Cell lysis disrupts cellular structure to release proteins. Three main approaches:

1. Chemical Lysis:

• Uses detergents and buffers
• Example: SDS disrupts hydrophobic interactions among membrane lipids
• Gentle, but may interfere with downstream analysis

2. Enzymatic Lysis:

• Uses specific enzymes to digest cell walls or extracellular matrix
• Examples: Lysozyme (bacteria), Zymolyase (yeast)
• Specific and gentle

3. Physical Lysis:

After lysis: Centrifugation separates debris from soluble proteins (supernatant).

Both are pre-analytical complexity management steps to reduce sample complexity and compress dynamic range.

Depletion:

• Purpose: Remove high-abundance proteins that mask low-abundance ones
• When used: Essential for plasma/serum (albumin = ~60% of protein)
• Methods: Immunoaffinity columns, protein A/G

Enrichment:

• Purpose: Isolate specific sub-proteomes of interest
• Methods:
  • Selective Dialysis: Membrane with tiny pores acts as sieve
  • Microdialysis: Collect small molecules through diffusion
  • Selective Precipitation: Salts/solvents isolate by solubility
  • Immunoprecipitation: Antibodies isolate target protein

Approach-specific needs:

• Bottom-up: Complexity reduced physically on 2D gel
• Shotgun & Top-down: Complexity must be managed strictly during extraction to avoid overloading LC-MS

Final steps of sample preparation to ensure proteins remain denatured and accessible to trypsin.

Reduction:

• Reagent: DTT (dithiothreitol) or TCEP
• Purpose: Break disulfide bonds (S-S → SH + SH)
• Unfolds protein structure

Alkylation:

• Reagent: IAA (iodoacetamide) or IAM
• Purpose: Block free thiol groups (prevents disulfide reformation)
• Adds ~57 Da (carbamidomethyl) to each cysteine

Why important:

• Ensures complete denaturation
• Makes all sites accessible to trypsin
• Prevents protein refolding/aggregation
• Produces reproducible digestion

Approach differences:

• Bottom-up: Essential for proper IEF/SDS-PAGE
• Shotgun: Essential for making protein accessible to trypsin
• Top-down: Alkylation often skipped to measure true intact mass

Five main goals:

1. Solubilize all protein classes reproducibly
   • Including hydrophobic membrane proteins
   • Use chaotropes (urea, thiourea) to disrupt hydrogen bonds
2. Prevent protein aggregation
   • Keep solubility high during IEF or digestion
   • Use appropriate detergents
3. Prevent chemical/enzymatic modifications
   • Use protease inhibitors
   • Work at low temperature
4. Remove interfering molecules
   • Digest or remove: nucleic acids, salts, lipids
5. Enrich target proteins
   • Reduce dynamic range
   • Deplete high-abundance proteins

2D-PAGE separates proteins by TWO independent (orthogonal) properties for maximum resolution.

First Dimension: Isoelectric Focusing (IEF)

• Separates by isoelectric point (pI)
• Uses immobilized pH gradient (IPG) strip
• High voltage applied
• Positively charged proteins → cathode
• Negatively charged proteins → anode
• Each protein migrates until net charge = 0 (at its pI)
• Result: Proteins aligned horizontally by pI

Second Dimension: SDS-PAGE

• Separates by molecular weight (MW)
• IPG strip placed on top of polyacrylamide gel
• SDS denatures and gives uniform negative charge
• Smaller proteins migrate faster
• Result: Horizontal band resolved vertically

Final result: 2D map of spots — each spot = specific protein with unique pI and MW.

The resolution of IEF depends on the pH range of the IPG strip:

Why?

• Wide range: Same physical strip length covers more pH units → proteins with similar pI hard to distinguish
• Narrow range: Same length covers fewer pH units → better separation of proteins with close pI values

Strategy:

1. Start with wide range (pH 3-10) for overview
2. Use narrow range strips to "zoom in" on regions of interest

Common staining methods:

Silver staining is the most sensitive method, capable of detecting very low-abundance proteins.

*Silver staining compatibility with MS depends on the protocol — some fixatives can interfere.

After staining:

1. Gel is digitized (scanner or camera)
2. Image imported to software (e.g., Melanie)
3. Spot detection and analysis performed

Master Gel: A synthetic reference map created from multiple gel replicates.

