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4.1: Protein Purification - Biology

4.1: Protein Purification - Biology


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Assays, Specific Activity, Initial Fractionation

A successful protein purification procedure can be nothing short of amazing. Whether you are starting off with a recombinant protein which is produced in E. coli, or trying to isolate a protein from some mammalian tissue, you are typically starting with gram quantities of a complex mixture of protein, nucleic acids, polysaccharide, etc. from which you may have to extract milligram (or microgram!) quantities of desired protein at high purity, and hopefully with high yield.

The first step in any purification is the development of a specific assay for the protein of interest. The specific assay can be based upon some unique characteristic of the protein of interest

  • Enzymatic activity
  • Immunological activity
  • Physical characteristics (e.g. molecular mass, spectroscopic properties, etc.)
  • Biological activity
  • Ideally, an assay should be
    • Specific (you don't want a false positive)
    • rapid (you don't want to wait a week for the results)
    • sensitive (you don't want to consume all your sample in order to assay it)
    • quantitative (you need an accurate way to measure the quantity of your protein at each step in the purification)

Western Blotting

Antibodies can be used in a method called Western blotting, which is useful for determining levels of protein expression and for assaying proteins during purification. This method usually involves the following steps:

  1. A protein sample is subjected to polyacrylamide gel electrophoresis.
  2. After this the gel is placed over a sheet of nitrocellulose and the protein in the gel is electrophoretically transfered to the nitrocellulose.
  3. The nitrocellulose is then soaked in gelatin to "block" its ability to non-specifically bind proteins.
  4. The nitrocellulose is then incubated with the specific antibody for the protein of interest.
  5. The nitrocellulose is then incubated with a second antibody which is specific for the first antibody. For example, if the first antibody was raised in rabbits, the second antibody might be termed "goat anti-rabbit immunoglobulin". What this means is that rabbit immunoglobulins were used to elicit an antibody response in goats. The goat antibodies (polyclonal) will include those which recognize the conserved region in the rabbit antibodies. Since the Fc region is conserved, it will bind to any and all rabbit antibodies, including those on the nitrocellulose paper.
  6. The second antibody will typically have a covalently attached enzyme which, when provided with a chromogenic substrate, will cause a color reaction.
  7. Thus the molecular weight and amount of the desired protein can be characterized from a complex mixture (e.g. crude cell extract) of other proteins.

In a variation of the above, the protein sample may be blotted directly on a nitrocellulose paper (called a dot blot) without first running a gel. This may be desirable if, for example, the antibody is monoclonal and recognizes an epitope which is dependent upon native structure (which would be destroyed upon running an SDS PAGE).

In addition to their varied uses, antibodies can also be used to purify proteins.

  • If relatively large amounts of an antibody can be obtained, they can be covalently attached to a chromatography resin (e.g. sephadex beads).
  • If a crude cell extract is run over such a column, only the protein of interest should bind, and everything else will flow through.
  • The bound protein can then be eluted. This is typically achieved by moderately low pH conditions (using acetic acid). As long as the protein of interest is not irreversibly denatured by such conditions, the method will work quite well.
  • One potential pitfall involves that of monoclonal antibodies being utilized to purify mutant proteins. The regions of the protein comprising the epitope cannot be modified without destroying the ability of the antibody to bind. Thus, the use of monoclonal antibodies in a purification scheme may preclude its use in purifying certain mutants.

Protein purification can be thought of as a series of fractionation steps designed so that:

  • The protein of interest is found almost exclusively in one fraction (and with good yield)
  • A significant amount of the contaminants can be found in a different fraction

During purification you will need to monitor several parameters, including:

  1. Total sample volume
  2. Total sample protein (can be estimated by A280; 1.4 ~ 1.0 mg/ml)
  3. Units of activity of desired protein (based on specific assay)

This basic information will allow you to keep track of the following information during each step of purification:

  1. % yield for each purification step
  2. Specific activity of the desired protein (units/mg total protein)
  3. Purification enhancement of each step (e.g. "3.5x purification)

In designing a purification scheme you typically have to balance purification with yield.

  • For example, it may be relatively straightforward to obtain 90% pure material with good yield.
  • However, it may be difficult to improve that purity an additional few percentile with good yield.
  • The planned application of the purified protein determines the target purity.
  • If the protein is to be used to determine amino acid sequence information, maybe 90% is acceptable. However, if the material is to be used in clinical trials, 99.99+% may be the target purity.

Initial steps in purification

Figure 4.1.1: Purification steps

  • It is extremely helpful to have some information not only on the general physical and chemical characteristics of the protein you are trying to purify, but also on the contaminating components.
  • For example, many E. coli proteins are generally low molecular weight (<50,000 Da) and somewhat acidic in isoelectric point

Usually the initial steps in purification make use of general physical and/or chemical differences between soluble proteins and other cell components.

  • For example, soluble proteins can be separated from general cellular debris, and intact cells, by centrifugation.
  • Thus, cells are physically disrupted (via homogenization or a cell press) to allow release of cell contents. This is then followed by centrifugation to separate generally soluble components from those which are insoluble.
  • It is at this point that data collection begins in order to monitor the purification.

Nucleic acids can sometimes be readily removed from the sample by the addition of large cationic compounds such as polyethylene imine, or streptomycin sulfate.

  • The nucleic acids bind to these compounds via electrostatic interactions and the complex precipitates and can be removed via centrifugation.
  • The same general result can be obtained by mixing in ion exchange resins which are anion exchangers (i.e. the resins contain cationic groups) and then filtering or centrifuging to remove. As with either method, it should be confirmed that the desired protein is not bound as well.

Crude fractionations of proteins can be achieved by adding various quantitites of precipitants such as ammonium sulfate, or polyethylene glycol (PEG).

  • For this type of purification step an initial experiment is performed to monitor the fraction of overall protein, as well as desired protein, remaining in solution (and pellet) as a function of precipitant concentration.

Ammonium Sulfate (% saturated)

0

10

20

30

40

50

60

70

80

90

Sample A280

1000

900

600

200

100

75

50

40

25

20

Activity assay(units)

200

200

200

190

170

100

30

5

0

0

Figure 4.1.2: Protein activity as a function of precipitant concentration

  • In this particular example we are in luck: at around 30% ammonium sulfate we can precipitate about 80% of the total protein concentration in our sample, yet our activity assay for our desired protein indicates that about 95% of our desired protein is still soluble.
  • At 80% ammonium sulfate all of our desired protein has precipitated. Thus, from these results we would do the following:
  1. Add ammonium sulfate to our sample to a concentration of 30% saturation
  2. Centrifuge and discard the pellet
  3. Add ammonium sulfate to 80% saturation
  4. Centrifuge and keep the pellet. Resuspend the pellet in buffer to solubilize the protein.
  • We would expect about a 5-fold purification with about 95% yield.

Column Chromatography - Ion exchange; Dialysis and Concentration

Column chromatography

After initial fractionation steps the typical procedure is to move to column chromatography.

  • In column chromatography we have a glass tube (column) which is filled with a material ("resin") which has certain physical/chemical characteristics.
  • These characteristics allow it to interact in various ways with different proteins.
  • Some common types of chromatographic resins include:
  1. Ion exchange
  2. Hydrophobic
  3. Gel filtration
  4. Affinity

Ion exchange

Ion exchange resins contain charged groups.

  • These may be acidic in nature (in which case the resin is a cation exchanger)
  • or basic (in which case it is an anion exchanger).
  • Cation and anion exchangers may be broken down further into weak and strong exchangers (reflecting binding affinity).

Type of exchanger

Functional group

Common name

Weak cation exchanger

carboxymethyl

CM cellulose/sephadex

Strong cation exchanger

sulfopropyl

SP sephadex

Weak anion exchanger

diethylaminoethyl

DE cellulose/sephadex

Strong anion exchanger

quaternary amine

QAE sephadex

  • Usually, samples are loaded under low ionic strength conditions and bound material is eluted using either a step or gradient elution of buffer with higher ionic strength.
  • Generally speaking, a protein will bind to a cation exchange resin if the buffer pH is lower than the isoelectric point (pI) of the protein, and will bind to an anion exchange resin if the pH is higher than the pI.

Figure 4.1.3: Protein binding to resins

  • Knowledge of the pI of the protein is therefore helpful in designing a purification protocol using ion exchange resins (however, you can always simply try different resins to see which works best).

Generally speaking, ion exchange columns are short and fat in dimensions.

Elution of proteins from ion exchange resins

  • Proteins bound to ion exchange resins are bound via non-covalent ionic (salt-bridge) interactions. We can compete for these ionic binding sites on the resin with other ionic groups, namely, salts
  • There are two general types of methods when eluting with a salt solution: 1. Gradient elution and 2. Step elution
  • A gradient elution refers to a smooth transition of salt concentration (from low to high) in the elution buffer. Weakly binding proteins elute first, and stronger binding proteins elute last (i.e. they require higher salt concentrations in the buffer to compete them off the column)
  • A gradient salt concentration can be made using a gradient maker. In its simplest form, this consists of two containers (must be the same shape) connected by a siphon (or tube at the bottom). One container contains the low salt buffer, and the other contains high salt buffer. The buffer is withdrawn from the low salt container:

Figure 4.1.4: Gradient maker

  • This will produce a linear gradient from low to high salt concentrations over the total volume of the gradient

Figure 4.1.5: Salt concentration and volume

  • If we know the concentration range of salt over which a protein of interest will elute we can simply elute with a buffer containing that concentration of salt. This is known as a step elution.
  • Step elutions are generally faster to run, and elute the protein in a smaller overall volume than with gradient elutions. They generally work best when contaminants elute at a significantly different salt concentration than the protein of interest

Figure 4.1.6: Step elution

Note that after ion exchange chromatography the protein of interest will be in a buffer with a potentially high salt concentration. This must be taken into account before proceeding with the next step in the purification scheme

