Block A Lecture 2 - Methods for Protein Purification

Question 1

What is gel filtration?

Answer 1

A column chromatography technique used to seperate proteins based on shape and size.

The column contains porous beads which make up the stationary phase, which results in smaller proteins entering the pores of the bead, taking a longer path through the column, whereas larger molecules cannot enter the pores, and move faster through the column

(Slide 5)

Question 2

What is the general buffer recommendation which is used in gel chromatography, and why is it used?

Answer 2

A buffer with intermediary ionic strength, which suppresses ionic effects and does not support hydrophobic interactions

(Slide 6)

Question 3

What are 3 factors which can affect the resolution in gel filtration?

Answer 3

Diffusion (the particle size of the column material and flow rate)

The volume of the sample

Viscosity of the sample

Relative amounts of proteins to be separated

(Slide 7)

Question 4

What does a smaller particle size and lower flow rate effect in the context of gel filtration chromatography, and what can this result in?

Answer 4

Smaller particles reduce diffusion, resulting in a better resolution.

A lower flow rate can improve separation but may lead to band spreading due to increased diffusion

(Slide 7)

Question 5

What are 3 uses of gel filtration?

Answer 5

Answers Include

Change of buffer (desalting)

Removal of reaction produce, co-factors or inhibitors

Removal of phenol red from culture medium prior to anion exchange chromatography

Fractionation

Determination of molecular mass

(Slide 9)

Question 6

How is a calibration curve generated in gel filtration chromatography?

Answer 6

By injecting standard proteins of known molecular weights into the column which will partition between the pores of the Sephadex G-200 matrix

(Slide 11)

Question 7

What is covalent chromatography, how are proteins eluted and what proteins is it usually used to purify?

Answer 7

The stationary phase has reactive disulphides (such as 2-pyridyl disulphide) covalently bound in order to enable selective covalent bonding to proteins containing free thiol groups.

These bonds are stable and after the rest of the proteins have passed through the analytes can be eluted by changing conditions such as pH, temperature, or adding a ligand, which breaks the covalent bonds.

It is usually used to purify proteins with cysteine residues (as they contain thiol groups) or for selectively removing specific proteins from a mixture

(Slide 13)

Question 8

What has to be removed from the sample before covalent chromatography, and how is this done?

Answer 8

Low molecular weight thiols (such as glutathione), which can be removed by dialysis or gel filtration

(Slide 14)

Question 9

How is binding monitored in covalent chromatography?

Answer 9

By using UV absorption to measure absorbance (at 343 nm) in the eluate (mobile phase leaving the column)

(Slide 14)

Question 10

What is the principle behind ion exchange chromatography and how does it work?

Answer 10

It separates proteins based on net electrical charge at a given pH.

Proteins with a positive charge will bind to a negatively charged stationary phase (a cation exchanger) whereas negatively charged proteins will bind to a positively charged stationary phase (an anion exchanger)

(Slides 18 - 26)

Question 11

What are ion exchangers in the context of ion exchange chromatography?

Answer 11

A stationary phase which contains functional groups capable of binding and releasing ions.

These sites are usually charged (positively or negatively) and can attract and hold ions of the opposite charge from the solution, separating them.

(Slide 18)

Question 12

What is the difference between an anion and a cation exchanger?

Answer 12

An anion exchanger is a basic ion exchanger with positively charged functional groups whereas a cation exchanger is an acid ion exchanger which has negative functional groups

(Slide 18)

Question 13

What is the difference between a strong and a weak ion exchanger? Give an example of each.

Answer 13

A strong ion exchanger (such as those which contain sulphonate or quaternary amino functional groups) remain ionised over a broad pH range, ensuring consistent binding capacity and therefore separation.

Weak ion exchangers such as those which contain carboxymethyl (-OCH2COOH) functional groups (pKa of 4-5) have a binding capacity which is pH dependent and decreases when pH approaches their pKa

(Slide 19)

Question 14

How can all proteins display cationic or anionic behaviour?

