In biochemistry, a protein is a long chain of amino acids that are all linked together. Proteins are an important part of how cells are built and how they work.
But if you want to study proteins, you need to find and separate them first. Biochemists do something called “high-throughput protein purification” to make this happen.
First, each protein is taken out on its own, and then they are put into groups based on their molecular weight, solubility, charge, and ability to bind to specific sites.
This process helps scientists study how proteins are made and how they interact with each other.
Protein purification is a series of steps that are done to get the protein you want out of a complex mixture so you can study it. Getting a protein to itself will help scientists figure out:
Size and structure
Affinity for binding
Activity in living things
Physical and chemical traits
Proteins talk to each other
This process has a lot of uses in research, especially for making recombinant proteins on a large scale. (However, it can be hard to purify native proteins. When this happens, affinity tags like Polyhistidine, Glutathione-S-transferase, and His-tags are used to separate and bind them.)
Protein Purification Methods
Creating a Crude Protein Extract – Extraction
A single-step purification process can’t be used to get a pure plant protein because plant tissues have many different kinds of proteins, a tough cell wall made of cellulosic material, and phenolic compounds that can break down proteins.
The process of extracting starts with putting a fresh or frozen plant sample of the right weight under liquid N2.
Mix this all together in a cold mortar with the right amount of extraction buffer (3 x of the sample).
The type of extraction buffer and its pH are unique to the type of sample. To improve the quality of the extract that is made, some additives should be added to the extraction buffer. Among these additives are:
To stop protease activity, phenylmethylsulfonyl fluoride (PMSF) is dissolved in a very small amount of propanol right before extraction.
Compounds with flavin (FAD)
Dithiothreitol (DTT), to keep the sulfhydryl group from breaking down.
As a chemical chelator, EDTA works best with phosphate buffer.
Sodium fluoride can stop phosphatase from working.
Polyphenylpyrovate (PVP) 25 gm/k g fresh weight of sample, which is an insoluble compound that binds to phenolic compounds in the sample. The resulting compound is thrown away by centrifugation.
30% glycerol, which may help some very unstable proteins stay in place.
Antibiotics like Hibitane are often used to get protein from rhizomes or other underground plant parts.
Use nylon mush to filter the homogenate, and then use a cooling centrifuge to spin it at 10,000 g for 10 minutes.
Repeat the process of extracting and spinning the tissue three times to make sure that all of the proteins are taken out.
Almost the same method was used to get Rubisco out of wheat leaves, protein out of Peganum harmala seeds, and protein out of chickpea seeds.
In the crude extracts, the total amount of protein and the activity should be measured. If the target protein is an enzyme, the activity should also be measured.
A Yeast Protein Extraction Buffer Kit could be used for a microorganism like yeast.
This buffer is made from organic buffering agents, which use mild nonionic detergents and a secret mix of different salts and agents to make it easier to get proteins out of the solution and keep them stable.
There is also a Zymolyase preparation that is ready to use.
Precipitation and Differential Solubilisation – Concentration of crude extract
In bulk protein purification, ammonium sulphate (NH4)2SO4 is often used as the first step to separate proteins.
To do this, you add more and more ammonium sulphate and collect the different amounts of protein that precipitate.
One good thing about this method is that it can be done cheaply and in very large amounts.
Water-soluble proteins are the first ones to be cleaned up. Integral membrane proteins can’t be separated from other proteins in the same membrane compartment without breaking the cell membrane.
Sometimes, a certain membrane tract can be isolated first. For example, mitochondria can be separated from cells before a protein in a mitochondrial membrane is purified.
Sodium dodecyl sulphate (SDS) is a detergent that can be used to break down cell membranes and keep membrane proteins in solution during purification. However, because SDS denatures proteins, milder detergents like Triton X-100 or CHAPS can be used to keep the proteins in their natural shape during purification.
Ammonium sulfate precipitation
Solid ammonium sulphate was added to the crude extracts to bring the final concentration to 70% (w/v) or look at Table 1 to choose a different concentration. This was done to concentrate or reduce the total amount of crude extract.
After the solid ammonium sulphate was fully dissolved, the mixture was left to sit at 4°C for 24 hours. The precipitate was then collected by spinning the mixture at 10,000 g for 10 minutes in a cooling centrifuge.
The proteins that had clumped together were dissolved in the least amount of extraction buffer. Dialysis was done against the same buffer to get rid of the extra ammonium ions.
Fractional precipitation with acetone
For this protein fraction, the best range for precipitation is between 37.5% and 50% (v/v).
Cruse protein extract is chilled in an ice-salt bath in the right amount.
Add 0.60 ml of acetone for every 1 ml of protein solution (dropwise with constant stirring).
After the acetone has been added, keep stirring and keeping an eye on the temperature for 10 minutes.
When the acetone-protein mixture is spun at 3000 xg for 10 minutes, the formed precipitate is collected.
The protein that has clumped together is taken out and retrieved using the least amount of extraction buffer.
The amount of supernatant is measured, and 0.25 ml of acetone is added for every ml of protein solution.