How it's created:

1. Run multiple replicates of the same sample
2. Use image alignment (matching) software
3. Apply warping algorithms to correct geometric distortions
4. Combine all spots detected across all gels

What it contains:

• Every spot detected across the entire experiment
• Characterizes a "typical profile"
• Assigns unique coordinates to each protein

How it's used:

• Reference for comparing samples (e.g., healthy vs. diseased)
• Enables consistent spot identification across experiments
• Facilitates quantitative comparison

Software features:

• Contrast adjustment
• Background subtraction
• 3D visualization
• Spot detection and splitting

Sample-Related Limitations:

• Hydrophobic proteins: Membrane proteins poorly soluble in IEF buffers
• Extreme pI: Very acidic (<3) or basic (>10) proteins hard to focus
• Extreme MW: Large (>200 kDa) don't enter gel; small (<10 kDa) run off
• Low-abundance proteins: Masked by high-abundance proteins
• Limited dynamic range: ~10⁴ vs. proteome range of 10⁶-10⁷

Technical Limitations:

• Poor reproducibility: Gel-to-gel variation requires triplicates
• Labor-intensive: Manual, time-consuming, hard to automate
• Low throughput: Cannot be easily scaled
• Co-migration: Similar pI/MW proteins in same spot

Practical Issues:

• Keratin contamination (especially manual spot picking)
• Streaking from degradation
• Background from staining

Trypsin Specificity:

• Cleaves at the C-terminal side of Lysine (K) and Arginine (R)
• Exception: Does NOT cleave when followed by Proline (P)

Why it's the gold standard:

1. Robustness: Stable and active across wide pH and temperature range
2. High Specificity: Predictable cleavage sites enable accurate database searching
3. Ideal Peptide Length: Generates peptides of 6-20 amino acids — optimal for MS detection
4. Internal Calibration: Autolysis peaks (trypsin digesting itself) serve as mass standards
5. Basic C-terminus: K and R promote ionization in positive mode

When to use alternatives:

• Proteins rich in K/R → use Glu-C (cleaves after Glu) for longer peptides
• Different sequence coverage needed → Chymotrypsin (cleaves after Phe, Tyr, Trp)

Bottom-up digestion:

• Called "in-gel digestion"
• Spot excised, destained, then digested
• Peptides extracted from gel

Shotgun digestion:

• Called "in-solution digestion"
• Whole lysate digested
• Produces complex peptide mixture

ZipTip: A 10 µL pipette tip packed with reverse-phase (RP) material.

Purpose:

• Desalt peptides (remove salts that interfere with ionization)
• Concentrate samples
• Remove detergents and buffers

How it works:

1. Condition tip with solvent
2. Bind peptides to RP material
3. Wash away salts (they don't bind)
4. Elute clean, concentrated peptides

When used:

• Bottom-up (gel-based): Preferred offline method for cleaning peptides from single gel spot
• Before MALDI-TOF analysis
• Improves MS sensitivity for low-abundance proteins

Shotgun & Top-down: Use online RP-HPLC instead (performs both desalting and high-resolution separation).

Reverse-Phase (RP) Chromatography: The dominant mode for peptide separation in proteomics.

Why "reverse-phase"?

• Normal-phase: Polar stationary phase, non-polar mobile phase
• Reverse-phase: Non-polar (hydrophobic) stationary phase, polar mobile phase
• It's the "reverse" of traditional chromatography

How it works:

• Stationary phase: C18 hydrocarbon chains (hydrophobic)
• Mobile phase: Water/acetonitrile gradient
• Peptides bind via hydrophobic interactions
• Increasing organic solvent elutes more hydrophobic peptides

Use in proteomics:

Summary:

• MALDI: Favored for speed, simplicity, and tolerance to contaminants
• ESI: Gold standard for high-sensitivity proteomics and complex LC-MS/MS analyses

PMF: Protein identification technique based on the mass spectrum of proteolytic peptides.

Principle: Each protein produces a unique "fingerprint" of peptide masses when digested with a specific enzyme.

Complete workflow:

1. Spot Recovery: Excise protein spot from 2D gel (robotic or manual)
2. Destaining: Remove Coomassie or silver stain
3. Reduction/Alkylation: Break disulfide bonds, block cysteines
4. In-gel Digestion: Trypsin digestion overnight
5. Peptide Extraction: Recover peptides from gel pieces
6. Cleanup: ZipTip desalting
7. MALDI-TOF Analysis: Acquire mass spectrum
8. Database Search:
   • Compare experimental masses to theoretical "digital digests"
   • Databases: UniProt, Swiss-Prot
   • Software assigns Mascot score (statistical probability)

Identification criteria:

• Significant number of peptides must match
• Typically need 4-6 matching peptides
• ~40% sequence coverage considered good

Limitation: Only works if protein is in database.