Dialysis

  • After an ammonium sulfate precipitation step, or an ion exchange chromatography step, the protein of interest may be in a high salt buffer. This may be undesirable for several reasons. How do we get rid of salt in our sample?
  • One of the most common methods is that of dialysis
  • The method of dialysis makes use of semi-permeable membranes. In the simplest example, this membrane is manufactured in the form of tubing (looking much like a sausage casing)
  • The main feature of this membrane is that it is porous. However, the pore size is such that while small salt ions can freely pass through the membrane, larger protein molecules cannot (i.e. they are retained). Thus, dialysis membranes are characterized by the molecular mass of the smallest typical globular protein which it will retain.
  • This is commonly referred to as the cutoff of the tubing (e.g. Spectrapore #6 dialysis tubing has a cutoff of 1,000 Daltons, meaning that a 1,000 Dalton protein will be retained by the tubing but that smaller molecular mass solutes will pass through the tubing)
  • Dialysis proceeds by placing a high salt sample in dialysis tubing (i.e. the dialysis "bag") and putting it into the desired low salt buffer:

Figure 4.1.7: Dialysis

  • Over time the concentration of low molecular mass solutes within the bag, and in the low salt buffer, will come achieve equilibrium. In practical terms (for the above case) salt molecules will diffuse out of the bag into the low salt buffer:

Figure 4.1.8: Salt diffusion

  • At equilibrium the salt concentration of the sample can be calculated as follows:

$$frac {(sample : volume) imes (sample : salt : concentration) + (buffer : volume) imes (buffer : salt : concentration)}{total :volume} = final :salt : concentration$$

Note

Often the buffer salt concentration is 0 M

  • The buffer volume for the dialysis is a function of the required final concentration of salt in the sample

Example 4.1.1:

Dialysis example

We have a 10ml protein sample from an ion exchange column elution pool which contains 1.0M NaCl. For our next step in the purification we can have no more than 1mM NaCl in the sample.

Therefore, the required buffer volume would be (total vol - sample vol) = 9.990 L (or ~ 10 L)

  • Thus, if we dialyzed 10mls of sample (with 1.0M NaCl conc) in 10 L of water after equilibrium the NaCl concentration in the sample would be 1.0 mM.

  • Note that in the above example this would commonly be referred to as a "1:1,000" dialysis.
  • Suppose that we don't want to make up 10 L of buffer? We can actually achieve the same results with two sequential "1:32" dialyses (i.e. the square root of the 1:1,000 dialysis - in other words, two sequential 1:32 dialyses is equivalent to a single 1:1,000 dialysis):

First dialysis versus 310 ml of buffer: sample NaCl conc will be (10*1.0)/(320) = 31 mM

Second dialysis versus 310 ml of buffer: sample NaCl conc will be (10*0.031)/(320) = 0.97 mM

Thus, instead of making 10 L of buffer, we could make only 620 ml and achieve the same results with two dialysis steps

  • In this case, removing the salt would take twice as long, i.e. we need to perform two dialysis steps. How long does dialysis take?

A useful rule of thumb is that for most types of dialysis tubing the dialysis is 80% compete after four hours

  • One consequence of dialysis to watch out for is that while salt ions are moving out of the bag, water molecules are moving into the bag. Thus the volume of sample may actually increase (the bag will swell) and, therefore, the protein concentration will decrease
  • In the extreme case, the bag may actually swell to the point of rupture. Therefore, it is a good idea not to fill the bag completely, but leave a void to allow for potential swelling.

Concentration

  • What if our protein sample is too dilute for our needs? How can we concentrate our samples?
  • One common method is, again, to use a semi-permeable membrane for this purpose.
  • A very simple method is to place our sample in a dialysis bag and coat it with a high molecular weight solute which can readily be dissolved by the buffer.
  • For example, polyethylene glycols and polyvinyl pyrolidones can have very large molecular masses (i.e. 20,000 Da). These compounds are also readily dissolved in water. If our sample in a dialysis bag is coated with dry forms of the above polymers, water will leave the dialysis bag (it can go through the pores) and hydrate the polymers. The result is a decrease in volume of buffer in the dialysis bag (the protein will be concentrated).
  • In another variation, the semi-permeable membrane is manufactured into a flat disk and placed at the bottom of a container which holds our sample. In one method the container is pressurized and forces buffer out of the container (protein is retained and is concentrated). In another method, the vessel is centrifuged and the centripetal force achieves the same goal as pressure in the previous example.

For both dialysis and concentration, it is essential that the membrane does not interact with the protein (i.e. has no affinity for, and will not bind, the protein)

  • With the pressure type concentrators, dialysis and concentration can be achieved in tandem. For example, the sample can be concentrated and then buffer added to the sample. The sample is then concentrated again. Every time buffer is added the salt concentration is reduced. After repeated cycles of this, the salt concentration is at the desired level and the sample is concentrated to the desired final volume.

Gel Filtration, Affinity and Hydrophobic resins; Preparation of Resin, Plumbing

Gel filtration

Gel filtration does not rely on any chemical interaction with the protein, rather it is based on a physical property of the protein - that being the effective molecular radius (which relates to mass for most typical globular proteins).

  • Gel filtration resin can be thought of as beads which contain pores of a defined size range.
  • Large proteins which cannot enter these pores pass around the outside of the beads.
  • Smaller proteins which can enter the pores of the beads have a longer, tortuous path before they exit the bead.
  • Thus, a sample of proteins passing through a gel filtration column will separate based on molecular size: the big ones will elute first and the smallest ones will elute last (and "middle" sized proteins will elute in the middle).

Figure 4.1.9: Gel filtration

  • If your protein is unusually "small" or "large" in comparison to contaminating proteins then gel filtration may work quite well.

Where will a protein elute in a gel filtration experiment?

  • There are two extremes in the separation profile of a gel filtration column.
  • There is a critical molecular mass (large mass) which will be completely excluded from the gel filtration beads. All solutes in the sample which are equal to, or larger, than this critical size will behave identically: they will all eluted in the excluded volume of the column
  • There is a critical molecular mass (small mass) which will be completely included within the pores of the gel filtration beads. All solutes in the sample which are equal to, or smaller, than this critical size will behave identically: they will all eluted in the included volume of the column
  • Solutes between these two ranges of molecular mass will elute between the excluded and included volumes

Figure 4.1.10: Excluded vs. included volume

As a general rule of thumb, the excluded volume (Vo) is approximately equal to one third of the column volume, the included volume is approximately equal to two thirds of the column volume

  • In gel filtration the resolution is a function of column length (the longer the better)
  • However, one drawback is related to the maximum sample volume which can be loaded. The larger the volume of sample loaded, the more the overlap between separated peaks. Generally speaking, the sample size one can load is limited to about 3-5% of the total column volume.
  • Thus, gel filtration is best saved for the end stages of a purification ,when the sample can be readily concentrated to a small volume.
  • Gel filtration can also be used to remove salts from the sample, due to its ability to separate "small" from "large" components.
  • Finally, gel filtration can be among the most "gentle" purification methods due to the lack of chemical interaction with the resin.

Affinity chromatography

Affinity chromatography is a general term which applies to a wide range of chromatographic media. It can be basically thought of as some inert resin to which has been attached some compound which has a specific affinity for your protein of interest.

  • Thus, a specific antibody attached to an inert resin would be a type of affinity chromatography.
  • Other examples might include: a protease inhibitor attached to some matrix, designed to bind a specific protease
  • a cofactor bound to some matrix, designed to bind to a particular enzyme
  • a metal ion bound to a matrix, designed to chelate a protein with a metal binding site, and so on.

In each case, the type of resins used and the method of attachment may vary, as will the method of elution. One generalization regarding method of elution is that the bound ligand can be competed off of the column's functional group by including in the elution buffer a high concentration of the free functional group. For example, if the functional group of the column is a cofactor, then the bound protein can be competed off the column by passing a buffer containing a high concentration of cofactor (or cofactor analog) through the column.

Other methods of elution include changing the buffer conditions such that the protein is no longer in the native state (since it is the native state which confers the structure required for the specific binding interaction). This can be achieved by changing pH or by adding denaturing agents such as urea or guanidine.

With affinity chromatography, typically the purification achieved in a single step can be dramatic - on the order of several thousand fold. Single step purifications with specific affinity columns are not unheard - in fact it is an ideal goal of purification - a matrix which recognizes only the protein of interest and none other.

Hydrophobic resins

Hydrophobic resins contain a non-polar functional group, such as an alkane or aromatic group.

  • Many proteins are able to sequester such groups on their surface and this exclusion from solvent provides the basis of the binding energy (i.e. the "hydrophobic effect").
  • This interaction is enhanced by increasing ionic strength, such that proteins may bind under high salt conditions and elute under low salt conditions.
  • As such these columns may be used to not only provide purification, but to desalt samples (for example after an initial ammonium sulfate precipitation).
  • It is usually not possible to predict in advance which particular resin will bind a given protein, this is usually determined empirically. However, the longer the alkane, or the larger the aromatic compound, the stronger the binding typically will be.

Due to the nature of hydrophobic interactions and ionic strength, hydrophobic chromatography and ion exchange chromatography can be conveniently used sequentially. For example, after ion exchange the protein is in high salt conditions, thus it can be loaded directly onto a hydrophobic column. Conversely, a hydrophobic column is eluted in low salt, which is a requirement for binding to an ion exchange resin.

