Chapter 3/4 Methods

Protein Sequencing and Identification

• Primary Sequence Determination from Genomic Data:

  • The primary amino acid sequence of a protein is routinely predicted directly from the host organism's fully sequenced genome.

  • Triplet nucleotide codons in the gene sequence are translated into predicted amino acid sequences by identifying the translation start and stop codons.

  • Limitation: DNA sequence translation cannot identify post-translational modifications (PTMs), such as phosphorylation, acylation, or glycosylation. Identification of PTMs requires direct protein characterization.

• Edman Degradation (Classical Sequencing Method):

  • Involves successive rounds of N-terminal modification, cleavage, and chemical identification to sequentially remove and identify amino acid residues starting from the N-terminus.

  • Mechanism:

  1. High pH conditions: Phenylisothiocyanate reacts with the unprotonated $\text{N-terminal}$ amino group to form a phenylthiocarbamyl derivative.

  2. Low pH conditions: The N-terminal residue is cleaved as an anilinothiazolinone derivative, leaving a shortened peptide chain.

  3. Low pH conversion: The anilinothiazolinone derivative is converted into a stable phenylthiohydantoin (PTH) amino acid derivative, which is identified via chromatography.

  4. The remaining peptide fragment is isolated and recycled through subsequent Edman cycles.

  • Scope and Utility: Typically used to sequence the first $10\text{--}15$ amino acid residues from the N-terminus. This sequence length is usually sufficient to identify a protein with a known sequence, because the probability of identical $10 ext{--}15$ residue sequences occurring in multiple distinct proteins is extremely low.

  • Drawbacks: Highly time-intensive and strictly limited by sequence length per run.

• Mass Spectrometry (Modern Sequencing and Identification Method):

  • General Principles:

  • Charged gaseous ions are accelerated through an electric or magnetic field inside a mass spectrometer.

  • The flight behavior and trajectory of the ions correlate directly with their mass-to-charge ratio ($m/z$).

  • Proteins are enzymatically digested into small peptide fragments prior to analysis using proteases with known cleavage site specificities (e.g., trypsin).

  • Cleavage generates short peptides typically $3\text{--}15$ amino acids in length.

  • Complete enzymatic digestion of a typical protein yields approximately $20\text{--}25$ distinct peptide fragments.

  • Ionization Techniques:

  • MALDI-MS (Matrix-Assisted Laser Desorption/Ionization Mass Spectrometry): Volatilizes and ionizes intact peptides out of a light-absorbing matrix using laser pulses.

  • ESI-MS (Electrospray Ionization Mass Spectrometry): Desolvates and ionizes peptides directly from a liquid solution into a gaseous charged state.

  • Tandem Mass Spectrometry ($\text{MS/MS}$ or $\text{MS1}$ / $\text{MS2}$):

  • $\text{MS1}$ (Peptide Mass Fingerprinting):

    • Measures the overall mass-to-charge ratios ($m/z$) of all intact, unfragmented peptide ions in a digested sample.

    • Experimental peptide masses are matched against theoretical peptide masses derived from genomic databases.

    • High-confidence protein identification usually requires as few as $5$ distinct high-quality peptide masses.

    • Limitations of $\text{MS1}$: Cannot determine the linear sequence of amino acids within an individual peptide, nor can it pinpoint exact PTM attachment sites.

  • $\text{MS2}$ (Peptide Cleavage and Sequencing):

    • Measures full amino acid sequences or precise localization of PTM sites.

    • Ion Selection: The first mass analyzer ($\text{MS-1}$) isolates a specific target peptide ion identified in $\text{MS1}$ and discards all other ions.

    • Accumulation & Fragmentation: The isolated peptide ion is accumulated for a few seconds in a collision cell and bombarded with high-energy inert gas collisions to break backbone peptide bonds.

    • Fragmentation Spectrum: Cleavage occurs at various peptide bonds across multiple copies of the peptide chain, yielding $b$ ions (N-terminal fragments) and $y$ ions (C-terminal fragments).

    • Mass Analysis: The second mass analyzer ($\text{MS-2}$) determines the masses of the fragment ions, enabling precise reconstruction of the amino acid sequence and identification of PTM locations.