Answer 14

In acidic pH amino groups of lysine, arginine and histidine are protonated, so the protein can display cationic behaviour

In basic pH the negative charges of carboxyl groups of aspartate and glutamate take over, so the protein displays anionic behaviour

(Slide 20)

Question 15

How does the isoelectric point tell us if a protein is basic or acidic?

Answer 15

If IP is above 8 - protein is basic.

If IP is below 6 - protein is acidic.

(Slide 20)

Question 16

What are 3 requirements of a buffer used for ion exchange chromatography?

Answer 16

Answers Include:

Its ionic strength must be high enough to guarantee protein stability

It must have a high buffer capacity -the pKa of buffer is not more than 0.5 pH units from the pH of the sample.

The pH of the buffer should be at least 1 - 2 units above or below (depending on whether you are trying to bind a positively or negatively charged protein) the isoelectric point of the target protein.

Buffer ions should have the same sign as the charged groups of the ion exchanger, otherwise they would act as “counter-ions”

(Slide 22)

Question 17

If freeze drying is planned in ion exchange chromatography, what property should the buffer used have?

Answer 17

It should be volatile

(Slide 22)

Question 18

How does a salt concentration gradient work in ion exchange chromatography?

Answer 18

The elution buffer initially has a low concentration of salt. This allows for weaker or less-affinity interactions between the target analytes and the ion exchange resin, meaning the analytes stay bound to the resin.

As the salt concentration is gradually increased, ions from the buffer (such as Na⁺ or Cl⁻) begin to compete with the analytes for the binding sites on the resin.

This results in analytes being displaced from the resin, with the higher the concentration of salt, the more analytes which are displaced, creating a concentration gradient

Note: This is a type of gradient separation technique, with this term being mentioned in lecture 1.

(Slides 23 - 25)

Question 19

How does a pH gradient work in ion exchange chromatography?

Answer 19

pH is altered towards the isoelectric point of the substance to be eluted, as this is where many proteins have their lowest solubility.

Substances with weak interactions come off first.

This method results in aggregation via reduced repulsion - allowing proteins to be sorted

(Slide 26)

Question 20

What is hydrophobic interaction chromatography?

Answer 20

A form of chromatography which separates proteins based on their hydrophobicity. It relies on the reversible interaction between the hydrophobic regions on the surface of a protein and hydrophobic groups immobilized on the stationary phase.

Protein binding is supported by an increased concentration of anti-chaotropic (neutral) salts, as they bind strongly with the water molecules, leaving less water available to stop the hydrophobic interaction.

(Slides 30 and 32)

Question 21

What are 2 common methods used for eluting proteins in hydrophobic interaction chromatography?

Answer 21

1. Gradual reduction in the anti-chaotropic salt concentration in the buffer, resulting in the weakening of the hydrophobic interactions.
2. Gradual addition of chaotropic agents, such as water-miscible alcohols, detergents, or chaotropic salts, which disrupt hydrophobic interactions.

(Slide 30)

Question 22

What protein purification step is hydrophobic interaction chromatography well suited to follow?

Answer 22

Protein precipitation (via ammonium sulphate)

(Slide 31)

Question 23

Why is hydrophobic interaction chromatography (HIC) suited to be the first chromatographic step?

Answer 23

As most biological starting material already has a high ionic strength (high conductivity)

(Slide 31)

Question 24

Why is hydrophobic interaction chromatography suited to be used after ion exchange chromatography?

Answer 24

As the sample is already in a buffer with high ionic strength

(Slide 31)

Question 25

What does increasing pH and lowering temperature do to the strength of hydrophobic interactions?

Answer 25

Increasing pH increases the strength of hydrophobic interactions whereas lowering the temperature reduces strength (Slide 31)

Question 26

What is the difference between precipitating agents and additives that reduce hydrophobic interactions?