A distinction should be noted between hydrophobic interaction chromatorgraphy and reverse phase chromatography

  • Hydrophobic interaction chromatography is performed in aqueous solvent conditions and changes in ionic strength are used to elute the column. The protein typically binds in the native state via hydrophobic groups located on the surface of the protein. The native state is retained during the elution conditions
  • Reverse phase chromatography utilizes a hydrophobic solvent (typically acetonitrile) and the binding of a ligand is a function of the phase partition between the hydrophobic nature of the solvent and column functional group. Proteins are typically denatured in such solvents and bind due to the hydrophobic nature of the entire polypeptide sequence. Since the majority of hydrophobic groups are located in the core of globular proteins, the binding is related to the denaturation of the protein and the accessibility of these groups to the column functional groups. Proteins can be purified using reverse phase chromatography, but usually must be refolded in some way to regain functionality (i.e. the native state)

Preparation of resins

The steps in preparing a chromatographic resin typically involve:

  1. Hydration of resin
  2. Decanting fines
  3. Equilibrating the resin and preparing a slurry
  4. Degassing the slurry
  • Resins come either dry or preswollen. If they are dry they need to be hydrated. This is usually accomplished by mixing the dry resin with buffer and letting it hydrate slowly overnight (or faster at higher temperatures
  • After the resin has hydrated and settled, very fine particles will settle at the top. These "fines" slow the flow rate of the packed resin. The settled resin is therefore carefully decanted to discard these fines.
  • The resin is then equilibrated in the buffer to be used for the analysis. Equilibration usually involves pH'ing the resin, or buffer exchanges. Never use a stir bar when pH'ing the resin (it can mechanically shear the resin and produce fines), rather stir the resin slurry with a stir rod.
  • After the equilibrated resin has settled, an equal volume of buffer is added to produce a 50% slurry of resin. This is usually "thin" enough to allow air bubbles to escape when packing the column.
  • Finally, the slurry is degassed prior to packing the column. This will help minimize the formation of air bubbles.

Packing the column

Low pressure columns are typically packed using gravity.

  • Add a small amount of buffer to the bottom of the column.
  • Place a packing reservoir on the top of the column. Since we will be using a 50% slurry we will have a volume which is 2x the column volume and its best to pour the resin in all at one time. Thus, the packing reservoir should have a volume equal to, or greater, than the column volume.
  • Carefully pour the resin slurry into the packing reservoir/column, avoiding the introduction of air bubbles as much as possible
  • Let the column sit for about 5 minutes to allow large air bubbles to escape
  • Open the column valve at the bottom and allow the column to pack under gravity
  • Note the top of the resin bed. It will move down as the column packs. When the column is packed the top of the resin bed will no longer move down.

Plumbing

Chromatography systems may be run using only gravity and a beaker to collect the appropriate fraction. Most common systems, however, will include the following:

  • A pump. Usually a peristaltic pump with variable flow rate and a communications port for a controller. The pump is usually set up to push buffer through the column, rather than sucking buffer out of the column (which can cause a low pressure condition with production of air bubbles)
  • A detector. This is typically a UV (A280) detector. Most detectors are of the two-cell type - meaning that you can have plain buffer as a blank in the detector while analyzing your column fractions. The detector sends the absorbance information to a chart recorder to be displayed (printed)
  • A fraction collector. This allows you to collect fractions either by number of drops (~30 per ml) or by time. In conjunction with a controllable pump, time collection translates to volume. The fraction collector will typically have an communications port to output a signal when it changes fractions and to receive commands from the detector/controller on some sophisticated systems.
  • A chart recorder. This will print a continuous trace of the detector output and the fraction collector event marker (signalling when a fraction changes). Fractions can also be read individually on a UV spectrophotometer if a chart recorder is unavailable.

Figure 4.1.11: Chromatography setup

  • If a gravity system is used, a safety loop should be installed to prevent the column from drying up if the buffer is used up when the column is unattended

Figure 4.1.12: Safety loop

Note that the bottom of the safety loop is lower than the outlet to the fraction collector.

Running the Experiment, Resolving Peaks

The following represents an example of a low pressure liquid chromatography (ion exchange resin) experiment.

Sample:

  • Volume = 90 mls
  • A280 = 1.8
  • Total A280 = 162

Column:

  • DE-52 (diethylaminoethyl cellulose; anion exchange)
  • Size = 1.0 x 12.7 cm
  • Volume = = 40 mL

Fraction Collector:

  • 10 mls / fraction (~300 drops/fraction)

The chromatogram for this experiment looked like this:

Figure 4.1.13: Chromatogram

The following events took place during this chromatography run:

1. Note the tick marks on the chromatogram.

  • The "event" marker from the fraction collector notifies the chart recorder when a tube change takes place.
  • The experiment begins with the tick next to the '0' on the x-axis ("tick 0"); this indicates the startof fraction (tube) number 1.
  • The next tick mark ("tick 1") indicates the end of fraction number 1, and the start of fraction number 2.
  • Thus, fractions span the gap between the tick marks.

2. The sample loading is begun at tick 0.

3. Sometime during fraction 5 we begin to notice the absorbence of the column effluent increasing

  • It has taken about (5 fractions x 10mls per fraction) or 50 mls from the start of loading until the detector notes any absorbance.
  • This compares well with the fact that the column volume is about 40 mls and there is some volume associated with the tubing going in and out of the column.
  • Thus, this 'delay' from sample load to sample detection is the dead volume of the system

4. Obviously, some material is not binding to the resin during the loading step. This is the flow-through. Is this some component of the sample which does not have affinity for the resin, or, does it represent that we have exceeded the capacity of the resin?

  • If we have exceeded the capacity of the resin, then the flow-through will have an A280 similar to the sample being loaded
  • Also, prior to exceeding the capacity, the flow-through will have some characteristic A280 which will then transition to another A280 (that of the loaded sample), resulting in a double-plateau chromatogram.
  • In the above experiment the flow-through plateaus around A280=0.5 or about 25% of the absorbance of the load. This would seem to indicate that a component, or component(s), representing one quarter of our sample, does not have affinity for the resin in the column

5. Around fraction 9 we begin to wash the column

  • This makes sense because 9 fractions x 10mls per fraction = 90 mls have loaded and this is equivalent to our original sample volume (i.e. all the sample has loaded)
  • The column is typically washed using the same buffer conditions in the protein sample

6. Around fraction 14 we note the A280 begins to decrease

  • This makes sense given that we determined the dead volume of the system to be approximately 50 mls or 5 fractions. Thus, a wash which was begun at fraction 9 is observed to decrease the absorbance around fraction 14
  • We continue washing the column until the A280 approaches 0 (baseline). In other words, all of the non-binding material in the sample has been washed away

7. After the A280 comes back down to baseline we begin our elution protocol. In this particular experiment we will use a linear gradient of increasing salt (NaCl) concentration (in wash buffer) to compete off the material bound to the ion exchange resin.

8. Our elution has produced two peaks: a small peak centered around fraction 42 and a larger peak centered around fraction 50

  • We will have to assay each peak (and the flow through) to find out where our protein of interest has gone
  • The two elution peaks are fairly well resolved. We could combine fractions 40-44 and call that "peak 1", and combine fractions 46-55 and call that "peak 2".

9. Is there any material left on the column? The integrated areas (i.e. summing the A280's of each fraction in a pool) of the flow-through, peak 1 and peak 2 are as follows

  • Flow through: 4
  • Peak 1: 2
  • Peak 2: 10
  • This gives a total integrated area of 16. Each fraction is 10 mls, so this gives a total A280 = 16 x 10 = 160 which is quite close to the total A280 of our loaded sample.
  • In other words, it looks like our chromatogram is accounting for all the components in our original sample.

10. If our protein of interest was actually peak 1 (and if our yield was 100%), then this column has provided an eight fold purification ( 2 x 10 / 162).


Resolving peaks

  • Contaminating peaks will not necessarily be completely separated from the peak which contains our protein of interest
  • In the following picture there are two components being resolved, and they are present in equimolar amounts (thus, the starting purity is 50%). The yield and purity are listed for the situation where we were to pool each peak by splitting at the midpoint between them (in this particular example the yield and purity are identical in each case)

Figure 4.1.14: Contaminating peaks

  • This gives you some idea of the amount of cross-contamination in each peak as a function of their separation from one another.
  • Software to fit gaussians to a chromatogram can provide this type of information

Pooling for purity verses yield

Do we try to pool to maximize yield or to maximize purity?

Usually, you will probably be pooling fractions in such a way as to maximize the recovery of your protein of interest. However, you always have the option of pooling to increase purity, and if you have lots of protein to work with this may allow you to achieve the desired purity with fewer steps. Here's an example of how it's done:

Figure 4.1.15: Yield vs. purity

  • These are all the same chromatogram, however, we can pool them differently to get better purity (at the expense of yield
  • The blue peak is the peak of interest and it is not resolved from a contaminating peak (in red).
  • The vertical line represents the left-most fraction we use to pool the peak (we pool all fractions to the right of the vertical line to get our protein of interest)
  • In the last panel we see that we can achieve about 98.8% purity if we are willing to part with half our protein!

Monitoring the purification

How do you know when you are finished purifying a protein?

There are several criteria. One criteria is that we cannot improve upon the specific activity of our sample. This value refers to the functional activity of our sample in relationship to the total protein concentration of the sample.

  • In the initial stages of purification this value will be low (not much activity in relationship to the total amount of protein).
  • This value will increase after each purification step as we remove other proteins from the sample.
  • At some point the specific activity will plateau, and by definition, if it is pure we cannot increase the specific activity.
  • There may be a published value for the specific activity which we can compare ours to.

Also, each step of the purification should be monitored by gel electrophoresis.

  • In the initial stages of purification we will probably see a variety of bands, of various molecular weights, on our gel.
  • After the different purification steps, we should see the disappearance of certain bands concomitant with the increasing concentration of a certain band (or bands) representing our protein.
  • If we have successfully purified our protein (and if it is a single polypeptide) we should arrive at a constant specific activity and a single band on a gel.
  • Analytical methods like HPLC or densitometer scanning of a stained gel can give us a quantitative idea of the purity of our final sample.

The following chart represents the typical data one would monitor during a purification:

Step
Total protein (mg)
Total activity (units)
Specific activity (units/mg)
Purification
% Yield
Crude cell lysate
5500
6600
1.2
30-70% Ammonium sulfate cut
1020
5910
5.8
4.8
89.5
DEAE Sephadex pool
187
5070
27.1
4.7
85.8
CM Sephadex pool
102
4420
43.3
1.6
87.2
Phenyl Sepharose pool
56
3930
70.2
1.6
88.9
Gel Filtration pool
32
2970
92.8
1.3
75.6
Affinity resin type #1 pool
5.8
2520
434.5
4.7
84.8
Affinity resin type #2 pool
5.3
2390
450.9
1.0
94.8
Total purification
376
Total yield (%)
36

Protein Purification: Meaning, Principle Strategies, Evaluation and Other Details

Read this article to learn about the meaning, principle strategies, selection and combination of purification techniques, purification of a tagged protein, evaluation of purification yield, concentration of purified protein and analysis of isolated proteins.

