Genomics and Proteomics (1-2)

What is SDS-PAGE?

• Sodium Dodecyl Sulfate – Polyacrylamide Gel Electrophoresis.

• A laboratory technique used to separate proteins based on their molecular weight.

Chemical properties of SDS

• Structure:

  • Long hydrophobic alkyl chain (dodecyl, 12 carbons).

  • Negatively charged sulfate head group (–OSO₃⁻) with sodium counterion (Na⁺).

• Type: Anionic surfactant/detergent.

• Amphipathic nature

  • Hydrophobic tail + hydrophilic sulfate head.

  • Allows SDS to dissolve hydrophobic molecules in water (detergent property).

• Ionic character

  • Sulfate group makes SDS strongly negatively charged in aqueous solution.

  • This is what gives proteins a uniform negative charge during SDS-PAGE.

• Denaturing ability

  • SDS disrupts non-covalent bonds (hydrogen bonds, hydrophobic interactions).

  • This unfolds (denatures) proteins, giving them a rod-like shape.

What does SDS do to proteins?

Before SDS

• The protein is in its native folded state.

• Structure stabilized by:

  • Hydrophobic interactions (H in the diagram = hydrophobic patches tucked inside).

  • Charged R-groups (+ and –) interacting on the surface.

• Result: A complex 3D shape, and each protein has its own unique net charge (depending on amino acids).

After SDS

• SDS (a strong anionic detergent) binds along the length of the protein.

• It:

  1. Denatures the protein → unfolds it into a linear chain.

  2. Masks natural charges → coats the protein with negative charges (–).

• Now, all proteins have:

  • Similar rod-like shapes.

  • A uniform negative charge proportional to their length.

How much SDS binds to protein?

About 1.4 g SDS per 1 g of protein, binding uniformly along hydrophobic regions.

What happens if a cell is incubated with SDS?

Membranes dissolve, all proteins are solubilized, and all proteins are coated with many negative charges.

If proteins are denatured with SDS and placed in just an electric field, what happens?

They all move toward the positive pole at the same rate → no size-based separation.

What medium is used to separate proteins of different sizes?

Polyacrylamide gel, which slows large proteins more than small ones.

What is the name of the process combining SDS and polyacrylamide gel?

SDS-PAGE (Sodium Dodecyl Sulfate – Polyacrylamide Gel Electrophoresis).

Why do small proteins move faster in SDS-PAGE?

Because polyacrylamide gel acts like a molecular sieve → small molecules move more easily through pores.

Where do large proteins stay in SDS-PAGE?

Closer to the well (top of the gel), since they move more slowly.

What are the two important outcomes of SDS-PAGE?

• Proteins lose all structure beyond primary structure.

• All proteins have a large negative charge and migrate toward the positive pole.

PTM crosstalk: Biology

• Shows how PTMs occur in biological systems.

• PTMs can:

  • Be added at specific amino acids (red/blue dots).

  • Occur in patterns or multiple sites along a protein.

  • Cause folding/unfolding changes or regulate protein-protein interactions.

PTM crosstalk: proteomic methods

• Methods to detect PTMs using mass spectrometry (MS).

• Workflow: proteins are digested, PTMs are mapped, then peptides are analyzed by MS.

• Specialized MS strategies allow detection of different modifications.

PTM crosstalk:

• PTMs affect protein function, stability, localization, and interactions.

• Experimental strategies:

  • Knockdown/knockout of modified proteins.

  • Enrichment of modified peptides.

  • MS-based functional analysis.

Is MALDI-TOF a soft ionization technique or hard?

• Soft ionization methods use lower energy so that molecules stay largely intact, producing fewer fragments. This is especially important for analyzing large, delicate, or thermally unstable biomolecules (like proteins, peptides, nucleotides).

Two step process of MALDI-TOF

1. sample prep

2. desoprtion

3. Ionization

MALDI-TOF desoprtion process

2. Laser Irradiation

• A pulsed UV laser (e.g., 337 nm nitrogen laser) strikes the matrix crystals.

• The matrix molecules absorb the laser photons because they have strong chromophores.

• The absorbed energy excites the matrix molecules to a high energy state.

