Lab 7 - Introduction to Protein Column Chromatography

What is chromatography?

- It's the name given to a broad selection of processes that are designed to separate different molecules from each other

what do you know about the component of chromatography equipment where separation of molecules takes place?

- It is normally comprised of an immobile stationary phase and a moving mobile phase

- the stationary phase usually resides on the outer edges of some kind of rectangular or cylindrical chamber but it can fill the entirety of the chamber

what is the role of the stationary phase?

- It will interact to varying degrees with different types of molecules in the mobile phase that are moving past it

- molecules that interact weakly or not at all with the stationary phase will spend more of their time moving but molecules that interact strongly with the stationary phase will spend a certain portion of the time immobilized

what does the stationary phases interaction with the mobile phase results in?

- Molecules will exit the chamber in which the separation is taking place at different times depending on the particular molecule

what do you know about the stationary phases for protein purification columns?

- They are almost always an inert matrix at which functional groups are covalently attached. The inert matrix is frequently spherical beads made from some kind of polymer such as polystyrene or an agarose based polysaccharide such as Sepharose

- The functional groups give the specificity to the column and can be either charged molecules or molecules modeled after biological molecules to which proteins can interact

what is the buffer or protein mixture of a chromatography column regarded as?

- It can be regarded as the mobile phase. The proteins will separate depending upon the specific properties of the proteins and the stationary phase

what are fractions?

- The different collected samples from a chromatography column are known as fractions. The first fraction collected often only contains the solvent and the second contains the faster moving molecules and the third contains the slower moving molecules

What are the three types of protein column chromatography we will consider?

- Ion exchange or iex, affinity chromatography, and size exclusion chromatography or SEC which is also sometimes known as gel filtration

what is ion exchange chromatography based on?

- It is based upon reversible electrostatic interactions between charged molecules and charged functional groups bound to inert resin beads such as polystyrene or a polysaccharide packed into a column

why is ion exchange chromatography a useful protein purification method?

- Different proteins have different amino acid compositions and therefore different proteins will have different number of positively or negatively charged amino acid side chains at any particular pH

how do functional groups on the IEX resin affect how the column binds different molecules?

- If the functional groups on the IEX resin are positively charged, the column will bind negatively charged proteins, and conversely, if the functional groups are negatively charged, the column will bind molecules with positive charges

how can molecules be selectively bound and unbound from the column?

- By either changing the pH which will affect the charge on the protein, or by changing the salt concentration in the column by increasing or decreasing the concentration of sodium chloride

how would changing the salt concentration in the column affect binding and unbinding from the column?

- Changing the salt conditions can affect the electrostatic interactions because the increase or decrease of the sodium or chloride ions changes the competition between the charged particles for binding spots on charged resin surface

what is protein purification by affinity chromatography based on?

- This specific and reversible high affinity interaction between a protein and a ligand molecule that is covalently linked to an insoluble solid support packed into a column

what is the biggest challenge to using affinity chromatography techniques?

- Finding a ligand that will bind to your protein but not other proteins

how do we get over the biggest challenge to using affinity chromatography techniques?

- Proteins can be synthesized to include sequences of amino acids which are known to bind to ligands that can easily be attached to a column. These additional sequences added to the proteins are called tags of which the most commonly used is the histidine tag

what is a his tag?

- Typically a sequence of six consecutive histidine residues that is attached to the beginning or to the end of a protein sequence

how will proteins with his tags bind?

- They will bind to columns that have a nickel ion immobilized on the column. Since consecutive histidine residues are rare in natural proteins, it ability to bind to immobilized nickel makes a protein with the tag unique compared to other proteins in the mixture and allows for easy purification using the appropriate column

what do we add to the column once a sample has entered the column and why do we add it?

- A wash buffer is continuously added to the top of the column to promote the flow of protein solution within the column and to prevent the column from becoming dry

Ideally at this point only buffer and protein of interest remain in the column. How do we remove the bound protein from the column?

- There are multiple ways to do this but the most common ways to add high concentrations of another molecule usually very similar in structure to the ligand that is covalently bound to the resin, that also has the ability to bind to the protein of interest. The high concentration of the added free ligand outcompetes the column for binding of the protein and as a result the protein elutes from the column and it can be collected in a fraction relatively pure for the protein

The buffer that is used to remove the protein from the beads is known as what?

- elution buffer

it is highly likely that the high concentration of ligand in elution buffer is undesirable for further analysis of the protein. How do we separate the ligand from the protein?

- In most cases the ligand is quite small relative to the size of the protein so size can be used as the basis for separating the protein and the ligand from each other

how does size exclusion chromatography also known as gel filtration work?