Objectives and Rationale of Protein Purification

• Why Purify Proteins?:

  • To characterize the biological function, enzymatic activity, and kinetic parameters of a specific protein.

  • To determine the three-dimensional atomic structure, which provides insight into function and molecular mechanism.

  • To study individual biochemical mechanisms and pathways without interference.

• The Fundamental Challenge:

  • Cells contain tens of thousands of distinct proteins, nucleic acids, lipids, and metabolites forming crowded, complex biological mixtures.

  • Purification isolates the target protein from all other cellular components to ensure experimental observations reflect the intrinsic properties of the target protein alone.

• The Overall Process of Protein Purification and Analysis:

  1. Select the specific target protein.

  2. Select an appropriate cell source (native tissue or recombinant expression system).

  3. Extract proteins from cells via cell lysis (crude extract formation).

  4. Separate proteins from crude cellular debris and organelles using centrifugation.

  5. Purify the target protein or protein complex using liquid chromatography techniques.

  6. Analyze purity, yield, molecular weight, function, and structure.

Protein Expression Sources: Native vs. Recombinant

• Native / Endogenous Source:

  • Target protein is extracted and purified directly from the natural organism, tissue, or cell type in which it is naturally expressed.

  • High Abundance Native Examples: Structural or oxygen-transport proteins such as collagen or hemoglobin are natively expressed at high concentrations.

  • Low Abundance Native Examples: Regulatory proteins, transcriptional repressors, or signal transduction factors are expressed natively at extremely low intracellular levels.

  • Drawbacks: Low yield for scarce proteins, requiring massive tissue volumes; potential ethical concerns regarding tissue harvesting; high financial costs.

• Recombinant Source (Heterologous Expression Systems):

  • Genetically engineering host cells ("cell factories") to express high levels of the target protein.

  • Procedure: The gene encoding the target protein is cloned from the native species and inserted into a expression vector, which is transformed into a host microorganism (most commonly Escherichia coli bacteria). If bacterial expression fails or lacks required modifications, eukaryotic expression hosts (e.g., Saccharomyces cerevisiae, insect cells, or mammalian cell lines) are used.

  • Advantages: Inexpensive, high yield potential, scalable production.

  • Challenges: Protein misfolding; formation of insoluble inclusion bodies; chaperone machinery discrepancies between host and native species; extensive optimization required for expression and folding protocols.

Cell Lysis and Extraction

• Cell Lysis (Extracting Intracellular Contents):

  • Destroys cell membranes to release the mixture of cytosolic proteins, generating a crude cellular extract.

  • Detergent Cleavage / Chemical Lysis:

  • Uses amphipathic detergent molecules to interact with and solubilize the lipid bilayer.

  • Detergents form pores in plasma membranes and dismantle lipid structures, releasing soluble intracellular contents.

  • Limitations: Detergents alone are often insufficient to disrupt rigid cell walls (found in plants, fungi, and bacteria), which require supplementary mechanical disruption (e.g., French press, sonication, bead beating) or enzymatic digestion (e.g., lysozyme).

Fractionation by Differential Centrifugation

• Centrifugation Principles:

  • Particles in a liquid suspension are separated based on size, mass, and particle density by applying centrifugal acceleration ($g$-force).

  • Heavier, larger, and denser components sediment faster to form a condensed pellet at the bottom of the tube; smaller, less dense components remain suspended in the liquid supernatant above the pellet.

• Sequential Steps in Differential Centrifugation:

  • Low-Speed Centrifugation: $5{,}000 \times g$ for $5\text{ min}$

  • Pellet: Unbroken intact cells, heavy cellular fragments, and dense cellular debris.

  • Supernatant: Transferred to a fresh tube for subsequent spin.

  • Medium-Speed Centrifugation: $20{,}000 \times g$ for $20\text{ min}$

  • Pellet: Subcellular organelles (mitochondria, lysosomes, peroxisomes).

  • Supernatant: Collected and transferred to a fresh tube.

  • High-Speed Centrifugation: $100{,}000 \times g$ for $1\text{ h}$

  • Pellet: Small membrane fragments, microsomes, small vesicles, and large macromolecular complexes.

  • Supernatant: Contains the soluble cytosolic protein fraction.