Introduction:

Protein purification is a series of processes intended to isolate a single type of protein from a complex mixture.

Protein purification is vital for the characterization of the function, structure and interactions of the protein of interest. The starting material is usually a biological tissue or a microbial culture.

The various steps in the purification process may free the protein from a matrix that confines it, separate the protein and non-protein parts of the mixture, and finally separate the desired protein from all other proteins. Separation of one protein from all others is typically the most laborious aspect of protein purification. Separation steps exploit differences in protein size, physicochemical properties and binding affinity.

Purification may be preparative or analytical. Preparative purifications aim to produce a relatively large quantity of purified proteins for subsequent use. Examples include the preparation of commercial products such as enzymes (e.g., lactase), nutritional proteins (e.g., soy protein isolate), and certain biopharmaceuticals (e.g., insulin).

Analytical purification produces a relatively small amount of protein for a variety of research or analytical purposes, including identification, quantification, and studies of the protein’s structure, post-translational modifications and function. Among the first purified proteins were urease and Concanavalin-A.

Principle Strategies:

Most purification protocols require more than one step to achieve desired level of purity. Each step in the process will cause some loss of product, a yield of 80% in each step is assumed, and therefore, it is advisable to have as few purification steps as possible. Choice of a starting material is key to the design of a purification process.

In a plant or animal, a particular protein usually is not distributed homogeneously throughout the body different organs or tissues have higher or lower concentra­tions of the protein. Use of only the tissues or organs with the highest concentration decreases the volumes needed to produce a given amount of purified protein.

If the protein is present in low abundance, or if it has a high value, scientists may use recombinant DNA technology to develop cells that will produce large quantities of the desired protein (this is known as an expression system) Recombinant expression allows the protein to be tagged, e.g., by a His-tag, to facilitate purification, which means that the purification can be done in fewer steps. In addition to this recombinant expression usually starts with a higher fraction of the desired protein than is present in a natural source.

With background information, assays and sample procedures in place the three phase purification strategies can be considered. The purification has three phases of capture, intermediate purification and polishing, each with specific objective. In capture phase the objectives are to isolate, concentrate and stabilize the target procedure.

During intermediate purification phase the objective is to remove bulk impurities such as other proteins, nucleic acids, endotoxins and viruses. In polishing phase the objective is to remove any trace impurities and closely related substances. Therefore selection and optimum combination of purification techniques for capture, intermediate purification and polishing is necessary to ensure good yield and pure product.

The final purification process ideally consists of sample preparation, including extraction and clarification when required followed by above described three phases of purification. The number of steps will always depend on purity required and intended use of protein.

An analytical purification generally utilizes three properties to separate proteins. First, proteins- may be purified according to their isolectric points by running them through a pH graded gel or an ion exchange column. Second, proteins can be separated according to their size or molecular weight via size exclusion chromatography or by SDS-PAGE (sodium dodecyl sulphate-polyacrylamide gel electrophoresis) analysis.

Proteins are often purified by using 2D-PAGE and are then analysed by peptide mass fingerprinting to establish the protein identity. This is very useful for scientific purposes and the detection limits for protein are nowadays very low and Nano gram amounts of protein are sufficient for their analysis.

Selection and Combination of Purification Techniques:

The aim of this combination is to evolve a fastest route to a product of required purity. For any chromatographic separation each different technique will offer a different performance with respect to recovery, resolution, speed and capacity. A technique can be optimized to focus on one of these parameters for example, resolution to achieve the best between two parameters such as speed and capacity.

A separation optimized for one of these parameters will produce result quite different in appearance from those produced using the same technique but focused on alternative parameters. Therefore, it is always preferable to select a technique to meet the objectives in purification step.

Capacity in the simple model shown represents the amount of target protein loaded during the purification. In some cases the amount which can be loaded may be limited by volume (as in gel filtration) or by large volume of contaminants rather than the amount of target proteins.

Speed is of highest importance at the beginning where contaminants like proteases must be removed as quickly as possible. Recovery becomes increasingly important as the purification progresses because of the increased value of the purified product. The recovery’ is influenced by destructive processes in the samples and unfavorable conditions on the column.

Resolution is achieved by the selectivity of the technique and efficiency of the chromatographic matrix to produce narrow peaks. In general, resolution is most difficult to achieve in final phases of purification where target protein and impurities have very similar properties.

Every technique offers a balance between capacity, speed, resolution and recovery and should be selected to meet the objectives of each purification step. In general, the optimization of any of these parameters can be achieved only at the expense of other and purification step will be a compromise.

The importance of each parameter will vary depending upon whether a purification step is used for capture, intermediate purification or polishing. This will steer the optimization of critical parameters as well as for the selection of suitable media for the step.

Purification of a Tagged Protein:

Adding a tag to the protein gives the protein a binding affinity it would not otherwise have. Usually the recombinant protein is the only protein in the mixture with this affinity, which aids in separation. The most common tag is the Histidine-tag (His-tag) that has affinity towards nickel or cobalt ions. Thus by immobilizing nickel or cobalt ions on a resin, an affinity support that specifically binds to histidine tagged proteins can be created. Since the protein is the only component with a His-tag, all other proteins will pass through the column, and leave the His-tagged protein bound to the resin.

The protein is released from the column in a process called elution, which in this case involves adding imidazole, to compete with the His-tags for nickel binding, as it has a ring structure similar to histidine. The protein of interest is now the only protein component in the eluted mixture, and can easily be separated from any minor unwanted contaminants by a second step of purification, such as size exclusion chromatography or RP-HPLC.

Another way to tag proteins is to add an antigen peptide to the protein, and then purify the protein on a column containing immobilized antibody. This generates a very specific interaction usually only binding the desired protein. When the tags are not needed anymore, they can be cleaved off by a protease. This often involves engineering a protease cleavage site between the tag and the protein.

Evaluating Purification Yield:

The most general method to monitor the purification process is by running a SDS PAGE, of the different steps. This method only gives a rough measure of the amounts of different proteins in the mixture, and it is not able to distinguish between proteins with similar molecular weight.

If the protein has a distinguishing spectroscopic feature or an enzymatic activity, this property can be used to detect and quantify the specific protein, and thus to select the fractions of the separation, that contains the protein. If antibodies against the protein are available, then western blotting and ELISA can specifically detect and quantify the amount of desired protein. Some proteins function as receptors and can be detected during purification steps by a ligand binding assay, often using a radioactive ligand.

In order to evaluate the process of multistep purification, the amounts of the specific protein have to be compared to the amount of total protein. The latter can be determined by the Bradford total protein assay or by absorbance of light at 280 nm, however some reagents used during the purification process may interfere with the quantification.

For example, imidazole (commonly used for purification of polyhistidine-tagged recombinant proteins) is an amino acid analogue and at low concentrations will interfere with the bicinchoninic acid (BCA) assay for total protein quantification. Impurities in low-grade imidazole will also absorb at 280 nm, resulting in an inaccurate reading of protein concentration from UV absorbance.

Concentration of the Purified Protein:

At the end of protein purification, the protein often has to be concentrated.

Different method exist for this purpose is:

1. Lyophilization:

If the solution does not contain any other soluble component than the protein in question the protein can be lyophilized (dried). This is commonly done after a HPLC run. This simply removes all volatile components leaving the proteins behind.

2. Ultrafiltration:

Ultrafiltration concentrates a protein solution using selective permeable membranes. The function of the membrane is to let the water and small molecules pass through while retaining tin- protein. The solution is forced against the membrane by mechanical pump or gas pressure or centrifugation.

Analysis of Isolated Proteins:

This analysis is done by following techniques:

1. Denaturing-Condition Electrophoresis:

Gel electrophoresis is a common laboratory technique that can be used both as preparative and analytical methods. The principle of electrophoresis relies on the movement of a charged ion in an electric field. In practice, the proteins are denatured in a solution containing a detergent (SDS). In these conditions, the proteins are unfolded and coated with negatively charged detergent molecules. The proteins in SDS-PAGE are separated on the sole basis of their size.

In analytical methods, the protein migrates as bands based on size. Each band can be detected using stains such as Coomassie blue dye or silver stain. Preparative methods to purify large amounts of protein require the extraction of the protein from the electrophoretic gel. This extraction may involve excision of the gel containing a band, or eluting the band directly off the gel as it runs off the end of the gel.

In the context of a purification strategy, denaturing condition electrophoresis provides an improved resolution over size exclusion chromatography, but does not scale to large quantity of proteins in a sample as well as the late chromatography columns.

2. Non-Denaturing-Condition Electrophoresis:

An important non denaturing electrophoretic procedure for isolating bioactive metalloproteins in complex protein mixtures is termed ‘quantitative native continuous polyacrylamide gel electro­phoresis’ (QNC-PAGE).


Chromatography

Column chromatography is one of the most common methods of protein purification. Like many of the techniques on this site, it is as much an art form as a science. Proteins vary hugely in their properties, and the different types of column chromatography allow you to exploit those differences. Most of these methods do not require the denaturing of proteins.

To be very general, a protein is passed through a column that is designed to trap or slow up the passing of proteins based on a particular property (such as size, charge, or composition).

There are three main steps to protein purification:

1. Capture. You need to get your protein into a concentrated form. If, for example, you are trying to isolate a protein you have synthesized in an E. coli cell, you could be looking at a protein to junk ratio of 1:1,000,000. For capture purification you need a high capacity method that is also fast. You need a speedy method because your crude solution is very likely to also contain proteases and these can quickly chew up your protein.

2. Intermediate. Intermediate purification requires both speed and good resolution.

3. Polishing. For the final step of purification you need a system that has both good resolution and speed. Capacity is usually irrelevant at this stage.