3. Desorption (Explosion of Matrix & Analyte)

• Excited matrix molecules rapidly convert absorbed energy into heat and vibrational energy.

• This causes rapid sublimation (solid → gas) of the matrix, carrying analyte molecules with it into the gas phase.

• Importantly, this process is gentle → the analyte is desorbed without fragmenting.

MALDI-TOF sample prep

• The analyte (protein, peptide, etc.) is mixed with a matrix (usually a small organic acid like sinapinic acid or α-cyano-4-hydroxycinnamic acid).

• The mixture is dried on a metal plate, forming matrix crystals with embedded analyte molecules.

• Role of matrix:

  • Absorbs laser energy instead of the fragile biomolecule.

  • Protects analyte from direct laser damage.

  • Helps transfer energy efficiently to analyte during desorption.

MALDI-TOF ionization process

4. Ionization of the Analyte

• During desorption, ionization occurs by proton transfer:

  • Matrix molecules (M) get excited and form [M+H]⁺ ions.

  • These ions then transfer protons to the analyte (A), producing [A+H]⁺.

• Thus, analyte ions are generated in the gas phase, mostly as singly charged ions.

5. Acceleration into the TOF Analyzer

• The generated analyte ions are accelerated in an electric field.

• All ions receive the same kinetic energy (E = qV).

• Since velocity depends on mass-to-charge ratio (m/z), lighter ions travel faster, heavir ions slower.

6. Detection (Time-of-Flight Measurement)

• The TOF tube measures how long ions take to reach the detector.

• Time ∝ √(m/z).

• By recording ion arrival times, the m/z spectrum of the analyte is obtained.

MALDI-TOF: properties of a good matrix

1. Strong absorption at the laser wavelength (usually UV, e.g., 337 nm).

2. Ability to form crystals with the analyte (co-crystallization).

3. Efficient proton donor/acceptor properties (to assist ionization).

4. Volatility so it can vaporize quickly during desorption.

5. Chemical compatibility (should not react with the analyte).

MALDI-TOF: good matrices

• α-Cyano-4-hydroxycinnamic acid (CHCA)

  • Best for small peptides and proteins < 10 kDa.

  • Produces sharp peaks with high resolution.

• Sinapinic Acid (SA)

  • Used for larger proteins (> 20 kDa).

  • Gives good ionization of intact biomolecules.

• 2,5-Dihydroxybenzoic Acid (DHB)

  • Good for carbohydrates and glycoproteins.

  • Produces smoother spectra for complex molecules.

MALDI-TOF:
 laser characteristics

🔹 Characteristics of the Laser

1. Pulsed laser → delivers short bursts of energy (nanosecond pulses).

2. UV wavelength (commonly used):

   • Nitrogen laser: 337 nm

   • Nd:YAG laser: 355 nm (3rd harmonic of Nd:YAG)

   • These wavelengths are strongly absorbed by common matrices.

3. Energy range: Carefully tuned so it excites the matrix but doesn’t directly destroy the analyte.

MALDI-TOF desoprtion process

1. Laser hits matrix crystals

   • The pulsed UV laser excites the matrix molecules (not the analyte directly).

2. Matrix absorbs the energy

   • Because the matrix has strong UV absorption, it takes in most of the photon energy.

3. Rapid heating & sublimation

   • Excited matrix molecules convert photon energy into heat.

   • The solid matrix sublimates (explodes into gas) almost instantly.

4. Analyte carried along

   • As the matrix vaporizes, the analyte molecules (proteins, peptides, etc.) embedded in the crystals are dragged into the gas phase intact.

   • This is the actual desorption part.

MALDI-TOF: why is desoprtion important?

• Desorption ensures large, fragile biomolecules (proteins, nucleic acids) can enter the gas phase without fragmenting.

• The matrix acts like a shock absorber, preventing analyte destruction.

MALDI-TOF: ionization process

1. Proton Transfer from Matrix

   • The matrix molecules, when excited by the laser, can become proton donors ([M+H]⁺).

   • These protons are transferred to the analyte (A).

   • Result: [A+H]⁺ (a protonated analyte ion).