  • Very-High-Speed Centrifugation: $150{,}000 \times g$ for $3\text{ h}$

  • Pellet: Microsomes (endoplasmic reticulum membrane fragments) and small vesicles.

  • Supernatant: Purified soluble protein fraction.

• Distribution of Cellular Proteins:

  • Approximately $60\text{--}80\%$ of cellular proteins are soluble cytosolic proteins.

  • Approximately $20\text{--}40\%$ of cellular proteins are membrane-bound or organelle-associated.

• Purification Strategy:

  • Target Organelle or Membrane Protein: Isolate and collect the pellet fraction obtained during medium-speed or high-speed centrifugation.

  • Target Soluble Cytosolic Protein: Collect the supernatant fraction remaining after high-speed centrifugation.

Dialysis

• Purpose and Principles:

  • Dialysis separates small solute molecules from large macromolecules (proteins).

  • Alters sample buffer composition, adjusts pH, removes unwanted small ions/salts, and equilibrates the protein sample in the exact buffer required for subsequent purification steps.

• Procedure:

  • The protein solution is loaded into sealed dialysis tubing made of a semipermeable membrane containing uniform, microscopic pores of defined size (molecular weight cut-off, MWCO).

  • The sealed tubing is submerged in a large volume of dialysate buffer.

  • Small molecules, unwanted ions, and salts diffuse freely across the semipermeable membrane down their concentration gradients until thermodynamic equilibrium is achieved.

  • High molecular weight proteins that exceed the pore dimensions cannot pass through the membrane and are retained inside the dialysis tubing.

  • Replacing the external dialysate buffer repeatedly reduces small solute concentrations inside the tubing to negligible levels.

Protein Separation Properties and Techniques

• Summary of physical and chemical properties utilized for protein separation:

  • Solubility $\rightarrow$ Ammonium sulfate precipitation ("salting out")

  • Size / Molecular Weight $\rightarrow$ Size-exclusion (gel filtration) chromatography, Gel electrophoresis (SDS-PAGE), Ultracentrifugation

  • Net Charge $\rightarrow$ Ion-exchange chromatography, Isoelectric focusing (IEF), Gel electrophoresis

  • Hydrophobicity / Polarity $\rightarrow$ Hydrophobic interaction chromatography (HIC), Reverse-phase chromatography (RP)

  • Function / Binding Specificity $\rightarrow$ Affinity chromatography

Separation by Solubility: Salting Out

• Principles of Salting Out:

  • All proteins exhibit specific solubility limits in aqueous solutions based on the balance of hydrophilic and hydrophobic interactions with surrounding water molecules.

  • Protein solubility can be systematically lowered by increasing the salt concentration of the solution, a process called salting out.

  • Ammonium Sulfate $(NH_4)_2SO_4$ is the standard salt used because of its high solubility in water and high ionic strength.

• Molecular Mechanism:

  • As ammonium sulfate concentration increases, added $NH_4^+$ and $SO_4^{2-}$ ions compete with the protein for interactions with solvent water molecules.

  • The salt ions sequester water molecules to form tight hydration shells around themselves, effectively stripping hydrating water from the protein surface.

  • Loss of surface hydration shells exposes hydrophobic surface patches on the protein.

  • Exposed nonpolar patches aggregate via hydrophobic interactions, causing the protein to aggregate and precipitate out of solution as a solid precipitate.

  • Because different proteins lose their hydration shells at different salt thresholds, target proteins can be selectively precipitated and collected by centrifugation.

Principles of Column Chromatography

• Components of Column Chromatography:

  • Stationary Phase: A solid, porous matrix consisting of synthetic polymer beads or resin packed inside a vertical column.

  • Mobile Phase: A liquid buffer solution containing the protein mixture that flows through the stationary phase packed in the column.

• General Mechanism:

  • The protein sample is applied to the top of the column and mobilized by the continuous flow of mobile phase buffer supplied by a reservoir and pump.

  • Proteins interact differently with the stationary phase based on their specific chemical or physical properties.

  • Proteins with low affinity for the stationary phase move rapidly through the matrix and wash off (elute) first.

  • Proteins with high affinity for the stationary phase are retained longer and elute in later fractions.