Some of the more common columns include:

IEX: Ion exchange chromatography. Good for capture, intermediate, and polish.

HIC: Hydrophobic interaction column. Good for intermediate purification.

AC: Affinity chromatography. Good for capture and intermediate purification.

GF: Gel filtration (size exclusion) chromatography. Good polishing step.

Let's look at these types of columns in more detail.

Ion exchange chromatography

Ion exchange chromatography is based on the charge of the protein you are trying to isolate. If your protein has a high positive charge, you'll want to pass it through a column with a negative charge. The negative charge on the column will bind the positively charged protein, and other proteins will pass through the column. You then use a procedure called "salting out" to release your positively charged protein from the negatively charged column. The column that does this is called a cation exchange column and often uses sulfonated residues. Likewise, you can bind a negatively charged protein to a positively charge column. The column that does this is called an anion exchange column and often uses quaternary ammonium residues.

Salting out will release, or elute, your protein from the column. This technique uses a high salt concentration solution. The salt solution will out-compete the protein in binding to the column. In other words, the column has a higher attraction for the charge of salts than for the charged protein, and it will release the protein in favor of binding the salts instead. Proteins with weaker ionic interactions will elute at a lower salt, so you will often want to elute with a salt gradient. Different proteins elute at different salt concentrations, so you will want to be sure you know well the properties of your protein best results.

Also be aware that changes in pH alter the charges in proteins. Be sure you know the isoelectric point of your protein (the isoelectric point is the pH at which the charge of a protein is zero) and make sure the pH of your system is adjusted and buffered accordingly.

The basic steps in using an ion exchange column are:

1. Prep the column. Pour your buffer over the column to make sure it has equilibrated to the required pH.

2. Load your protein solution. Some proteins in the solution don't bind and will elute during this loading phase.

3. Salt out. Increase the salt concentration to elute the bound proteins. It is best to use a salt gradient to gradually elute proteins with different ionic strengths. At the end bump the system with a very high salt concentration (2-3M) to make sure all proteins are off the column.

4. Remove salts. Use dialysis to remove the salts from your protein solution.

Temperature doesn't have a huge effect on column chemistry. However, it is better to work cold since proteins are more stable cold.

Hydrophobic interaction chromatography

Where ion exchange chromatography relies on the charges of proteins to isolate them, hydrophobic interaction chromatography uses the hydrophobic properties of some proteins. Hydrophobic groups on the protein bind to hydrophobic groups on the column. The more hydrophobic a protein is, the stronger it will bind to the column.

Load the proteins in the presence of a high concentration of ammonium sulfate (not ammonium persulfate). Ammonium sulfate is a chaotropic agent. It increases the chaos (entropy) in water, and thereby increases hydrophobic interactions (the more disordered the water, the stronger the hydrophobic interactions). Ammonium sulfate also stabilizes proteins. So as a result of using an HIC column you can expect your protein to be in its most stable form.

The hydrophobic column is packed with a phenyl agarose matrix. In the presence of high salt concentrations the phenyl groups on this matrix binds hydrophobic portions of proteins. You can control elution of different column-bound proteins by reducing the salt concentration or by adding solvents.

Affinity chromatography.

Affinity chromatography relies on the biological functions of a protein to bind it to a column. The most common type involves a ligand, a specific small biomolecule. This small molecule is immobilized and attached to a column matrix, such as cellulose or polyacrylamide. Your target protein is then passed through the column and bound to it by its ligand, while other proteins elute out. Elution of your target protein is usually done by passing through the column a solution that has in it a high concentration of free ligand. This is a very efficient purification method since it relies on the biological specificity of your target protein, such as the affinity of an enzyme for a substrate.

Gel filtration (size exclusion) chromatography d

Gel filtration, or size exclusion, chromatography separates proteins on the basis of their size. The column is packed with a matrix of fine porous beads.

It works somewhat like a sieve, but in reverse. The beads have in them very small holes. As the protein solution is poured on the column, small molecules enter the pores in the beads. Larger molecules are excluded from the holes, and pass quickly between the beads.

These larger molecules are eluted first. The smaller molecules have a longer path to travel, as they get stuck over and over again in the maze of pores running from bead to bead. These smaller molecules, therefore, take longer to make their way through the column and are eluted last.


Purification of Polyhistidine-Tagged Proteins

Rapid Purification of Polyhistidine-Tagged Proteins Using Magnetic Resins

There is a growing need for high-throughput protein purification methods. Magnetic resins enable affinity-tagged protein purification without the need for multiple centrifugation steps and sequential transfer of samples to multiple tubes. There are several criteria that define a good protein purification resin: minimal nonspecific protein binding, high binding capacity for the fusion protein and efficient recovery of the fusion protein. The MagneHis™ Protein Purification System meets these criteria, enabling purification of proteins with a broad range of molecular weights and different expression levels. The magnetic nature of the binding particles allows purification from crude lysates to be performed in a single tube. In addition, the system can be used with automated liquid-handling platforms for high-throughput applications.

MagneHis™ Protein Purification System

The MagneHis™ Protein Purification System uses paramagnetic precharged nickel particles (MagneHis™ Ni-Particles) to isolate polyhistidine-tagged protein directly from a crude cell lysate. Figure 2 shows a schematic diagram of the MagneHis™ Protein Purification System protocol. Polyhistidine-tagged protein can be purified on a small scale using less than 1ml of culture or on a large scale using more than 1 liter of culture. Samples can be processed in a high-throughput manner using a robotic platform such as the Beckman Coulter Biomek® FX or Tecan Freedom EVO® instrument. Polyhistidine-tagged proteins can be purified under native or denaturing (2–8M urea or guanidine-HCl) conditions. The presence of serum in mammalian and insect cell culture medium does not interfere with purification. For more information and a detailed protocol, see Technical Manual #TM060 and the MagneHis™ Protein Purification System Automated Protocol.

Figure 2. Diagram of the MagneHis™ Protein Purification System protocol.

Protocol: MagneHis™ Purification of Proteins Expressed in Bacterial Cells

Materials Required:
    and protocol
  • 37°C incubator for flasks or tubes
  • shaker
  • magnetic separation stand
  • 1M imidazole solution (pH 8.0 for purification from insect or mammalian cells or culture medium)
  • additional binding/wash buffer (may be required if processing numerous insect cell, mammalian cell or culture medium samples)
  • solid NaCl (for purification from insect or mammalian cells or culture medium)

Purification using Denaturing Conditions. Proteins expressed in bacterial cells may be present in insoluble inclusion bodies. To determine if your protein is located in an inclusion body, perform the lysis step using FastBreak™ Cell Lysis Reagent, 10X, as described in Technical Manual #TM060. Pellet cellular debris by centrifugation, and check the supernatant and pellet for the polyhistidine-tagged protein by gel analysis.

Efficient purification of insoluble proteins requires denaturing conditions. Since the interaction of polyhistidine-tagged fusion proteins and MagneHis™ Ni-Particles does not depend on tertiary structure, fusion proteins can be captured and purified using denaturing conditions by adding a strong denaturant such as 2–8M guanidine hydrochloride or urea to the cells. Denaturing conditions must be used throughout the procedure so that the proteins do not aggregate. We recommend preparing denaturing buffers by adding solid guanidine-HCl or urea directly to the MagneHis™ Binding/Wash and Elution Buffers. For more information, see Technical Manual #TM060.

Note: Do not combine FastBreak™ Cell Lysis Reagent and denaturants. Cells can be lysed directly using denaturants such as urea or guanidine-HCl.

Purification from Insect and Mammalian Cells. Process cells at a cell density of 2 × 10 6 cells/ml of culture. Adherent cells may be removed from the tissue culture vessel by scraping and resuspending in culture medium to this density. Cells may be processed in culture medium containing up to 10% serum. Processing more than the indicated number of cells per milliliter of sample may result in reduced protein yield and increased nonspecific binding. For proteins that are secreted into the cell culture medium, remove any cells from the medium prior to purification. For more information, see Technical Manual #TM060.

MagneHis™ System Protocols

More information and detailed protocols for use of the MagneHis™ System are available in Technical Manual #TM060. A protocol for automating MagneHis™ system on liquid handlers is also available (#EP011).

Protocol: MagZ&trade Purification of Proteins Expressed in Reticulocyte Lysate

Purification of a polyhistidine-tagged protein that is expressed in rabbit reticulocyte lysate is complicated by copurification of hemoglobin in the lysate and the protein of interest. Hemoglobin copurification limits downstream applications (e.g., fluorescence-based functional assays, protein:protein interaction studies) and reduces the amount of protein purified. The MagZ&trade Protein Purification System provides a simple, rapid and reliable method to purify expressed polyhistidine-tagged protein from rabbit reticulocyte lysate with minimal copurification of hemoglobin. The paramagnetic, precharged MagZ&trade Binding Particles are used to isolate polyhistidine-tagged protein from 50&ndash500&mul of T n T® Rabbit Reticulocyte Lysate, resulting in polyhistidine-tagged proteins that are 99% free of contaminating hemoglobin.

The MagZ&trade System is flexible enough to be used with different labeling and detection methods. Polyhistidine-tagged proteins expressed in rabbit reticulocyte lysate can be labeled with [35S]methionine or the FluoroTect&trade GreenLys in vitro Translation Labeling System. FluoroTect&trade dye-labeled polyhistidine-tagged proteins can be visualized by gel analysis and analyzed using a FluorImager® instrument. Figure 3 shows a schematic diagram of the MagZ&trade Protein Purification System protocol. For more information and a detailed protocol, see Technical Bulletin #TB336.

Materials Required:

Figure 3. Schematic diagram of the MagZ™ Protein Purification System. A TnT® reaction expressing polyhistidine-tagged proteins is diluted with MagZ™ Binding/Wash Buffer and added to MagZ™ Particles. The polyhistidine-tagged proteins bind to the particles during incubation and then are washed to remove unbound and nonspecifically bound proteins.

MagZ™ System Protocol

More information and a detailed protocol for use of the MagZ&trade System are available in Technical Manual #TM336.

