Premed · Premed · Biochemistry
Lecture 6: Protein Purification and Analysis Techniques
Biochemistry
Learning Objectives
By the end of this lecture, students will be able to:
- Explain the principles behind common protein purification techniques
- Describe how chromatographic methods separate proteins based on different properties
- Interpret results from SDS-PAGE and isoelectric focusing
- Explain how mass spectrometry and Edman degradation are used to identify and sequence proteins
- Describe how protein concentration and purity are assessed
- Design a basic protein purification strategy
Lecture Content
I. Overview of Protein Purification
The goal of protein purification is to isolate a single protein from a complex mixture, given that a cell lysate contains thousands of different proteins. The general strategy proceeds through several stages: cell lysis and extraction from the source material, crude fractionation (such as ammonium sulfate precipitation or differential centrifugation), multiple chromatographic steps that exploit different protein properties, and assessment of purity at each step.
Several key considerations guide the process. Work should be performed at 4 degrees C to minimize proteolysis and denaturation. Protease inhibitors such as PMSF, leupeptin, and EDTA should be included. Appropriate pH and ionic strength must be maintained throughout, and the protein of interest should be tracked with an activity assay or antibody. A purification table is used to monitor total protein, total activity, specific activity, yield, and fold purification at each step.
II. Cell Lysis and Initial Fractionation
Cells can be lysed by mechanical means (homogenization, sonication, French press), chemical means (detergents such as Triton X-100 or SDS, osmotic shock), or enzymatic means (lysozyme for bacteria, cellulase for plants).
Differential centrifugation separates cellular components by size and density. Low speed centrifugation at 1,000 x g pellets nuclei and cell debris. Medium speed at 10,000 x g pellets mitochondria, lysosomes, and peroxisomes. High speed at 100,000 x g pellets microsomes (ER fragments) and ribosomes, leaving the cytosol in the supernatant.
Ammonium sulfate precipitation (salting out) works by reducing the water available to solvate proteins. Different proteins precipitate at different salt concentrations depending on their surface hydrophobicity. Fractions collected at increasing salt concentrations are tested for the target protein. While crude, this is an effective first step that also concentrates the protein.
III. Chromatographic Methods
Ion Exchange Chromatography
Ion exchange chromatography separates proteins by net charge. Cation exchange resins such as CM-cellulose carry negative charges and bind positively charged proteins, while anion exchange resins such as DEAE-cellulose carry positive charges and bind negatively charged proteins. Bound proteins are eluted by increasing salt concentration through a NaCl gradient or by changing pH. Binding behavior can be predicted from the protein's pI and the buffer pH.
Size Exclusion (Gel Filtration) Chromatography
Size exclusion chromatography separates proteins by their hydrodynamic radius. The column is packed with porous beads of defined pore size. Small proteins enter the pores and follow a longer path, eluting later, while large proteins are excluded from the pores and elute first. This method can estimate native molecular weight by comparison to standards and is also useful for buffer exchange and desalting.
Affinity Chromatography
Affinity chromatography is the most selective method, exploiting specific binding interactions. The column contains an immobilized ligand specific for the target protein. Examples include immobilized antibodies to capture target antigens, Ni-NTA resin to bind His-tagged recombinant proteins, glutathione-Sepharose for GST-tagged proteins, lectin columns for glycoproteins, and substrate analogs for enzymes. The target protein binds while contaminating proteins wash through, and elution is achieved with free ligand, pH changes, or imidazole (for His-tags). This method often provides high purity in a single step.
Hydrophobic Interaction Chromatography (HIC)
HIC separates proteins by surface hydrophobicity. Proteins bind at high salt concentration and are eluted by decreasing salt. This technique complements ion exchange chromatography nicely.
<image>A four-panel figure illustrating the four major chromatographic techniques used in protein purification. Panel A: Ion exchange chromatography showing a positively charged column resin binding negatively charged proteins, with a salt gradient elution profile. Panel B: Size exclusion chromatography showing large proteins excluded from beads eluting first and small proteins entering pores eluting later, with an elution profile showing peaks in order of decreasing size. Panel C: Affinity chromatography showing a specific ligand immobilized on beads binding only the target protein (shown in color) while others wash through, followed by elution with free ligand. Panel D: A flowchart of a typical purification scheme combining these methods in sequence.</image>
IV. Electrophoretic Methods
SDS-PAGE (Sodium Dodecyl Sulfate - Polyacrylamide Gel Electrophoresis)
SDS-PAGE is the standard method for assessing protein purity and determining subunit molecular weight. SDS is an anionic detergent that denatures proteins and coats them with a uniform negative charge at approximately 1.4 g SDS per gram of protein, giving all proteins approximately the same charge-to-mass ratio. As a result, proteins separate solely by size, with smaller proteins migrating faster. Beta-mercaptoethanol or DTT reduces disulfide bonds. Molecular weight is determined by comparison to a protein ladder of standards, and proteins are visualized by Coomassie blue staining (detecting approximately 0.1 microgram) or silver staining (detecting approximately 1 ng). A single band on SDS-PAGE indicates purity at the resolution of the gel.
