Key Area 1

(c) Separation Techniques

Centrifugation

Centrifugation is a separation technique used to separate substances of differing density. More dense components settle as a pellet; less dense components remain in the supernatant. This is really useful technique, often used when trying to extract enzymes from tissue, e.g. Dopa oxidase can be found in many tissues, including banana - centrifugation of a mashed banana allows separation of crude, dense cell components from less dense cellular materials, including enzymes.

Inside a centrifuge is a metal rotor that can hold samples. The centrifuge spins at a set speed, causing the components of the mixture to separate based on their relative densities - most dense materials settle at the bottom (forming a "pellet"), while least dense can remain in the upper liquid portion (forming the "supernatant").

This image shows the tubes of samples within the centrifuge. The rotor spins the samples at high speed. It is important to carry out a Risk Assessment if you are intending on using a centrifuge in your school lab.

Following centrifugation, your sample should have separated to form a pellet and supernatant - cells and cruder cell structures form the pellet, while less dense cellular substances, including enzymes, will form the supernatant. The supernatant can be extracted very carefully, using a pipette, and used in assays.

PPQ6

2023, Section 1, Qu 3.

Chromatography

Chromatography refers to a set of techniques which separates the components of a mixture. It can be used to separate mixtures of amino acids and proteins. The speed that each solute travels along the chromatogram depends on its differing solubility in the solvent used.

There are three types of chromatography:

The migration distance of a solute in the mixture is known as the Rf value.

The Rf value is often used as a defining point of information which allows identification of a particular compound in a mixture.

Task 9

Billy cascaded into the class on Monday - "Alright Miss, how's it going? I've got those leaves you were after".

Dr McRobbie smiled with relief - she knew she could count on Billy to remember. Today was Chromatography day and the class were comparing the photosynthetic pigments found in different plants. 

Billy used a cork borer to extract 1 cm2 of the leaf and ground it up in a mortar containing 750 ml propanone. He poured the liquid into small bottle.



He then prepared his chromatogram, using a pencil to mark 1.5cm from the bottom of the plate. Using a piece of capillary tubing, he added drops of the leaf mixture to the chromatogram, repeating the addition to form one small concentrated blot of sample.


The mobile phase solvent was then added to a chromatography chamber and the chromatogram was gently placed into this, centrally. It was removed from the chamber when the solvent front had nearly reached the top of the stationary phase.


The solvent front was marked using a pencil.

Billy recorded the distance travelled by each band and the solvent front, as shown below.

Task:



Answers are available here.

Chromatography Reference - Rf values and identification of photosynthetic pigments.

Chromatography activity

Try this practical activity on TLC from the SSERC website. There is a powerpoint to go along with it here. 

Affinity Chromatography: principles and its uses 

Affinity chromatography can be used to separate a mixture of proteins. A solid matrix or gel column is created with specific molecules bound to the matrix or gel. 

Soluble, target proteins in a mixture, with a high affinity for these molecules, become attached to them as the mixture passes down the column. 

Other non-target molecules with a weaker affinity are washed out.

Affinity chromatography is a useful technique to separate a protein of interest from a crude mixture. The protein of interest often contains a tag that has high affinity for a molecule bound to the column. This allows its separation from the rest of the mixture.

Task 10

Dr McRobbie had just finished delivering the lesson on Affinity Chromatography and presented her class with the following image. She asked the class to explain what was happening. Wee Jonny was sat up the back swinging on his chair - which Dr McRobbie hates. "Wee Jonny", she said, "how about you have a go explaining this to us all".  

Oh no, wee Jonny wasn't listening at all. His head was in the clouds thinking about Catriona.


Can you help wee Jonny out and explain the image opposite? What is happening and how can this be used to produce a pure sample of the "red protein"?


Answers are available here.

Gel Electrophoresis

Gel electrophoresis is used to separate mixtures of proteins and mixtures of nucleic acids. Charged macromolecules move through an electric field applied to the gel matrix. Smaller proteins migrate faster due to less resistance from the gel matrix. Other influences on the rate of migration through the gel matrix include the structure and charge of the proteins. There are two main types you need to be aware of:

Native gel electrophoresis

Native gel electrophoresis involves separating fully folded proteins. Separation occurs on the basis of shape, size and net charge. This type of electrophoresis can be useful because information can be gained about the quaternary structure of a protein - this refers to the number of subunits a protein has, e.g. haemoglobin has 4 subunits, which can be seen in a native gel (this would not be possible in a denaturing gel environment). Furthermore, because the native structure of the protein is retained, enzyme activity usually is also. This means that native gel electrophoresis can be used to purify proteins - the band can be cut out of the gel and the protein stored and used later!