Medium- to Large-Scale Purification of Polyhistidine-Tagged Proteins In Column or Batch Formats

The two most common support materials for resin-based, affinity-tagged protein purification are agarose and silica gel. As a chromatographic support, silica is advantageous because it has a rigid mechanical structure that is not vulnerable to swelling and can withstand large changes in pressure and flow rate without disintegrating or deforming. Silica is available in a wide range of pore and particle sizes including macroporous silica, which provides a higher capacity for large biomolecules such as proteins. However, two of the drawbacks of silica as a solid support for affinity purification are the limited reagent chemistry that is available and the relatively low efficiency of surface modification.

The HisLink™ Protein Purification Resin (Cat.# V8821, V8823) overcomes these limitations by using a new modification process for silica surfaces that provides a tetradentate metal-chelated solid support with a high binding capacity and concomitantly eliminates the nonspecific binding that is characteristic of unmodified silica. HisLink™ Resin is a macroporous silica resin modified to contain a high level of tetradentate-chelated nickel (>20mmol Ni/ml settled resin). Figure 4 show a schematic diagram of HisLink™ Resin and polyhistidine tag interaction. The HisLink™ Resin has a pore size that results in binding capacities as high as 35mg of polyhistidine-tagged protein per milliliter of resin.

The HisLink™ Resin enables efficient capture and purification of bacterially expressed polyhistidine-tagged proteins. This resin also may be used for general applications that require an immobilized metal affinity chromatography (IMAC) matrix (Porath et al. 1975 Lonnerdal and Keen, 1982). HisLink™ Resin may be used in either column or batch purification formats. For a detailed protocol, see Technical Bulletin #TB327.

Figure 4. Schematic diagram of HisLink™ Resin and polyhistidine interaction. Two sites are available for polyhistidine-tag binding and are rapidly coordinated with histidine in the presence of a polyhistidine-tagged polypeptide.

Column-Based Purification using HisLink™ Resin

The HisLink™ Resin provides a conventional means to purify polyhistidine-tagged proteins and requires only a column that can be packed to the appropriate bed volume. When packed to 1ml under gravity-driven flow, HisLink™ Resin shows an average flow rate of approximately 1ml/minute. In general a flow rate of 1–2ml/minute per milliliter of resin is optimal for efficient capture of polyhistidine-tagged protein. Gravity flow of a cleared lysate over a HisLink™ column will result in complete capture and efficient elution of polyhistidine-tagged proteins however, the resin also may be used with vacuum filtration devices (e.g., Vac-Man® Vacuum Manifold, Cat.# A7231) to allow simultaneous processing of multiple columns. HisLink™ Resin is also an excellent choice for affinity purification using low- to medium-pressure liquid chromatography systems such as fast performance liquid chromatography (FPLC).

Example Protocol Using the HisLink™ Resin to Purify Proteins from Cleared Lysate by Gravity-Flow Column Chromatography

Materials Required:

  • HisLink™ Protein Purification Resin (Cat.# V8821) and protocol
  • HEPES buffer (pH 7.5)
  • imidazole
  • HisLink™ Binding Buffer
  • HisLink™ Wash Buffer
  • HisLink™ Elution Buffer
  • column

Cell Lysis: Cells may be lysed using any number of methods including sonication, French press, bead milling, treatment with lytic enzymes (e.g., lysozyme) or use of a commercially available cell lysis reagent such as the FastBreak™ Cell Lysis Reagent (Cat.# V8571). If lysozyme is used to prepare a lysate, add salt (>300mM NaCl) to the binding and wash buffers to prevent lysozyme binding to the resin. Adding protease inhibitors such as 1mM PMSF to cell lysates does not inhibit binding or elution of polyhistidine-tagged proteins with the HisLink™ Resin and is highly recommended to prevent degradation of the protein of interest by endogenous proteases. When preparing cell lysates from high-density cultures, adding DNase and RNase (concentrations up to 20μg/ml) will reduce the lysate viscosity and aid purification.

  1. Prepare the HisLink™ Binding, Wash and Elution Buffers
    Note: Polyhistidine-tagged proteins can be eluted using 250–1,000mM imidazole. Polyhistidine tags containing less than six histidines typically require less imidazole for elution, while polyhistidine proteins containing more than six polyhistidines may require higher levels of imidazole.
  2. Determine the column volume required to purify the protein of interest. In most cases 1ml of settled resin is sufficient to purify the amount of protein typically found in up to 1 liter of culture (cell density of O.D.600 < 6.0). In cases of very high expression levels (e.g., 50mg protein/liter), up to 2ml of resin per liter of culture may be needed.
  3. Once you have determined the volume of settled resin required, precalibrate this amount directly in the column by pipetting the equivalent volume of water into the column and marking the column to indicate the top of the water. This mark indicates the top of the settled resin bed. Remove the water before adding resin to the column.
  4. Make sure that the resin is fully suspended fill the column with resin to the line marked on the column by transferring the resin with a pipette. Allow the resin to settle, and adjust the level of the resin by adding or removing resin as necessary.
    Note: If the resin is not pipetted within 10–15 seconds of mixing, significant settling will occur, and the resin will need to be resuspended. Alternatively, a magnetic stir bar may be used to keep the resin in suspension during transfer. To avoid fracturing the resin, do not leave the resin stirring any longer than the time required to pipet and transfer the resin.
  5. Allow the column to drain, and equilibrate the resin with five column volumes of binding buffer, allowing the buffer to completely enter the resin bed.
  6. Gently add the cleared lysate to the resin until the lysate has completely entered the column. The rate of flow through the column should not exceed 1–2ml/minute for every 1ml of column volume. Under normal gravity flow conditions the rate is typically about 1ml/minute. The actual flow rate will depend on the type of column used and the extent to which the lysate was cleared and filtered. Do not let the resin dry out after you have applied the lysate to the column.
  7. Wash unbound proteins from the resin using at least 10–20 column volumes of wash buffer. Divide the total volume of wash buffer into two or three aliquots, and allow each aliquot to completely enter the resin bed before adding the next aliquot.
  8. Once the wash buffer has completely entered the resin bed, add elution buffer and begin collecting fractions (0.5–5ml fractions). Elution profiles are protein-dependent, but polyhistidine-tagged proteins will generally elute in the first 1ml. Elution is usually complete after 3–5ml of buffer is collected per 1.0ml of settled resin, provided the imidazole concentration is high enough to efficiently elute the protein of interest.

Batch Protein Purification Using HisLink™ Resin

One of the primary advantages of the HisLink™ Resin is its use in batch purification. In batch mode, the protein of interest is bound to the resin by mixing lysate with the resin for approximately 30 minutes at a temperature range of 4–22°C. Once bound with protein, the resin is allowed to settle to the bottom of the container, and the spent lysate is removed. Washing requires only resuspension of the resin in an appropriate wash buffer followed by a brief period to allow the resin to settle. The wash buffer is then carefully poured off. This process is repeated as many times as desired. Final elution is best achieved by transferring the HisLink™ Resin to a column to elute the protein in fractions. The advantages of batch purification are: 1) less time is required to perform the purification 2) large amounts of lysate can be processed and 3) clearing the lysate prior to purification is not required.

Purification of Polyhistidine-Tagged Proteins by FPLC

The rigid particle structure of the silica base used in the HisLink™ Resin make this material an excellent choice for applications that require applied pressure to load the lysate, wash or elute protein from the resin. These applications involve both manual and automated systems that operate under positive or negative pressure (e.g., FPLC and vacuum systems, respectively). To demonstrate the use of HisLink™ Resin on an automated platform we used an AKTA explorer from GE Healthcare to purify milligram quantities of polyhistidine-tagged protein from 1 liter of culture. The culture was lysed in 20ml of binding/wash buffer and loaded onto a column containing 1ml of HisLink™ Resin. We estimate the total amount of protein recovered to be 75–90% of the protein expressed in the original lysate.

Protein purification under denaturing conditions: Proteins that are expressed as an inclusion body and have been solubilized with chaotrophic agents such as guanidine-HCl or urea can be purified by modifying the protocol to include the appropriate amount of denaturant (up to 6M guanidine-HCl or up to 8M urea) in the binding, wash and elution buffers.

HisLink™ Resin may be used in either column or batch purification formats. For a detailed protocol, see Technical Bulletin #TB327.


RAP tag: A new protein purification approach

Whether it's our diets, building strength, or as part of medical advancements, it is no secret that proteins form an important part of our lives. Tracking how proteins work and move in cells, and purifying engineered proteins, are important tools for researchers. Traditional approaches to label proteins of interest, called "tagging," have the disadvantage of interfering with protein characteristics, including function and localization. Sometimes, these tags can also cross-react, which makes the information they provide nonspecific. A successful protein tagging system needs to be highly specific and have high affinity.

In a study published in September 2020 in Frontiers in Plant Science, researchers from the University of Tsukuba, led by Professor Kenji Miura, have described a new tagging system for detecting and purifying proteins in plant cells. This approach uses a short sequence called a "RAP tag" to label proteins. An antibody, PMab-2, is then able to specifically recognize the RAP tag and can be used to purify the proteins of interest.

In describing this approach, Professor Miura says, "The high affinity and specificity of immunoaffinity chromatography using monoclonal antibodies makes it a very powerful tool, especially for the purification of proteins expressed at low levels." A hurdle to applying this approach, however, is the high cost of reagents, especially that of antibodies.

To get around this, Professor Miura and colleagues explored whether they could produce the PMab-2 antibody in the plant model Nicotiana benthamiana, a relative of the tobacco plant. Not only could they successfully produce PMab-2, they went on to show that the plant-produced PMab-2 behaved similarly to that produced in animal cells. This discovery opens the door to reducing the cost of antibody production, and could be applied more widely across scientific fields.

Testing the feasibility of a RAP-tagged/ PMab-2 affinity purification approach, the researchers then expressed RAP-tagged proteins in plant cells. They found that these tagged proteins could be specifically identified using the PMab-2 antibody. Moreover, RAP-tagged recombinant proteins, involving the fusion of sequences from more than one protein, and protein complexes were also expressed in these cells and identified by PMab-2. These proteins could also be purified from plant cells using the PMab-2 antibody, indicating that the RAP tag can be used for both protein detection and purification from soluble plant extracts.