As in the last step, centrifugation is used to dissolve the protein that has settled to the bottom. To get rid of any leftover acetone, the centrifuge tubes with filter paper on top are turned upside down.
The pellet is kept for the next step of purification in a small amount of buffer.
Ultracentrifugation
Centrifugation is a process that uses the force of spinning to separate particles of different sizes or densities that are floating in a liquid.
When a tube or bottle with a mixture of proteins or other small particles, like bacterial cells, is rotated quickly, the angular momentum gives each particle an outward force that is proportional to its mass.
Because of this force, a particle might try to move through a liquid. However, the liquid pushes back against the particle.
When the sample is “spun” in a centrifuge, the result is that small, dense, and heavy particles move outward faster than less dense or less heavy particles or particles that “drag” more in the liquid.
When a centrifuge is used to “spin” a suspension of particles, a “pellet” may form at the bottom of the vessel. This pellet is made up of the most dense particles that don’t move much in the liquid.
The liquid with the non-compacted particles that are still mostly in it is called the “supernatant,” and the supernatant can be taken out of the vessel to separate it from the pellet.
The sample’s angular acceleration, which is usually measured in relation to the g, tells us how fast the centrifuge is going.
If samples are spun long enough, the particles in the vessel will reach equilibrium. This means that the particles will gather at a point where their buoyancy and centrifugal force are both equal.
With this kind of “equilibrium” centrifugation, a particle can be cleaned up a lot.
Sucrose gradient centrifugation
In a tube, sugar (usually sucrose glycerol or Percoll) is put in such a way that the highest concentration is at the bottom and the lowest concentration is at the top.
Then, a protein sample is put on top of the gradient, and an ultracentrifuge is used to spin it very fast. This makes it so that the heavier macromolecules move to the bottom of the tube faster than the lighter ones.
When centrifugation is done without sucrose, particles get more and more centrifugal force as they move farther away from the centre of rotation (the further they move, the faster they move).
The problem with this is that the useful range of separation inside the ship is limited to a small window that can be seen.
If you spin a sample twice as long, the particle of interest won’t go twice as far; it will go much farther. But as the proteins move through a sucrose gradient, they meet liquid that is getting more dense and thick.
If the sucrose gradient is set up right, it will work against the increasing centrifugal force, so the particles will move in a way that is close to proportional to how long they have been in the centrifugal field.
“Rate zonal” centrifugations are used to separate samples based on these differences. After separating the protein/particles, the gradient is then divided and collected.
Chromatographic Methods
Most methods for purifying proteins have one or more chromatographic steps.
In chromatography, the basic step is to let the solution with the protein flow through a column full of different materials.
Different proteins interact with the column material in different ways. This means that the time it takes for a protein to pass through the column or the conditions it needs to get out of the column can be used to separate the proteins. Most of the time, the absorbance at 280 nm is used to find proteins as they are coming off the column.
Different chromatographic methods
1. Size Exclusion Chromatography
By using porous gels, chromatography can be used to separate proteins in solution or under conditions that cause them to break down.
Size exclusion chromatography is the name for this method.
In a porous matrix, smaller molecules have to move through a larger volume because they are smaller.
So, proteins in a certain size range will need a different amount of eluant (solvent) before they can be collected at the other end of the gel column.
In the process of purifying proteins, the eluant is usually put into different test tubes and mixed together.
All test tubes with no trace of the protein to be purified are thrown away.
So, the remaining solution is made up of the protein to be purified and any other proteins that are about the same size.
The type and strength of the charge decide which column will be used.
Anion exchange resins have a positive charge and are used to keep and separate negatively charged molecules, while cation exchange resins have a negative charge and are used to separate positively charged molecules.
Before the separation can start, a buffer is pumped through the column to make sure that the oppositely charged ions are all in the same place.
When the sample is injected into the resin, the solute molecules will trade places with the buffer ions as they try to bind to the resin.
The amount of time each solute is kept depends on how strong its charge is.
The compounds with the weakest charges will come out first, then those with stronger charges.
Because of how the separation works, pH, buffer type, buffer concentration, and temperature all play important roles in controlling the separation.
Ion exchange chromatography is a very effective way to get rid of impurities in proteins, and it is often used to separate substances for both analytical and practical purposes.
3. Affinity Chromatography
Affinity chromatography is a way to separate things based on how molecules are shaped. Often, application-specific resins are used.
On the surface of these resins are ligands that are specific to the compounds that need to be separated.
Most of the time, these ligands work in a way that is similar to how antibodies and antigens interact.
This “lock and key” fit between the ligand and its target compound makes it very specific, often producing a single peak while leaving behind everything else in the sample.
Many membrane proteins are glycoproteins, and lectin affinity chromatography can be used to clean them up.
Proteins that have been broken down by detergent can bind to a chromatography resin that has been changed so that a lectin is covalently attached to it.
By introducing a high concentration of a sugar that competes with the bound glycoproteins at the lectin binding site, specifically bound glycoproteins can be eluted.