- It separates molecules on the basis of their size

what is the difference between size exclusion chromatography and the two previously described methods in terms of beads?

- The beads for this form of purification do not have functional groups attached to them. Instead the beads are porous and filled with channels of varying sizes. The beads, as seem prior, are relatively inert and do not have any chemical interaction with proteins or other molecules in a buffer

How do the beads results in size exclusion in gel filtration?

- Molecules may or may not be able to enter into the pores of the beads and channels within the beads depending upon how big the molecules are. Small molecules can enter the bead move around in the bead before exiting the bead and returning to the general flow of the mobile phase. Larger molecules may not be able to enter the bead because they're too big which means they remain in the mobile phase moving through the column faster than the smaller beads

- in other words any molecule that cannot enter into the bead will elute first and smaller molecules will elute later according to their size with the smallest molecules alluding last

- This separation is due to the channels within the beads being of varying sizes

- it should be noted however that all of the molecules that are too big to fit within a bead will elute together even if they are different sizes and conversely all molecules that can fit within all of the pores will elute together even if they are different sizes

what can size exclusion chromatography also be used as?

- An analytical technique for relatively pure protein mixtures

Regarding lysozyme assays, why will the slope of the line be negative if there is lysosome in that fraction?

- The cell walls of the substrate are opaque to light at 450 nano meters, so at the start of the assay the spectrophotometer will show an absorbance reading well above 0, but if there is any lysozyme present, the bonds between the monomers of the cell walls will be hydrolyzed by the enzyme and the absorbance will go down because more light is now reaching the detector. Since the presence of lysozyme causes the absorbance to fall with time, the slope of the line will be negative

Why does SDS-PAGE require/depend additional technical details?

- Due to the high structural variation of proteins

Migration of a charge particle in solution under the influence of an electric field can be represented by what equation?

V = Eq/f

- V = velocity

- E Is strength of the electric field to which the charge particle is exposed

- q is the net charge on the particle

- f is the frictional coefficient of the particle as it moves through the solution

you saw that for the uniformity of DNA structure meant that the DNA molecules of different sizes had similar QF ratios and therefore would behave similarly in solutions. The situation is very different for proteins. How?

- Proteins are polymers of 20 different amino acids and some amino acids have ionizable side chains which may be deprotonated depending on the pH of the solution. Thus the charge on the protein depends upon the actual amino acids that are present and the pH of the solution

you saw that for the uniformity of DNA structure meant that the DNA molecules of different sizes had similar QF ratios and therefore would behave similarly in solutions. The situation is very different for proteins. What does this change?

- The effect of this is that proteins almost always have a charge and the net charge of the protein, Q, is the sum of all the various charges of these ionizable side chains on amino acids at any particular pH. Since different proteins have different amino acid compositions, different proteins will have a different charge at a particular pH which means of course at the numerator and the previous equation will be different for different proteins. This variety of charge or heterogeneity may appear to be a useful basis for separation of proteins

what is native structure of a protein?

- The inherent charge shape and size of a protein

what does the SDS PAGE version of electrophoresis do?

- It is a method that separates proteins from each other on the basis of length of their polypeptide chains and is probably the most commonly used technique in a biochemistry lab for the evaluation of protein purity and for obtaining some basic structural information about a protein

what are the two reasons why SDSPAGE can separate proteins on the basis of size?

1. During the preparation of the proteins for loading on the gel they all are structurally altered to give them the same QF ratio

2. They are electrophoresed through a sieving structural support which we're calling gels

what do you know about sample preparation for SDS PAGE?

- Protein sample is heated to about 95 degrees Celsius for five minutes in loading buffer that contains beta mercaptoethanol and sodium dodecyl sulfate (SDS)

In sample preparation, protein is denatured. How does this happen?

- Through the combination of high temperature and the reduction of disulfide bonds between cysteine residues by the beta mercaptoethanol

in sample preparation, why do we use SDS?

- It's a surfactant and after the protein is denatured it associates with the peptide bonds of the protein at a constant ratio of approximately 1.4 grams of SDS for every one gram of protein. Since SDS carries a negative charge the denatured protein is now covered with negative charges that greatly exceed the original charge on the protein

what is the combined effect of all of these substances and methods used in sample preparation?

- All proteins now roughly look like long rods coated with a negative charge. Proteins of different sizes will now be rods of different links but all will have similar QF values

what do you know about the loading buffer in relation to preparation of proteins?

- It will contain glycerol or sucrose to give the loading buffer greater density so that when it is applied to the wells of the gel the sample sinks directly to the bottom of the well

the gels for SDS PAGE are based upon what?

- Acrylamide

Polyacrylamide gels are formed by what?