"Plants are an extremely valuable resource for molecular biology," explains Professor Miura. "They can be used as bioreactors to produce large amounts of proteins because they are unlikely to suffer from contamination issues faced by bacterial and mammalian cell systems."

The results presented by the team show that this approach has the potential to be widely applied across the molecular sciences.


4.1: Protein Purification - Biology

Bacterial Protein Extraction - mini scale - Sonication

1) Resuspend pellet of 10ml cell culture in 1ml lysis buffer (or 100ml bacterial culture for very low expression level).

Suggested Lysis buffer : 140mM NaCl 2.7mM KCL 10mM Na2HPO4 1.8mM KH2PO4 pH 7.3 ( PBS)
or 100mM NaCl 25mM TrisHCl pH 8.0
optional 0.02% NaN3 (azide)
optional protease inhibitors

Optional additives to the lysis buffer
a) 1mM PMSF or protease inhibitor cocktail 1:200 (cocktail for bacterial cells #P-8849 from Sigma)
b) Dnase 100U/ml or 25-50ug/ml (SIGMA DN-25). Incubate 10min 4°C in the presence of 10mMMgCl2.
c) Lysozime 0.2mg/ml. Incubate 10min 4°C.
d) ßME, DTT or DTE up to 10mM for proteins with many cysteines.
e) 0.1-2% Triton X-100, NP40 or any other detergent that do not affect the biological activity of your protein.
f) 10% glycerol (for stabilization of the protein and prevention of aggregation).

2) Sonicate in ice bucket 3 x 10sec or more if the cells are not completely disrupted (Lysis is complete when the cloudy cell suspension becomes translucent. Avoid protein denaturation by frothing).

3) Spin 5min 13000rpm 4°C . Separate soluble proteins (supernatant) from insoluble or inclusion bodies proteins (pellet). Use supernatant for next step. Keep sample of 40ul of supernatant for PAGE-SDS and Western blot: soluble proteins

4) Resuspend pellet in another 1ml lysis buffer and keep sample of 40ul for PAGE-SDS and Western blot: insoluble proteins, or unlysed cells .


Drug Discovery Technologies

3.19.7.3 Capture Step with Affinity Tags

In the last decade, a broad variety of fusion proteins or peptides for simplified protein purification and detection have been described. The fusion partner is encoded on the plasmid or viral DNA and either C- or N-terminally linked to the protein of interest. For larger fusion tags the introduction of an appropriate protease cleavage site between fusion protein and the protein of interest is considered to remove the tag prior to crystallization. Careful removal of the protease before starting crystallization experiments is very important so as to avoid cleavage of the protein of interest during the crystallization procedure.

Plasmids containing DNA coding for proteins like glutathione-S-transferase, 58,59 maltose-binding protein, NusA, calmodulin-binding peptide, intein, thioredoxin, cellulose-binding protein, and Fc fragments of antibodies 17 are commercially available, together with the appropriate protein purification tools. Many of these large fusion partners increase the solubility of the expressed fusion protein (see, for example, Kapust and Waugh 60 ). Sometimes fusion partners simulate solubility of the expressed protein. After removal of the tag, the protein of interest precipitates irreversibly.

Small affinity tags such as Poly-His-tag and Strep-tag 61,62 are widely used as they do not change protein properties and do not need to be cleaved off. When using Poly-His-tags for protein purification, it has to be taken into account that E. coli strains produce protein SlyD (20 kDa) containing a potential metal-binding site domain. This protein binds tightly to metal chelate chromatography and is often detected as a contamination band. 63

Affinity tag chromatography can be performed in batch mode, by centrifugal or syringe filter devices, or on chromatographic equipment with prepacked columns according to the manufacturer's instructions. 58,62,64

In some cases, removal of the elution agent is necessary to maintain protein stability: Fc-fragment fusion proteins are eluted in extreme acid conditions: pH has to be adjusted to neutral values immediately after elution. Using imidazole in high concentrations in the elution of Poly-His-tagged proteins destabilizes proteins and it has to be removed by dialysis.


Contents

Protein purification is either preparative or analytical. Preparative purifications aim to produce a relatively large quantity of purified proteins for subsequent use. Examples include the preparation of commercial products such as enzymes (e.g. lactase), nutritional proteins (e.g. soy protein isolate), and certain biopharmaceuticals (e.g. insulin). Several preparative purifications steps are often deployed to remove bi-products, such as host cell proteins, which poses as a potential threat to the patient's health. [1] Analytical purification produces a relatively small amount of a protein for a variety of research or analytical purposes, including identification, quantification, and studies of the protein's structure, post-translational modifications and function. Pepsin and urease were the first proteins purified to the point that they could be crystallized. [2]

Extraction Edit

If the protein of interest is not secreted by the organism into the surrounding solution, the first step of each purification process is the disruption of the cells containing the protein. Depending on how fragile the protein is and how stable the cells are, one could, for instance, use one of the following methods: i) repeated freezing and thawing, ii) sonication, iii) homogenization by high pressure (French press), iv) homogenization by grinding (bead mill), and v) permeabilization by detergents (e.g. Triton X-100) and/or enzymes (e.g. lysozyme). [3] Finally, the cell debris can be removed by centrifugation so that the proteins and other soluble compounds remain in the supernatant.

Also proteases are released during cell lysis, which will start digesting the proteins in the solution. If the protein of interest is sensitive to proteolysis, it is recommended to proceed quickly, and to keep the extract cooled, to slow down the digestion. Alternatively, one or more protease inhibitors can be added to the lysis buffer immediately before cell disruption. Sometimes it is also necessary to add DNAse in order to reduce the viscosity of the cell lysate caused by a high DNA content.

Precipitation and differential solubilization Edit

In bulk protein purification, a common first step to isolate proteins is precipitation with ammonium sulfate (NH4)2SO4. [4] This is performed by adding increasing amounts of ammonium sulfate and collecting the different fractions of precipitated protein. Subsequently, ammonium sulfate can be removed using dialysis. During the ammonium sulfate precipitation step, hydrophobic groups present on the proteins are exposed to the atmosphere, attracting other hydrophobic groups the result is formation of an aggregate of hydrophobic components. In this case, the protein precipitate will typically be visible to the naked eye. One advantage of this method is that it can be performed inexpensively, even with very large volumes.

The first proteins to be purified are water-soluble proteins. Purification of integral membrane proteins requires disruption of the cell membrane in order to isolate any one particular protein from others that are in the same membrane compartment. Sometimes a particular membrane fraction can be isolated first, such as isolating mitochondria from cells before purifying a protein located in a mitochondrial membrane. A detergent such as sodium dodecyl sulfate (SDS) can be used to dissolve cell membranes and keep membrane proteins in solution during purification however, because SDS causes denaturation, milder detergents such as Triton X-100 or CHAPS can be used to retain the protein's native conformation during complete purification.

Ultracentrifugation Edit

Centrifugation is a process that uses centrifugal force to separate mixtures of particles of varying masses or densities suspended in a liquid. When a vessel (typically a tube or bottle) containing a mixture of proteins or other particulate matter, such as bacterial cells, is rotated at high speeds, the inertia of each particle yields a force in the direction of the particles velocity that is proportional to its mass. The tendency of a given particle to move through the liquid because of this force is offset by the resistance the liquid exerts on the particle. The net effect of "spinning" the sample in a centrifuge is that massive, small, and dense particles move outward faster than less massive particles or particles with more "drag" in the liquid. When suspensions of particles are "spun" in a centrifuge, a "pellet" may form at the bottom of the vessel that is enriched for the most massive particles with low drag in the liquid.

Non-compacted particles remain mostly in the liquid called "supernatant" and can be removed from the vessel thereby separating the supernatant from the pellet. The rate of centrifugation is determined by the angular acceleration applied to the sample, typically measured in comparison to the g. If samples are centrifuged long enough, the particles in the vessel will reach equilibrium wherein the particles accumulate specifically at a point in the vessel where their buoyant density is balanced with centrifugal force. Such an "equilibrium" centrifugation can allow extensive purification of a given particle.

Sucrose gradient centrifugation — a linear concentration gradient of sugar (typically sucrose, glycerol, or a silica based density gradient media, like Percoll) is generated in a tube such that the highest concentration is on the bottom and lowest on top. Percoll is a trademark owned by GE Healthcare companies. A protein sample is then layered on top of the gradient and spun at high speeds in an ultracentrifuge. This causes heavy macromolecules to migrate towards the bottom of the tube faster than lighter material. During centrifugation in the absence of sucrose, as particles move farther and farther from the center of rotation, they experience more and more centrifugal force (the further they move, the faster they move). The problem with this is that the useful separation range of within the vessel is restricted to a small observable window. Spinning a sample twice as long doesn't mean the particle of interest will go twice as far, in fact, it will go significantly further. However, when the proteins are moving through a sucrose gradient, they encounter liquid of increasing density and viscosity. A properly designed sucrose gradient will counteract the increasing centrifugal force so the particles move in close proportion to the time they have been in the centrifugal field. Samples separated by these gradients are referred to as "rate zonal" centrifugations. After separating the protein/particles, the gradient is then fractionated and collected.

Choice of a starting material is key to the design of a purification process. In a plant or animal, a particular protein usually isn't distributed homogeneously throughout the body different organs or tissues have higher or lower concentrations of the protein. Use of only the tissues or organs with the highest concentration decreases the volumes needed to produce a given amount of purified protein. If the protein is present in low abundance, or if it has a high value, scientists may use recombinant DNA technology to develop cells that will produce large quantities of the desired protein (this is known as an expression system). Recombinant expression allows the protein to be tagged, e.g. by a His-tag [5] or Strep-tag [6] to facilitate purification, reducing the number of purification steps required.