  • An automated fraction collector collects sequential liquid fractions at the column exit for subsequent analysis.

Ion-Exchange Chromatography

• Separation Basis: Differences in net surface charge of proteins at a given buffer pH.

• Stationary Phase Types:

  • Cation Exchanger: Column packed with polymer beads containing covalently attached negatively charged functional groups (e.g., carboxymethyl groups). Retains positively charged proteins (cations). Proteins with a net negative charge move faster and elute earlier.

  • Anion Exchanger: Column packed with polymer beads containing covalently attached positively charged functional groups (e.g., diethylaminoethyl groups). Retains negatively charged proteins (anions).

• Elution Protocols:

  • Salt Gradient Elution: An increasing concentration gradient of an ionic salt (e.g., $NaCl$) is passed through the column. Cations ($Na^+$) and anions ($Cl^-$) compete directly with the bound target proteins for charged binding sites on the resin, displacing the bound proteins.

  • pH Shift Elution: Changing the pH of the mobile phase alters the protonation state and net charge of the bound proteins. Extremes of pH should be avoided, as extreme pH can cause irreversible protein denaturation and structural damage.

Hydrophobic Interaction Chromatography (HIC)

• Separation Basis: Differences in exposed nonpolar/hydrophobic amino acid surface patches on proteins.

• Stationary Phase: Polymer beads covalently modified with short hydrophobic functional groups, such as phenyl (benzene) or alkyl chains.

• Binding Conditions:

  • The protein sample is loaded onto the column in a high-salt buffer (high ionic strength).

  • High ionic strength reinforces the hydrophobic effect, driving nonpolar surface patches on the protein to bind tightly to the hydrophobic resin beads.

• Elution Mechanism:

  • The column is washed with a buffer gradient of decreasing salt concentration.

  • Lowering the ionic strength weakens hydrophobic interactions, releasing bound target proteins from the matrix so they elute in order of increasing surface hydrophobicity.

Size-Exclusion (Gel Filtration) Chromatography

• Separation Basis: Differences in molecular size and hydrodynamic shape. No chemical binding to resin occurs.

• Stationary Phase: Polymer beads (e.g., cross-linked dextran or agarose) containing engineered microscopic pores of specific dimensions.

• Separation Mechanism:

  • Large Proteins: Exceeded by the pore dimensions (excluded from pores). They pass freely around the outside of the polymer beads, taking a short, direct path through the column, and elute first.

  • Small Proteins: Enter the labyrinthine internal pore network of the beads. They take a tortuous, longer pathway through the stationary phase, resulting in a delayed transit time, and elute last.

  • Intermediate Proteins: Penetrate a fraction of the bead pores and elute in intermediate fractions between large and small proteins.

Affinity Chromatography

• Separation Basis: Highly specific biological binding interactions between a target protein and a specific ligand.

• Stationary Phase: Polymer beads covalently cross-linked to a specific ligand (e.g., enzyme substrate, cofactor, receptor ligand, antibody, or metal ion).

• Procedure:

  1. Sample Application: The crude protein mixture is loaded onto the column.

  2. Washing: The target protein binds tightly to the immobilised ligand, while all non-binding unwanted proteins flow unimpeded through the column.

  3. Elution: Bound target protein is eluted by passing a solution containing a high concentration of free, soluble ligand. The excess free ligand competes directly with the stationary phase ligand for protein binding sites, displacing the target protein into the mobile phase.

Analytical Separation Techniques

• Purification Scheme Optimization:

  • Complete purification of a target protein typically requires a minimum of $2 ext{--}4$ sequential separation steps (e.g., cell lysis $\rightarrow$ centrifugation $\rightarrow$ ammonium sulfate precipitation $\rightarrow$ ion-exchange chromatography $\rightarrow$ size-exclusion chromatography).

  • Every additional step introduces sample loss; purification schemes are designed to minimize the total number of steps to achieve maximum purity while optimizing overall yield.

• General Electrophoresis Principles:

  • Charged macromolecules migrate through a porous gel matrix under the influence of an applied electric field.

  • Agarose Gel Electrophoresis: Cast horizontally in a buffer chamber. Typically used to separate nucleic acids (DNA/RNA) which possess a uniform negative charge-to-mass ratio and migrate toward the positive anode ($+$). Not suitable for high-resolution protein separation.