2. Other Ion Forms

   • Sometimes analytes can pick up other cations from the matrix or sample environment:

     • [A+Na]⁺ or [A+K]⁺ (sodium/potassium adducts).

   • But the dominant ion in MALDI is usually [M+H]⁺.

3. Singly Charged Ions

   • Unlike Electrospray Ionization (ESI), which often makes multiply charged ions, MALDI typically produces singly charged ions.

   • This simplifies the spectrum, making it easier to interpret.

MALDI-TOF: ion acceleration

MALDI-TOF: why ion acceleration matters

• Ensures all ions start with the same kinetic energy.

• Creates a direct relationship between m/z and flight time, making TOF analysis possible.

MALDI-TOF: time of flight analyzer

Why is TOF analyzer important?

• Allows analysis of very large molecules (up to hundreds of kDa).

• Unlimited mass range compared to quadrupoles or ion traps.

• High speed and sensitivity.

MALDI-TOF: detector process

1. Arrival of Ions

   • Ions strike the detector surface at the end of the TOF tube.

   • Lighter ions arrive first, heavier ions arrive later.

2. Detector Type

   • Most MALDI-TOF systems use an electron multiplier or a microchannel plate (MCP) detector.

   • When an ion hits, it triggers the release of secondary electrons.

3. Signal Amplification

   • The secondary electrons are multiplied through a cascade process, creating a measurable current pulse.

   • The strength of the signal is proportional to the number of ions arriving.

4. Time Recording

   • The detector records the exact arrival time of each ion.

   • Since the distance and voltage are fixed, arrival time can be directly converted to mass-to-charge ratio (m/z).

What does the MALDI-TOF output show?

1. X-axis: m/z (mass-to-charge ratio)

   • Horizontal axis represents the mass-to-charge ratio of detected ions.

   • Since MALDI produces mostly singly charged ions ([M+H]⁺), m/z ≈ molecular mass of the analyte.

2. Y-axis: Intensity (Relative Abundance)

   • Vertical axis shows signal strength (number of ions hitting the detector).

   • Taller peaks = more abundant ions.

3. Main Peak(s)

   • Represents the intact analyte (e.g., a peptide, protein, polymer).

   • Example: A peptide of 1500 Da will show a main peak near m/z = 1501 ([M+H]⁺).

4. Adduct Peaks

   • Additional peaks can appear due to sodium or potassium adducts: [M+Na]⁺, [M+K]⁺.

5. Matrix Peaks

   • Sometimes, small peaks from the matrix appear at low m/z.

6. Fragmentation (usually minimal)

   • Unlike hard ionization, MALDI produces little fragmentation, so the spectrum is clean and dominated by molecular ions.

ESI: sample prep

1. Analyte in Solution

   • The analyte (proteins, peptides, nucleotides, drugs, metabolites, etc.) is dissolved in a liquid solvent.

   • Typical solvents: mixtures of water + organic solvents (like methanol, acetonitrile).

   • Often a small amount of acid (formic acid, acetic acid) is added to improve protonation.

2. Delivery into the Source

   • The sample solution is fed into a narrow capillary needle (usually stainless steel).

   • A syringe pump or LC (liquid chromatography) system pushes the liquid at a controlled low flow rate (nL/min – µL/min).

3. Continuous Flow System

   • Unlike MALDI (which uses a dried sample spot), ESI is continuous.

   • This makes ESI ideal to couple directly with liquid chromatography (LC-MS), enabling separation and detection in one run.

ESI: set up

🔹 Setup

1. Capillary Needle

   • Very fine metal tube (tens of micrometers wide).

   • The analyte solution flows steadily through it.

2. High Voltage Application

   • A strong electric field is applied between the needle and the counter-electrode (entrance of the mass spectrometer).

   • Voltage: typically 2–5 kilovolts (kV).

   • Polarity can be set to generate positive ions ([M+H]⁺) or negative ions ([M–H]⁻).

What is the effect of a high voltage in ESI set up?

• The strong electric field charges the liquid at the needle tip.

• This creates electrostatic repulsion among the charges in the solution.

• The liquid surface elongates, forming a pointed shape called the Taylor cone.

ESI: Taylor cone formation

1. Electric Field Acts on the Liquid

   • At the needle tip, the high voltage pulls positive (or negative) charges in the solvent toward the surface.

