms, cellular debris, larger cellular organelles and proteins precipitates by ammonium sulphate

· Not use for viruses and smaller organelles like ribosome

4. Continuous flow centrifuge

· Relatively simple and high speed centrifuge

· Special design rotor (long and tubular) with non interchangeable system

· Have total capacity of 1-1.25 dm³/min with continuous flow

· Particles sediment at wall and excess clarified medium overflows through an outlet port

· Use to collect bacterial and yeast cells from their mass culture of about 100-500 dm³

5. Ultracentrifuge

· Powerful with speed

· 2 types

a) Preparative ultracentrifuge

Maximum rotor speed of 30000-80000rpm with RCF of 600000g

Highly sophisticated with refrigerated, sealed and evacuated to minimize excess heat generate

More sophisticated temperature monitoring system employing an infrared temperature sensor

Overspeed control system to prevent operation of rotor above its max rated speed

Vibration minimize system (a flexible drive shaft system) during unequal loading of the centrifuge tubes

Enclosed in heavy armour plating

Airfuse for some biochemical applications requiring high centrifugal force

Use for sediment macromolecule/ligand binding kinetic studies, steroid hormone receptor assays, separation of major lipoprotein from plasma and deproteinisation of physiological fluids for amino acid analysis

b) Analytical ultracentrifuge

Maximum rotor speed of 70000rpm with RCF of 500000g

Highly protective chambers with refrigerated and evacuated system also have an optical system to enable the sedimenting material to be observed throughout the process.

Three types of optical system, a light absorption system, alternative Schlieren system and Rayleigh interferometric system (both measures refractive index of solution)

Design and types of Preparative Rotors

These are rotating instruments in Centrifuges

During rotation at high speed, higher stress forces generated

Made up of aluminum alloy and titanium alloy which don’t rust-brass, steel or Perspex

Can tolerate nearly twice the centrifugal force of rotors

Protective coating to the metal surface by anodizing or by applying black epoxy paint

Various types of rotors

a) Swing bucket rotors

· Staring off in a vertical position but during acceleration of the rotor swing out to horizontal position

CENTRIFUGATION . RCF CALCULATION

The relative centrifugal force (RCF) can be calculated from the following equation:

RCF = (1.119 x 10-5) (rpm)²(r)

Where rpm is the speed of rotation expressed in revolutions per minute and r (radius) is the distance from the axis expressed in cm. The RCF units are "x g" where g represents the force of gravity. RCF can also be determined from the NOMOGRAPH below. Place a straight edge to intersect the radius and the desired RCF to calculate the needed rpm. Alternatively place the straight edge on the radius and the rpm to calculate the g-force. For example, spinning a sample at 2500 rpm in a rotor with a 7.7 cm radius results in a RCF of 550 x g.

SEPARATION METHODS IN PREPARATIVE ULTRACENTRIFUGATION

1. Differential Centrifugation: The process of differential centrifugation is based on the fact that organelles have differences in size, shape and density. As a result, the effect of gravity on each is different. We can use this principle to separate an organelle from a homogenous solution of particles by artificially controlling the gravity of a solution. This is done by putting the solution in a variable speed centrifuge and rotating them at a high rate of speed. This creates a force that can be much greater than the force of gravity, and particles that would normally stay in solution will fall out and form a pellet at the bottom of the tube.

Differential centrifugation schemes involve stepwise increases in the speed of centrifugation. At each step, more dense particles are separated from less dense particles, and the successive speed of centrifugation is increased until the target particle is pelleted out. The final supernatant is removed, the pellet is resuspended and further study or purification can be done on it. The fractionation of rat liver is an example of how this process works:

2. Density Gradient Centrifugation

Density gradient centrifugation is a technique that allows the separation of cells, organelles and macromolecules, depending on their size, shape and density.

A density gradient is created in a centrifuge tube by layering solutions of varying densities with the dense end at the bottom of the tube. Cells and large molecules are usually separated on a shallow gradient of sucrose or other inert carbohydrates even at relatively low centrifugation speeds, while macromolecules such as proteins and nucleic acids are separated at higher centrifugation using ultracentrifuges.

Criteria for an ideal density gradient centrifugation medium are:

the additive must form a solution within the required density range

the additive must not interfere with, or damage, the sample

the solvent must be compatible with the sample

the solution must have a refractive index within the practical range, as well as a low viscosity

The additive must be easily removable from the sample.