  • Polyacrylamide Gel Electrophoresis (PAGE):

  • Polyacrylamide gels are cast vertically between thin glass plates.

  • Acts as a molecular sieve to separate proteins.

Native PAGE vs. SDS-PAGE

• Native PAGE:

  • Separates proteins in their native, fully folded, active state.

  • Maintains protein quaternary structure and non-covalent subunit interactions.

  • Limitations: Protein migration velocity depends simultaneously on three distinct variables: net charge, molecular mass, and native three-dimensional shape. Consequently, two proteins of identical molecular weight can display completely different migration rates if their shapes or surface charges differ.

• SDS-PAGE (Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis):

  • Denaturation Strategy: Eliminates native shape and charge differences so that migration velocity depends solely on molecular mass ($M_r$).

  • Sodium Dodecyl Sulfate (SDS):

  • An amphipathic, strongly negatively charged detergent with the chemical formula $\text{CH}_3(\text{CH}_2)_{11}\text{OSO}_3^-\text{Na}^+$.

  • Unfolds native proteins into linear extended polypeptide chains.

  • SDS binds to unfolded proteins stoichiometrically at a constant mass ratio (approx. $1.4\text{ g SDS}$ per $1.0\text{ g protein}$, or roughly one SDS molecule per two amino acids).

  • Masks intrinsic amino acid side-chain charges, imparting a uniform negative charge density and uniform mass-to-charge ratio across all proteins in the sample.

  • Reducing Agent and Heat Treatment:

  • Samples are mixed with SDS and a reducing agent (e.g., $\beta\text{-mercaptoethanol}$ or dithiothreitol, DTT) and heated at $95^\text{o}\text{C}$ for $5\text{--}10\text{ min}$.

  • Heat and SDS disrupt non-covalent hydrophobic and electrostatic interactions; the reducing agent cleaves covalent disulfide bonds ($-\text{S}-\text{S}-$).

  • Migration Mechanics:

  • Unfolded, negatively charged SDS-protein complexes migrate downward through the vertical polyacrylamide gel toward the positive anode ($+$).

  • The polyacrylamide mesh acts as a physical sieve: small proteins pass easily through the pores and move quickly, while large proteins are retarded by the matrix and move slowly.

  • Applications of SDS-PAGE:

  • Monitoring purity across successive purification steps (a single, clean, crisp band indicates high purity).

  • Estimating the molecular weight ($M_r$) of target proteins by running a lane of standard marker proteins of known molecular weights.

  • Estimating relative protein concentration by comparing staining band intensities.

Isoelectric Focusing (IEF) and 2D Gel Electrophoresis

• Isoelectric Focusing (IEF):

  • Determines the isoelectric point ($pI$) of a protein (the specific pH at which a protein carries a net electrical charge of zero).

  • Mechanism:

  • A gel strip is prepared with an immobilized pH gradient (e.g., $pH\text{ }9$ down to $pH\text{ }3$).

  • Under high pH conditions, carboxylic acid groups ($-\text{COOH}$) are deprotonated ($-\text{COO}^-$), giving proteins a net negative charge.

  • Under low pH conditions, amino groups ($-\text{NH}_2$) are protonated ($-\text{NH}_3^+$), giving proteins a net positive charge.

  • An electric field is applied. Positively charged proteins migrate toward the negative cathode ($-\text{top}$) into regions of higher pH, becoming progressively deprotonated.

  • Negatively charged proteins migrate toward the positive anode ($+\text{bottom}$) into regions of lower pH, becoming progressively protonated.

  • When a protein reaches the pH region corresponding to its exact $pI$, its net charge becomes zero ($\text{net charge} = 0$), electrostatic attraction ceases, and migration stops.

• Two-Dimensional (2D) Gel Electrophoresis:

  • Combines two orthogonal separation techniques sequentially to resolve complex protein mixtures:

  • First Dimension (Horizontal): Isoelectric focusing on a gel strip, separating denatured proteins strictly by their intrinsic $pI$ values.

  • Second Dimension (Vertical): The IEF strip is placed atop a standard SDS-PAGE slab gel and subjected to electrophoresis, separating proteins strictly by their molecular weight ($M_r$).