An analytical purification generally utilizes three properties to separate proteins. First, proteins may be purified according to their isoelectric points by running them through a pH graded gel or an ion exchange column. Second, proteins can be separated according to their size or molecular weight via size exclusion chromatography or by SDS-PAGE (sodium dodecyl sulfate-polyacrylamide gel electrophoresis) analysis. Proteins are often purified by using 2D-PAGE and are then analysed by peptide mass fingerprinting to establish the protein identity. This is very useful for scientific purposes and the detection limits for protein are nowadays very low and nanogram amounts of protein are sufficient for their analysis. Thirdly, proteins may be separated by polarity/hydrophobicity via high performance liquid chromatography or reversed-phase chromatography.

Usually a protein purification protocol contains one or more chromatographic steps. The basic procedure in chromatography is to flow the solution containing the protein through a column packed with various materials. Different proteins interact differently with the column material, and can thus be separated by the time required to pass the column, or the conditions required to elute the protein from the column. Usually proteins are detected as they are coming off the column by their absorbance at 280 nm. Many different chromatographic methods exist:

Size exclusion chromatography Edit

Chromatography can be used to separate protein in solution or denaturing conditions by using porous gels. This technique is known as size exclusion chromatography. The principle is that smaller molecules have to traverse a larger volume in a porous matrix. Consequentially, proteins of a certain range in size will require a variable volume of eluent (solvent) before being collected at the other end of the column of gel.

In the context of protein purification, the eluent is usually pooled in different test tubes. All test tubes containing no measurable trace of the protein to purify are discarded. The remaining solution is thus made of the protein to purify and any other similarly-sized proteins.

Separation based on charge or hydrophobicity Edit

Hydrophobic interaction chromatography Edit

HIC media is amphiphilic, with both hydrophobic and hydrophilic regions, allowing for separation of proteins based on their surface hydrophobicity. Target proteins and their product aggregate species tend to have different hydrophobic properties and removing them via HIC further purifies the protein of interest. [7] Additionally, the environment used typically employs less harsh denaturing conditions than other chromatography techniques, thus helping to preserve the protein of interest in its native and functional state. In pure water, the interactions between the resin and the hydrophobic regions of protein would be very weak, but this interaction is enhanced by applying a protein sample to HIC resin in high ionic strength buffer. The ionic strength of the buffer is then reduced to elute proteins in order of decreasing hydrophobicity. [8]

Ion exchange chromatography Edit

Ion exchange chromatography separates compounds according to the nature and degree of their ionic charge. The column to be used is selected according to its type and strength of charge. Anion exchange resins have a positive charge and are used to retain and separate negatively charged compounds (anions), while cation exchange resins have a negative charge and are used to separate positively charged molecules (cations).

Before the separation begins a buffer is pumped through the column to equilibrate the opposing charged ions. Upon injection of the sample, solute molecules will exchange with the buffer ions as each competes for the binding sites on the resin. The length of retention for each solute depends upon the strength of its charge. The most weakly charged compounds will elute first, followed by those with successively stronger charges. Because of the nature of the separating mechanism, pH, buffer type, buffer concentration, and temperature all play important roles in controlling the separation.

Ion exchange chromatography is a very powerful tool for use in protein purification and is frequently used in both analytical and preparative separations.

Free-flow-electrophoresis Edit

Free-flow electrophoresis (FFE) is a carrier-free electrophoresis technique that allows preparative protein separation in a laminar buffer stream by using an orthogonal electric field. By making use of a pH-gradient, that can for example be induced by ampholytes, this technique allows to separate protein isoforms up to a resolution of < 0.02 delta-pI.

Affinity chromatography Edit

Affinity Chromatography is a separation technique based upon molecular conformation, which frequently utilizes application specific resins. These resins have ligands attached to their surfaces which are specific for the compounds to be separated. Most frequently, these ligands function in a fashion similar to that of antibody-antigen interactions. This "lock and key" fit between the ligand and its target compound makes it highly specific, frequently generating a single peak, while all else in the sample is unretained.

Many membrane proteins are glycoproteins and can be purified by lectin affinity chromatography. Detergent-solubilized proteins can be allowed to bind to a chromatography resin that has been modified to have a covalently attached lectin. Proteins that do not bind to the lectin are washed away and then specifically bound glycoproteins can be eluted by adding a high concentration of a sugar that competes with the bound glycoproteins at the lectin binding site. Some lectins have high affinity binding to oligosaccharides of glycoproteins that is hard to compete with sugars, and bound glycoproteins need to be released by denaturing the lectin.

Immunoaffinity chromatography Edit

Immunoaffinity chromatography uses the specific binding of an antibody-antigen to selectively purify the target protein. The procedure involves immobilizing a protein to a solid substrate (e.g. a porous bead or a membrane), which then selectively binds the target, while everything else flows through. The target protein can be eluted by changing the pH or the salinity. The immobilized ligand can be an antibody (such as Immunoglobulin G) or it can be a protein (such as Protein A). Because this method does not involve engineering in a tag, it can be used for proteins from natural sources. [9]

Purification of a tagged protein Edit

Another way to tag proteins is to engineer an antigen peptide tag onto the protein, and then purify the protein on a column or by incubating with a loose resin that is coated with an immobilized antibody. This particular procedure is known as immunoprecipitation. Immunoprecipitation is quite capable of generating an extremely specific interaction which usually results in binding only the desired protein. The purified tagged proteins can then easily be separated from the other proteins in solution and later eluted back into clean solution.

When the tags are not needed anymore, they can be cleaved off by a protease. This often involves engineering a protease cleavage site between the tag and the protein.

HPLC Edit

High performance liquid chromatography or high pressure liquid chromatography is a form of chromatography applying high pressure to drive the solutes through the column faster. This means that the diffusion is limited and the resolution is improved. The most common form is "reversed phase" HPLC, where the column material is hydrophobic. The proteins are eluted by a gradient of increasing amounts of an organic solvent, such as acetonitrile. The proteins elute according to their hydrophobicity. After purification by HPLC the protein is in a solution that only contains volatile compounds, and can easily be lyophilized. [10] HPLC purification frequently results in denaturation of the purified proteins and is thus not applicable to proteins that do not spontaneously refold.

At the end of a protein purification, the protein often has to be concentrated. Different methods exist.

Lyophilization Edit

If the solution doesn't contain any other soluble component than the protein in question the protein can be lyophilized (dried). This is commonly done after an HPLC run. This simply removes all volatile components, leaving the proteins behind.

Ultrafiltration Edit

Ultrafiltration concentrates a protein solution using selective permeable membranes. The function of the membrane is to let the water and small molecules pass through while retaining the protein. The solution is forced against the membrane by mechanical pump, gas pressure, or centrifugation.

The most general method to monitor the purification process is by running a SDS-PAGE of the different steps. This method only gives a rough measure of the amounts of different proteins in the mixture, and it is not able to distinguish between proteins with similar apparent molecular weight.

If the protein has a distinguishing spectroscopic feature or an enzymatic activity, this property can be used to detect and quantify the specific protein, and thus to select the fractions of the separation, that contains the protein. If antibodies against the protein are available then western blotting and ELISA can specifically detect and quantify the amount of desired protein. Some proteins function as receptors and can be detected during purification steps by a ligand binding assay, often using a radioactive ligand.

In order to evaluate the process of multistep purification, the amount of the specific protein has to be compared to the amount of total protein. The latter can be determined by the Bradford total protein assay or by absorbance of light at 280 nm, however some reagents used during the purification process may interfere with the quantification. For example, imidazole (commonly used for purification of polyhistidine-tagged recombinant proteins) is an amino acid analogue and at low concentrations will interfere with the bicinchoninic acid (BCA) assay for total protein quantification. Impurities in low-grade imidazole will also absorb at 280 nm, resulting in an inaccurate reading of protein concentration from UV absorbance.

Another method to be considered is Surface Plasmon Resonance (SPR). SPR can detect binding of label free molecules on the surface of a chip. If the desired protein is an antibody, binding can be translated directly to the activity of the protein. One can express the active concentration of the protein as the percent of the total protein. SPR can be a powerful method for quickly determining protein activity and overall yield. It is a powerful technology that requires an instrument to perform.

Denaturing-condition electrophoresis Edit

Gel electrophoresis is a common laboratory technique that can be used both as preparative and analytical method. The principle of electrophoresis relies on the movement of a charged ion in an electric field. In practice, the proteins are denatured in a solution containing a detergent (SDS). In these conditions, the proteins are unfolded and coated with negatively charged detergent molecules. The proteins in SDS-PAGE are separated on the sole basis of their size.

In analytical methods, the protein migrate as bands based on size. Each band can be detected using stains such as Coomassie blue dye or silver stain. Preparative methods to purify large amounts of protein, require the extraction of the protein from the electrophoretic gel. This extraction may involve excision of the gel containing a band, or eluting the band directly off the gel as it runs off the end of the gel.

In the context of a purification strategy, denaturing condition electrophoresis provides an improved resolution over size exclusion chromatography, but does not scale to large quantity of proteins in a sample as well as the late chromatography columns.

Non-denaturing-condition electrophoresis Edit

A non-denaturing electrophoretic procedure for isolating bioactive metalloproteins in complex protein mixtures is preparative native PAGE. The intactness or the structural integrity of the isolated protein has to be confirmed by an independent method. [11]


Description

Guide to Protein Purification, Second Edition provides a complete update to existing methods in the field, reflecting the enormous advances made in the last two decades. In particular, proteomics, mass spectrometry, and DNA technology have revolutionized the field since the first edition’s publication but through all of the advancements, the purification of proteins is still an indispensable first step in understanding their function. This volume examines the most reliable, robust methods for researchers in biochemistry, molecular and cell biology, genetics, pharmacology and biotechnology and sets a standard for best practices in the field. It relates how these traditional and new cutting-edge methods connect to the explosive advancements in the field. This "Guide to" gives imminently practical advice to avoid costly mistakes in choosing a method and brings in perspective from the premier researchers while presents a comprehensive overview of the field today.


Watch the video: Gel Filtration Sephadex G 50 (December 2022).