Answer 26

Precipitating agents promote protein aggregation and precipitation by encouraging proteins to become less soluble in the solution whereas additives that reduce hydrophobic interactions help proteins stay in solution by disrupting hydrophobic interactions that could cause aggregation, essentially increasing protein solubility. i.e they are essentially opposites; precipitating agents promote proteins precipitating out a solution whereas reducing hydrophobic interactions promotes proteins staying in solution (Slide 31)

Question 27

When would you use additives which reduce hydrophobic interactions over precipitating agents?

Answer 27

if the target protein is prone to aggregation (forming clumps) and you want to keep it soluble in the solution (Slide 31)

Question 28

What are 2 examples of salts which are commonly used in hydrophobic interaction chromatography?

Answer 28

Answers include: Na2SO4 NaCl (NH4)2SO4 (Slide 33)

Question 29

In what situation does hydrophobic interaction chromatography yield a fairly concentrated protein?

Answer 29

In a low ionic strength solution (Slide 34)

Question 30

What is affinity chromatography and what mixtures is it usually used to separate?

Answer 30

A specific type of chromatography where molecules are separated based on their specific interactions with a biomimetic ligand or a biologically functional partner. The ligand is immobilized onto the stationary phase, and the target molecule is selectively captured through this interaction. It allows highly selective purification from complex mixtures (Slide 36)

Question 31

How is the stationary phase produced in affinity chromatography and how does this work?

Answer 31

Activated gel materials are usually used which allows covalent coupling of the ligand (Slide 36)

Question 32

What are 3 features a ligand should have in affinity chromatography?

Answer 32

Answers include: It has a **reversible** complex formation with the protein to be isolated High specificity for the target protein Suitable dissociation constant to allow reversible complex formation without harming the protein (KD = 1 - 10 µM) Chemical features which allow immobilisation to a matrix (stable in solvent used for coupling, at least 1 functional group for immobilisation, functional group must also not take part in the interaction involved in purification) (Slide 37)

Question 33

What are the classifications of ligand which can be used in affinity chromatography?

Answer 33

They are somewhere between monospecific and group specific and between low molecular to macromolecular size (Slide 38)

Question 34

What are 3 ways in which elution can occur in affinity chromatography?

Answer 34

By changing pH Changing ionic strength Changing structure or polarity (can be done by adding chaotropic salts, urea or guanidinium hydrochloride, polarity can be decreased by adding ethylene glycol or dioxane, or addition to detergents close to critical micelle concentration) Specific elution by addition of a free ligand (Slide 39)

Question 35

What is the principle behind immobilised metal ion affinity chromatography (IMAC) and what proteins is it usually used to separate?

Answer 35

It uses a stationary phase that is covalently bound to metal-chelating groups, which bind metal ions. These metal ions then interact with proteins that have a high affinity for metal ions, such as those with histidine residues, allowing the proteins to bind to the stationary phase. This method enables the separation of proteins based on their affinity for metal ions. IMAC is particularly useful for isolating proteins with engineered polyhistidine tags (His-tags). (Slide 42)

Question 36

What is a metal-chelating group?

Answer 36

Functional groups which bind to metal ions at 2 (or more) co-ordination sites, donating a lone pair of electrons at each bond in order to form a stable co-ordination complex with the metal ion. (Slide 42)

Question 37

What results in a higher yield in immobilised metal ion affinity chromatography (IMAC)?

Answer 37

The metal-chelator group binding stronger to the ion, the stronger the binding, the higher the yield (Slide 42)

Question 38

What are free co-ordination sites?

Answer 38

They refer to the vacant positions or spaces on a metal ion where a ligand can bind (Slide 43)

Question 39

Why do free coordination sites need to be purified before immobilised metal ion affinity chromatography?

Answer 39

To ensure efficient and specific binding of target molecules to the column (Slide 43)

Question 40

What are 2 common methods used for eluting proteins in immobilised metal ion affinity chromatography (IMAC), and how do they do this?

Answer 40

Either by lowering pH, which protonates histidine residues, weakening their interaction with the metal ion or by adding imidazole or histidine residues, which compete with histidine residues on the protein for binding to the metal ion, displacing them from the stationary phase and resulting in the protein being eluted. (Slide 43)