Quantitating Protein Activity and Specific Activity

• Definitions:

  • Total Activity: The total enzymatic activity units present in a solution volume.

  • Total Protein: The total mass (mg) of all proteins present in the fraction.

  • Specific Activity: The ratio of total enzyme activity units to the total mass of protein:     $\text{Specific Activity} = \frac{\text{Total Activity (units)}}{\text{Total Protein (mg)}}$

• Trends During Purification:

  • As purification progresses through successive steps, Total Protein drops dramatically (as contaminants are removed) and Total Activity decreases slightly (due to unavoidable yield loss).

  • Specific Activity increases progressively with each purification step, serving as the quantitative metric for increasing protein purity.

• Example Purification Table:

Structural Determination Methods

X-Ray Crystallography

• Workflow:

  1. Expression and high-grade purification of large quantities of protein.

  2. Protein crystallization: Slowly precipitating protein out of solution under precise pH, salt, and temperature conditions so water evaporates slowly to form an ordered, 3D crystalline lattice.

  3. Diffraction data collection: X-ray beams strike the crystal lattice, generating a regular pattern of diffracted spots.

  4. Electron density calculation: Fourier transformation converts diffraction spot intensities and phase data into a 3D electron density map.

  5. Model fitting: Amino acid residues are fitted into the electron density map to resolve atomic coordinates.

• Pros:

  • No upper molecular weight / size limits.

  • Highly established technique yielding high-resolution atomic structures.

• Cons:

  • Highly challenging for membrane proteins.

  • Crystal growth is difficult, empirical, and not guaranteed.

  • The static conformation forced by crystal packing may not reflect the active protein conformation in solution or in vivo.

  • Flexible structural loops or disordered regions do not occupy fixed spatial positions within the crystal, resulting in weak, missing, or uninterpretable electron density.

  • Cannot capture dynamic conformational changes in real-time.

Biomolecular NMR (Nuclear Magnetic Resonance) Spectroscopy

• Workflow:

  1. Recombinant expression of protein in media enriched with NMR-active stable isotopes ($^{13}\text{C}$, $^{15}\text{N}$).

  2. Purification of highly concentrated protein solutions.

  3. Collection of 2D/multidimensional NMR spectra (measuring magnetic interactions transferred through chemical bonds or through space between nuclei separated by $< 5\text{ \AA}$).

  4. Assignment of resonance chemical shift cross-peaks.

  5. Calculation of an ensemble of low-energy 3D structures.

• Pros:

  • Requires no protein crystallization.

  • Determines protein structures directly in aqueous solution under physiological conditions.

  • Enables dynamic studies of conformational flexibility, domain motions, and real-time ligand-binding interactions (e.g., induced-fit mechanisms).

• Cons:

  • Inapplicable to insoluble proteins.

  • Strictly limited to relatively small proteins or domains ($< 30\text{--}50\text{ kDa}$).

  • Requires very high protein sample concentrations.

Cryo-Electron Microscopy (Cryo-EM)

• Workflow:

  1. High-grade purification of protein or macromolecular complex.

  2. Application of a minute sample volume ($5\text{--}10\text{ ng}$) onto a specialized micro-perforated grid.

  3. Rapid vitrification (flash-freezing) in liquid ethane to freeze water instantly into non-crystalline amorphous ice.

  4. Sample grid insertion into transmission electron microscope under cryogenic temperatures.

  5. Transmission of high-energy electron beams to record thousands of 2D projection micrographs.

  6. Single-particle alignment, 2D class averaging, and 3D density map reconstruction using computational algorithms.

• Pros:

  • Requires no protein crystallization.

  • Requires minuscule sample quantities ($5\text{--}10\text{ ng}$).

  • Ideally suited for extremely large proteins, multi-subunit assemblies, and macromolecular complexes (e.g., proteasomes, ribosomes).

  • Computationally resolves and classifies multiple distinct conformational states and orientations from a single sample batch.

• Cons:

  • Historically provided lower resolution than X-ray crystallography (though near-atomic resolution is now routinely achievable).

  • Extremely challenging for small, low-molecular-weight proteins ($< 50\text{--}100\text{ kDa}$) due to insufficient image contrast.