   • Electrostatic repulsion builds up at the liquid–air interface.

2. Cone Shape Formation

   • Normally, liquid at the needle would form a round droplet due to surface tension.

   • But under strong electric fields, electrostatic forces stretch it into a cone shape.

   • This cone is called the Taylor cone (described by Sir Geoffrey Taylor).

3. Jet Emission

   • At the cone’s tip, the forces are unbalanced, so a fine jet of charged liquid is emitted.

   • This jet rapidly breaks up into tiny charged droplets.

Importance of taylor cone (ESI)

• It is the first step of aerosolization — turning a liquid stream into charged droplets.

• The cone ensures droplets are very small, which helps solvent evaporate quickly.

• Without the cone, spraying would be unstable, and ionization efficiency would drop.

ESI: droplet formation

1. Emission of Charged Jet

   • The tip of the Taylor cone ejects a narrow jet of solution.

   • This jet is unstable and rapidly breaks into many droplets.

2. Droplet Charge

   • Because the liquid was under a high electric field, the droplets are highly charged.

   • For positive ion mode: droplets carry excess protons (H⁺).

   • For negative ion mode: droplets carry excess negative charges (e.g., deprotonated species).

3. Droplet Size

   • Droplets are extremely small (sub-micrometer to nanometer scale).

   • Smaller droplets mean higher surface charge density.

ESI: solvent evaporation

1. Droplets Enter Heated Region

   • After leaving the needle, droplets pass through a region with heated drying gas (often nitrogen) and reduced pressure.

   • This encourages solvent evaporation.

2. Droplet Shrinkage

   • As the solvent evaporates, droplets become smaller and smaller.

   • The analyte molecules + charges remain inside, so the charge density increases.

3. Approaching Instability

   • Eventually, the repulsion between like charges inside the droplet becomes too strong for the droplet’s surface tension to contain.

   • At this point, the droplet is unstable and undergoes Coulombic fission (next step).

ESI: importance of solvent evaporation

• Evaporation concentrates charge and brings the system closer to ion release.

• Without solvent evaporation, the analyte would never leave the liquid phase.

• Careful control (temperature, gas flow) is needed so analytes don’t degrade.

ESI: coulumic fission

1. Rayleigh Limit

   • A droplet can only hold a certain amount of charge before it becomes unstable.

   • This limit is called the Rayleigh limit, defined by the balance between:

     • Surface tension (trying to hold droplet together)

     • Coulombic repulsion (charges pushing apart).

2. Droplet Breakup

   • When the Rayleigh limit is exceeded, the droplet undergoes Coulombic fission.

   • It splits into several smaller droplets, each carrying part of the charge.

3. Repeated Fission

   • This process repeats: droplets shrink → charge density rises → fission again.

   • With each cycle, droplets become smaller and more charged.

ESI: Ion Release into Gas Phase

1. Charged Residue Model (CRM)

   • The droplet continues evaporating and fragmenting until it contains just one analyte molecule with leftover charges.

   • Eventually, solvent evaporates completely → leaving a bare charged analyte ion.

   • Works best for large biomolecules like proteins.

2. Ion Evaporation Model (IEM)

   • Before the droplet fully disappears, ions near the surface are ejected directly from the droplet due to strong electrostatic repulsion.

   • Works best for small molecules and salts.

ESI: charged state of ions in ESI and their effect on the mass spectra

🔹 Charge States of Ions in ESI

• Unlike MALDI (mostly singly charged ions), ESI produces multiply charged ions.

• Analyte molecules pick up several protons (or lose several in negative mode).

• Example: a protein might appear as [M+10H]¹⁰⁺, [M+15H]¹⁵⁺, etc.

• Reason: analytes remain in a solvent environment where multiple protonation sites are accessible (amines, carboxyls, etc.).

🔹 Effect on Mass Spectrum

1. Charge State Distribution

   • Large biomolecules produce a series of peaks in the spectrum, each corresponding to a different charge state.

   • Example: a 50 kDa protein may show peaks at m/z 2500, 2000, 1667… (corresponding to +20, +25, +30 charges).