The additives for density gradient centrifugation can be divided into four main categories:

Salts of Alkali Metals

These solutions fulfill most of the above requirements. However, due to the high ionic strength, hydrogen bonding within biological macromolecules (protein, nucleic acid - protein complexes) is impaired by a chaotropic effect. Therefore these salts are mainly used for DNA and RNA separations. Cesium chloride is used most frequently. Other useful salts include sodium iodide, sodium bromide, cesium sulfate and cesium acetate. Potassium tartrate has been used to separate viruses from host cells.

It should be kept in mind that the density of the sample is highly dependent on the hydration of the macromolecule, which in turn depends to a large extent on the dehydration power of the salt solution.

Neutral, Water-Soluble Molecules

In this class of compounds sucrose is most widely used. It has a useful density range of up to 1.29. This range can be increased to 1.37 by addition of glucose or by dissolving sucrose in D₂O. Sucrose has very little effect on macromolecules, but affects enzyme activity. Due to its high osmotic pressure, sucrose solution dehydrates cells and their organellae very efficiently. Glycerol solutions are the preferred media for the separation of enzymes because they do not affect enzyme activity. They exhibit a high viscosity, requiring prolonged centrifugation times. More importantly however, glycerol penetrates biological membranes.

Hydrophilic Macromolecules

Dextran gradients have been used for the separation of microsomes. Separations achieved with dextrans show similar results to those obtained by using synthetic sucrose/ epichlorohydrin co-polymers. In some cases bovine serum albumin has been applied, but the preparation of an appropriate solution is very difficult.

Synthetic Molecules

These additives are the sodium or methyl glucamine salt of triiodobenzoic acid and of metrizoic acid. It should be kept in mind that the parent acid of these salts may precipitate on adjusting the pH to acidic values. Metrizamide, a covalently bonded compound of glycosamine and metrizoic acid is most widely used. This additive forms solutions of relatively low viscosity. These solutions are stable over a wide range of pH and ionic strength, and show practically no interference with the analytes.

Density gradient centrifugation methods are of two types, the rate zonal technique and the isopycnic (isodensity or equal density) technique.

Rate Zonal Technique: When mixtures of cellular extracts are layered on top of a density gradient in a tube and subjected to centrifugation, the various components move through the gradient at different rates that are dependant on their sizes and shapes. These different components appear as distinct bands or zones in the gradient with large components migrating farthest in the tube in a given period of time. The rate with which a fraction moves the fixed distance in the gradient tube is dependant of its sedimentation value (S) that, in turn is determined by the size and shape of that fraction. By comparing the different position of the components in the gradient, it is possible to make an approximate measurement of their molecular weight. It is, however, difficult to precisely determine these molecular weights, as this requires knowledge about the shape of these molecules, which is hard to determine with accuracy. This density gradient separation technique is called rate zonal centrifugation and is usually performed with a shallow sucrose gradient. The different components being separated by this technique are denser than any of the sucrose concentrations used in the gradient. Samples are, therefore, centrifuged just long enough to separate the components of interest. Longer centrifugation than necessary would allow all components to form a pellet at the bottom of the tube. One of the most important applications of this technique over the past decades was the separation of transfer RNA (4S) from ribosomal RNA that forms three different classes with distinct sedimentation values 23S, 16S and 5S. This helped to facilitate the characterization of the protein synthesizing system.

Isopycnic Technique: A second density gradient technique, called equilibrium density-gradient centrifugation is used to separate cellular components on the basis of their buoyant density. In this case the cellular mixture is centrifuged through a steep density gradient that contains a high concentration of sucrose, or more often, cesium chloride (CsCl). In these gradients, the molecules being studied have a density somewhere in between the highest and lowest densities of sucrose or CsCl generated in the gradient. The components of a sample begin to move down this gradient in the same way as they do in a rate-zonal density gradient. When a component of the mixture reaches a point where the density of the solution is equal to its own density, it stops moving further and forms a distinct band. The position of the band in the tube is characteristic of the buoyancy of that component. Buoyancy or buoyant density of a substance is its tendency to float in a medium, which in this case is the density gradient. Hence soluble proteins which have similar density (p=1.3 g cm^-3in sucrose solution) can not usually be separated by this method, whereas subcellular organelles (e.g. Golgi apparatus p=1.11 g cm^-3, mitochondria p=1.19 g cm^-3 and peroxisomes p=1.23 g cm^-3in sucrose solution can be effectively separated.

Equilibrium density gradient centrifugation using CsCl was for decades the method of choice in the purification of highly pure plasmid DNA. Meselson and Stahl, who developed this technique, were the first to use it in an experiment that provided evidence for the semi-conservative replication of DNA and confirmed the double helix structure of DNA proposed by Crick and Watson.