- The crosslinking, polymerization, of acrylamide with bisacrylamide in a 37 to one ratio in the presence of the inhibitor catalyst system of ammonia persulfate and TEMED

What does TEMED do in the gel?

- It is the catalyst and it causes the formation of free radicals from the ammonium persulfate which induces the polymerization of the acrylamide molecules into long chains with the occasional bis acrylamide forming across link joining two linear polysaccharide polymers

The mesh network formed by the reaction of TEMED generates what?

- A gel that is characterized as having numerous pores of various sizes and the average pore size decreases with higher concentrations of acrylamide

what do you know about the percentage of acrylamide present in the gel?

- Once the percentage exceeds 4, the average pore size is sufficiently reduced such that solvent molecules can still easily pass through the gel unimpeded but larger molecules like the denatured proteins can only migrate through the larger pores

what is SDSPAGE frequently performed using?

- A discontinuous gel

what is a discontinuous gel?

- It is a gel where the bottom portion is called the separating or resolving gel and will typically have been acrylamide concentration of 6 to 15% depending on the application. A stacking gel made to 4% acrylamide sits atop the separating gel and functions primarily to focus the protein sample into a tight band before the separation by size that occurs in the separating gel

What do you know about loading the gel of an SDS PAGE?

- The micro pipettor tips that are used have long flexible tips to enable you to insert the tip between the glass plates and load samples directly into the well. Typically one loads between 10 to 20 microliters of mixture

after the electrophoresis is completed, you need to dye the gel to visualize the proteins. What do you know about this?

- Most common method is to use coomassie based stain - recall that this dye used in Bradford protein quantification assays

- typically the gel has to stain for 10 to 20 minutes in a mixture of the dye, methanol and glacial acetic acid which allows the die to reach their protein embedded in the gel

The method previously described to dye the proteins also dyes the gel. What do we have to do to effectively see the proteins?

- The gel needs to be destained to remove stain that is not bound to the protein in the gel. The destaining solution is usually a glacial acidic acid methanol mixture. It typically requires 24 to 48 hours depending on the destaining conditions

Regarding SDS PAGE, how can the purity of a protein sample be assessed?

- By the number of bands that are present. A single band suggests pure protein whereas numerous bands usually indicate that different types of protein are present in the sample that was electrophoresed

How can you estimate the size of your protein?

- Comparing the migration of your protein of interest to the standard protein of known size also known as the ladder

what does lactate dehydrogenase (LDH) do?

- Catalyzes the reversible reduction of pyruvate into lactate and is abundant in many tissues including the heart liver and skeletal muscle

How is LDH used in skeletal muscle?

- In tissue susceptible to the development of anaerobic conditions, such as heavily active skeletal muscle, the enzyme is crucial for the regeneration of the NAD+ that is required to sustain glycolysis

how is LDH used in the liver?

- The enzyme functions to catalyze the reverse reaction for the conversion of lactate back into pyruvate with the other enzymes for gluconeogenesis being responsible for the eventual regeneration of glucose

Regarding affinity chromatography, what was the ligand solid support that we used for the experiment?

- Cibacron Blue F3GA marketed as blue sepharose 6 fast flow which has a functional group that resembles NADH, and the bound LDH will be eluted with one micro molarity NADH

What is a spectrophotometric assay?

- It is a type of assay that can be used to measure the amount of protein that is present at any step of the purification process. since the protein we were attempting to purify was an enzyme, this specific assay was useful

the assay for our purification was based upon what fact?

- The fact that the coenzyme for LDH, (NAD/NADH), absorbs light differently depending upon its chemical state. The reduced form, NADH, absorbs light at 340, whereas the oxidized state, NAD, does not. Therefore the process of the enzymatic reaction can be followed by measuring the decrease in absorbance at 340. If the concentrations of NADH and pyruvate is the same for different assays, the rate at which the absorbance at 340 falls will be portion of the amount of LDH enzyme that is present in the assay cuvette

why did we have to wear gloves when using acrylamide?

- It is a potential neurotoxin

what did we add to the Falcon tube to promote the polymerization of the acrylamide?

- TEMED and APS

what can a vertical gel electrophoresis unit be used for?

- It can be used to run 2 gels simultaneously

what is Total activity?

- How much activity was loaded on top of column or how much activity was collected in fraction

what is % recovery of activity?

- How much enzyme activity did you collect in fraction compared to what you loaded on the column

what is Total protein?

- How much protein loaded on top of column or how much protein was collected in fraction

what is % recovery protein?

- How much protein did you collect in fraction compared to what you loaded on the column

what is specific lysozyme activity?

- Ratio of how much enzyme is there to how much protein is there

what are chelators?

- molecules specifically designed to remove ions from solution that are known to be required co-factors that can degrade biological molecules