2. Deconvolution

   • Software can convert this charge distribution back into the true molecular weight (M).

   • Formula:

     M=(m/z)×z−z×(mass of proton)M = (m/z) \times z - z \times (mass \, of \, proton)M=(m/z)×z−z×(massofproton)

   • Used repeatedly across charge states to confirm accurate mass.

3. Spectral Features

   • Many closely spaced peaks (charge envelope).

   • Very clean spectra for small molecules (often just [M+H]⁺, [M+Na]⁺).

   • For large proteins: “envelope” pattern of charge states.

How is an IPG gradient created?

• A pH gradient is formed by mixing several specially designed buffering compounds called immobilines.

• Typically, 6–8 different immobilines are used.

• These immobilines are weak acids and bases with well-defined pKa values.

• During gel preparation, these immobilines are co-polymerized with acrylamide, meaning they become chemically bound (immobilized) within the gel matrix.

This immobilization prevents the gradient from drifting during electrophoresis — making the pH gradient permanent and highly stable.

What is the working principle of IPG gradients?

• When an electric field is applied, proteins migrate through the pH gradient.

• Each protein moves until it reaches the region where the pH equals its isoelectric point (pI) — the point where it has no net charge.

• At that position, the protein stops migrating, effectively becoming “focused” into a sharp band.

• Since the pH gradient is chemically fixed in the gel, it remains stable and consistent between runs.

4 advantages of IPG

1. Customizable pH Ranges

   • IPG strips can be formulated for any desired range, commonly between pH 3 to 12.

   • Narrow-range gradients (e.g., pH 4–7) provide higher resolution, ideal for focusing similar proteins.

2. High Reproducibility

   • Because the gradient is covalently linked to the gel, it doesn’t shift or vary between runs — ensuring consistent results.

3. Increased Sample Capacity

   • IPG gels can hold more protein sample compared to traditional carrier ampholyte gels, which makes them ideal for proteomic studies.

4. Improved Focusing and Spot Sharpness

   • Proteins form sharper, more distinct spots due to stable gradients, improving downstream quantification and mass spectrometry.

Step 1 of 2-DE: protein extraction

• Process: Proteins are first extracted (solubilized) from cells or tissues under two different experimental conditions — shown as Condition A and Condition B.

• Purpose: These conditions may represent, for example, a healthy vs diseased state, or treated vs untreated cells.

• Goal: Obtain all proteins in soluble form by using lysis buffers containing detergents, reducing agents, and chaotropic agents.

Step 2 of 2-DE:first dimension

2. First Dimension – Isoelectric Focusing (IEF)

• Principle: Separation of proteins based on their isoelectric point (pI).

• How:

  • Proteins are applied to an immobilized pH gradient (IPG) strip (e.g., pH 3–10).

  • Under an electric field, each protein migrates until it reaches the pH that equals its pI (where its net charge is zero).

• Purpose: Resolve proteins according to their charge differences.

Step 3 of 2-DE: alkylation

• After IEF, proteins are treated with reducing agents (e.g., DTT) to break disulfide bonds, and alkylating agents (e.g., iodoacetamide) to prevent their reformation.

• Reason: This ensures proteins remain fully denatured and separated during the second dimension.

Step 4 of 2-DE: SDS-PAGE

4. Second Dimension – SDS-PAGE

• Principle: Separation by molecular weight (MW).

• How:

  • The IPG strip is placed on top of an SDS-PAGE gel.

  • SDS coats all proteins with a uniform negative charge, so they migrate through the gel solely according to size.

  • Smaller proteins move faster than larger ones.

• Result: A 2D pattern of protein spots, where each spot represents a unique protein species.

Step 5 of 2-DE: staining

5. Staining and Visualization

• Common stains:

  1. Silver stain (high sensitivity)

  2. Coomassie Brilliant Blue (general use)

  3. Fluorescent dyes (for quantitative analysis)

  4. Autoradiography (for radiolabeled proteins)

• Purpose: To visualize the protein spots clearly on the gel.

Step 6 of 2-DE: image analysis

6. Image Analysis

• Step: Compare gels from different conditions (A vs B).

• Goal: Identify protein spots that differ in intensity, presence, or position.