Some lectins have a hard time competing with sugars when it comes to binding to oligosaccharides of glycoproteins. The bound glycoproteins need to be released by denaturing the lectin.
4. Metal Binding
Adding a sequence of 6 to 8 histidines to the C-terminus of the protein is a common method.
Divalent metal ions, like nickel and cobalt, stick strongly to the polyhistidine.
The polyhistidine tag can be attached to the protein by passing it through a column with nickel ions that are stuck together.
All proteins that don’t have tags pass through the column.
The protein can be released from the column by using imidazole, which prevents the polyhistidine tag from binding to the column, or by lowering the pH (usually to 4.5), which makes the tag less likely to stick to the resin.
This method is usually used to clean up recombinant proteins with an engineered affinity tag, like a 6xHis-tag or Clontech’s HAT tag. However, it can also be used to clean up natural proteins that have a natural affinity for divalent cations.
5. Immunoaffinity
Immunoaffinity chromatography (IAC) is a type of liquid chromatography that uses antibodies to bind to an antigen that represents the target protein in a very specific way.
Immune-extraction, immune-depletion, chromatographic immunoassays, and post-column immune-detection are among the applications of IAC.
Because of how tightly and selectively antigens and antibodies stick to each other, they can be used in all of these different ways.
IAC has been used to separate wild-type and recombinant mutant amidases from Pseudomonas aeruginosa. This was done by using either immobilised antigens or antibodies.
For immunodiagnostics and biopharmaceutics, antibodies are separated by antibody class, and higher levels of purification help both diagnostic methods and therapeutic uses.
Because purifying antibodies is so important, more and more attention is being paid to IAC, which is a good way to separate proteins based on how target proteins interact with specific immobilised antibodies.
Monoclonal antibodies (MAbs) can be used to treat many different conditions, such as cancer, autoimmune diseases, infectious diseases, heart diseases, transplant rejection, and cardiovascular diseases.
In this way, specific antibodies were used to separate the IgG class, which plays the most important role in clinical applications (hyperimmune IgG).
Because purifying antibodies is so important, more and more attention is being paid to IAC, which is a good way to separate proteins based on how target proteins interact with specific immobilised antibodies.
New synthetic ligands have become the focus of research due to their low cost, low risk, and high safety (synthetic mimic ligands of proteins A and L).
By using monospecific antibodies and affinity chromatography, the Mycobacterium tuberculosis H37Ra protein antigens ES-31, ES-43, and EST-6 that can be used for immunodiagnostics were separated from detergent soluble sonicate (DSS) antigen.
Both culture filtrate and DSS antigen purified ES-31, ES-43, and EST-6 antigens showed similar seroreactivity, with an overall sensitivity of 85%, 80%, and 75%, respectively.
6. HPLC
High performance liquid chromatography, also called high pressure liquid chromatography, is a type of chromatography that uses high pressure to move the solutes through the column more quickly.
This means that there is less spreading, so the resolution is better. Most people use “reversed phase” HPLC, in which the material in the column doesn’t like water.
A gradient of increasing amounts of an organic solvent, like acetonitrile, is used to get the proteins out.
The proteins come out based on how much they like water.
After HPLC purification, the protein is in a solution with only volatile compounds. This solution can be easily lyophilized.
HPLC purification often causes the proteins to lose their shape, so it can’t be used on proteins that don’t refold on their own.
References
Structural Genomics Consortium; China Structural Genomics Consortium; Northeast Structural Genomics Consortium, Gräslund S, Nordlund P, Weigelt J, Hallberg BM, Bray J, Gileadi O, Knapp S, Oppermann U, Arrowsmith C, Hui R, Ming J, dhe-Paganon S, Park HW, Savchenko A, Yee A, Edwards A, Vincentelli R, Cambillau C, Kim R, Kim SH, Rao Z, Shi Y, Terwilliger TC, Kim CY, Hung LW, Waldo GS, Peleg Y, Albeck S, Unger T, Dym O, Prilusky J, Sussman JL, Stevens RC, Lesley SA, Wilson IA, Joachimiak A, Collart F, Dementieva I, Donnelly MI, Eschenfeldt WH, Kim Y, Stols L, Wu R, Zhou M, Burley SK, Emtage JS, Sauder JM, Thompson D, Bain K, Luz J, Gheyi T, Zhang F, Atwell S, Almo SC, Bonanno JB, Fiser A, Swaminathan S, Studier FW, Chance MR, Sali A, Acton TB, Xiao R, Zhao L, Ma LC, Hunt JF, Tong L, Cunningham K, Inouye M, Anderson S, Janjua H, Shastry R, Ho CK, Wang D, Wang H, Jiang M, Montelione GT, Stuart DI, Owens RJ, Daenke S, Schütz A, Heinemann U, Yokoyama S, Büssow K, Gunsalus KC. Protein production and purification. Nat Methods. 2008 Feb;5(2):135-46. doi: 10.1038/nmeth.f.202. Erratum in: Nat Methods. 2008 Apr;5(4):369. Hallberg, B Martin [added]. PMID: 18235434; PMCID: PMC3178102.