Native PAGE
In native PAGE, no SDS or reducing agents are used, so proteins maintain their native conformation and charge. Separation occurs based on size, shape, and charge, making it useful for assessing quaternary structure and native protein complexes.
Isoelectric Focusing (IEF)
IEF separates proteins based on their isoelectric point. The gel contains an immobilized pH gradient, and proteins migrate until they reach the pH equal to their pI, where their net charge is zero and migration stops. IEF provides very high resolution for separating proteins with similar size.
2D Gel Electrophoresis
Two-dimensional gel electrophoresis combines IEF in the first dimension with SDS-PAGE in the second dimension, resolving thousands of proteins in a single experiment. Each spot represents a protein with a unique pI and molecular weight. This technique is widely used in proteomics to compare protein expression between samples.
V. Protein Detection and Quantification
The Bradford assay uses Coomassie blue dye binding and measures absorbance at 595 nm; it is quick and convenient. The BCA assay uses bicinchoninic acid and is based on Cu2+ reduction; it is compatible with detergents. UV absorbance at 280 nm relies on tryptophan and tyrosine residues absorbing UV light, with concentration determined by the Beer-Lambert law (A = epsilon x c x l), where the extinction coefficient is protein-specific.
The Western blot (immunoblot) is a highly specific technique for detecting a particular protein. Proteins are first separated by SDS-PAGE, then transferred to a membrane (nitrocellulose or PVDF), probed with a primary antibody specific for the target protein, and detected with a secondary antibody conjugated to an enzyme (HRP or AP) or fluorophore. The technique can detect very low amounts of a specific protein.
<image>A step-by-step diagram of a Western blot procedure. Step 1: SDS-PAGE separation of protein mixture. Step 2: Transfer to nitrocellulose membrane via electrophoretic transfer (showing the sandwich setup with gel and membrane between filter paper). Step 3: Blocking with milk or BSA. Step 4: Incubation with primary antibody (shown binding to target band). Step 5: Incubation with enzyme-conjugated secondary antibody. Step 6: Detection by chemiluminescence or colorimetric substrate, showing a specific band on film. A parallel Coomassie-stained gel is shown for comparison.</image>
VI. Mass Spectrometry
Mass spectrometry determines the precise molecular weight of proteins and peptides by ionizing molecules, separating them by mass-to-charge ratio (m/z), and detecting them. In MALDI-TOF (Matrix-Assisted Laser Desorption Ionization - Time of Flight), the protein is mixed with a matrix and hit with a laser pulse. Ionized proteins fly through a vacuum tube, with smaller ions arriving at the detector first, yielding an accurate molecular weight. In ESI-MS (Electrospray Ionization), the protein solution is sprayed through a charged needle, generating multiply charged ions; this approach is often coupled with liquid chromatography (LC-MS/MS).
Peptide mass fingerprinting involves digesting the protein with trypsin (which cleaves after lysine and arginine), measuring the masses of the resulting peptides by MALDI-TOF, and comparing the pattern to a database of predicted tryptic peptides. Tandem MS (MS/MS) further fragments individual peptides for analysis and can determine amino acid sequences de novo. It is the gold standard for protein identification in proteomics.
VII. Protein Sequencing
Edman degradation is a chemical method for determining amino acid sequences. Phenylisothiocyanate (PITC) reacts with the N-terminal amino acid, which is then cleaved and identified by HPLC. The process repeats one residue at a time, with a practical limit of approximately 50 to 60 residues before signal degradation. Although largely replaced by mass spectrometry, Edman degradation is still used for N-terminal identification. The modern approach relies on DNA sequencing of the gene and computational prediction of the protein sequence, which is faster and more reliable for full-length sequences.
<image>A diagram of the Edman degradation process. Step 1: PITC reacts with the free alpha-amino group of the N-terminal residue under mildly alkaline conditions, forming a PTC-protein. Step 2: Treatment with anhydrous acid cleaves the PTC-amino acid as a thiazolinone derivative, leaving the rest of the polypeptide intact with a new N-terminal residue. Step 3: The thiazolinone is converted to a more stable PTH-amino acid and identified by HPLC. Step 4: The cycle repeats with the next amino acid. A chromatogram showing PTH-amino acid identification is included.</image>