• Interpretation:

  • Upregulated or downregulated proteins indicate differential expression between the two conditions.

Step 7 of 2-DE: spot excisison

7. Spot Excision and Mass Spectrometry (MS)

• Process:

  • Spots of interest are cut (excised) from the gel.

  • Proteins are digested (usually with trypsin) into peptides.

  • Peptides are analyzed using Mass Spectrometry (MS).

• Outcome:

  • Each spot is identified based on peptide mass fingerprinting or MS/MS sequencing.

  • Provides the protein’s identity, sequence, and possible post-translational modifications.

Key steps of sample prep in protein purification 2-DE

Key Steps

1. Lysis buffer — contains urea, thiourea, CHAPS, DTT (denaturants + detergents).

2. Remove salts & nucleic acids — interfere with IEF.

3. Use protease inhibitors — prevent degradation.

4. Protein quantification — Bradford or Lowry assay before loading.

Gel interpretation in 2-DE

Axes of 2-DE Map

• X-axis: pH gradient (acidic → basic)

• Y-axis: Molecular weight (large → small)

Reading the Gel

• Each spot = one protein species.

• Proteins that are upregulated/downregulated between samples → biologically significant.

• Post-translational modifications (PTMs) shift the protein’s pI or MW, creating spot trains.

Advantages of 2-DE

✅ High resolution — separates thousands of proteins.
✅ Detects isoforms and PTMs.
✅ Quantitative (when combined with fluorescent dyes).
✅ Compatible with downstream MS.

Limitations of 2-DE

🚫 Poor for hydrophobic or membrane proteins (don’t solubilize well).
🚫 Extremely large/small proteins may not focus properly.
🚫 Labor-intensive, time-consuming, and requires high-quality sample prep.
🚫 Quantitative analysis is less precise than LC-MS-based proteomics.

MALDI-TOF vs ESI for 2-DE

Why digest proteins into peptides in 2DE?

Why digest?

• Mass spectrometers cannot directly analyze large intact proteins.

• Peptides are smaller, ionize better, and produce characteristic mass spectra that can be matched to databases.

• Each protein yields a unique set of peptide masses → its “peptide mass fingerprint (PMF)”.

Why is trypsin used for protein digestion in 2-DE?

Trypsin

• The gold standard enzyme for proteomics digestion.

• Cleaves specifically at the C-terminal side of Lysine (K) and Arginine (R) residues (unless followed by Proline).

• Produces peptides of ideal length (8–20 amino acids).

• The cleavage pattern is predictable, aiding database matching.

(Optional) Automated In-Gel digestion in 2-DE

⚙ 3. Step-by-Step: Automated In-Gel Digestion Workflow Step 1 – Gel Spot Preparation

• Excise the protein spot with a clean scalpel or automated spot cutter.

• Transfer to a microtube or 96-well plate.

• Wash to remove stains (silver, Coomassie) and residual SDS.

• Dehydrate with acetonitrile (ACN) to shrink the gel piece.

Step 2 – Reduction and Alkylation (Optional but Standard)

• Reduction: Add DTT (Dithiothreitol) to break disulfide bonds (–S–S– → –SH).

• Alkylation: Add iodoacetamide to cap the thiol groups, preventing re-oxidation.

• This ensures complete unfolding and accessibility of the protein to the enzyme.

Step 3 – Rehydration with Trypsin

• Rehydrate the dried gel piece with a small volume of trypsin solution in ammonium bicarbonate buffer.

• The enzyme diffuses into the gel and binds to the protein.

Step 4 – Digestion

• Incubate at 37°C overnight (8–16 hours).

• Trypsin cleaves proteins into peptides within the gel matrix.

• The gel acts as a microreactor.

Step 5 – Peptide Extraction

• After digestion, peptides are extracted using alternating washes of:

  • Aqueous ammonium bicarbonate (to recover hydrophilic peptides),

  • Acetonitrile + formic acid (to recover hydrophobic peptides).

• Combine extracts, then dry (e.g., by vacuum centrifuge).

Step 6 – Sample Cleanup (Optional)

• Desalt using C18 ZipTips or reverse-phase columns to remove salts and contaminants before MS.