Denaturing SDS-PAGE gel electrophoresis

Denaturing gels gives all the molecules an equally negative charge and denatures them, separating proteins by size alone.  

It does this by using a chemical called SDS along with a reducing agent - SDS is a detergent with a strong protein-denaturing effect and binds to the protein backbone. The reducing agent serves to cleave disulfide bonds, which are critical in maintaining the 3D shape of a protein. 

As a result of these treatments, proteins unfold into linear chains with negative charges. The protein mixtures are then subjected to polyacrylamide gel electrophoresis (PAGE) and proteins are separated based on their size, with high molecular weight proteins migrated more slowly through the gel matrix compared to smaller proteins.


This image shows SDS-PAGE being carried out and protein migration can be seen. In each gel, 2 molecular mass markers have been included - each band in these "ladders" indicate a particular molecular mass so the researcher can work out the molecular mass of their protein of interest. This is a powerful identification tool.

During the process of SDS-PAGE, the sample is prepared and denatured before loading onto a polyacrylamide gel using a micropipette. A current is passed through the gel and the protein is separated based on its molecule mass.

This image shows a SDS-PAGE gel Dr McRobbie produced many moons ago during her PhD. The gel shows 3 purified proteins, all involved in DNA repair pathways. The Molecular Mass Ladder (left lane) provides a marker to indicate approximate sizes of these purified proteins, e.g. RadA is ~40kDa, which is close to theoretically-derived masses from bioinformatic tools.

Gel electrophoresis activity

Try this gel electrophoresis practical activity from SSERC.

There is a powerpoint to support this as well.

Task 11

In a former life, Dr McRobbie had a close relationship with a protein called XPD. She used an Archaeal model organism (called Sulfolobus) to study XPD which, when mutated in humans, is associated with a particular form of skin cancer. But Dr McRobbie had to clone XPD in the lab, based on the DNA sequence and introduce mutations. 

Here is XPD:


Dr McRobbie carried out some simple bioinformatics, like you have done in Higher Biology/Human Biology, and found out that the theoretical molecule mass of XPD was about 64kDa.

The process of producing XPD involved expressing the protein in bacteria - think back to your theory of genetic engineering. XPD gene was introduced to a bacterial plasmid and the GM-plasmid was then re-introduced back into the bacterium. The GM-bacterium was then grown in culture to produce many copies of XPD. 

However, the XPD must be purified from the bacterial cells and all its own proteins. Dr McRobbie needed to separate XPD from this mixture of mess and then make sure she had cloned what she really wanted.

Dr McRobbie performed the following procedures. Note: For the SDS-PAGE gel, look at the band on the far right under GF. 

M stands for Molecular Mass marker/ladder.

CL - crude extract containing all the proteins in the bacterial culture.

H - the extract after a first step purification step.

Ni - the extract after an affinity chromatography step.

GF - the extract after a gel filtration chromatography step.

Question: Describe what was happening during each of the processes shown in Dr McRobbie's procedure. Do you think she had successfully cloned and expressed archaeal XPD?


Answers are available here.

Isoelectric points

Proteins can be separated from a mixture using their isoelectric points (IEPs). The IEP is the pH at which a soluble protein has no net charge and will precipitate out of solution.

If the solution is buffered to a specific pH, only the protein(s) that have an IEP of that pH will precipitate.

Proteins can also be separated using their IEPs in electrophoresis. Soluble proteins can be separated using an electric field and a pH gradient. A protein stops moving through the gel at its IEP in the pH gradient because it has no net charge.

The IEP of casein, a milk protein, can be determined by adding HCl to milk, dropwise (image on right), while measuring the pH using a digital pH meter. Record the pH at which the milk starts to look lumpy - this is casein's IEP as it precipitates and comes out of solution. The image on the left shows casein separated and dried having precipitated from the milk solution.

The image shows that as pH changes, the net charge of the protein changes. The pH at which the protein has no net charge is called its isoelectric point (IEP).

Isoelectric point practical activity

Try this activity from SSERC - if you don't have a pH probe/meter, this is ok - you can simply add the same acid, drop-wise, to the sample of milk until you see coagulation. 

There is an accompanying bioinformatics resource for casein protein as well.

PPQ7

Specimen paper, Section 1, Question 3

PPQ8

Specimen paper, Section 1, Question 4

PPQ9

2022, Section 1, Question 1

PPQ10

Exemplar, Section 2, Question 2b

PPQ11

2017, Section 1, Question 2

Go to SCHOLAR "1.3 Separation techniques" for further insight.

You are now ready to move on to Topic 1, Key Area 1d: Detecting proteins using antibodies