Unit 5: Analysis of proteins - Subjective Questions

SDS-PAGE stands for Sodium Dodecyl Sulfate - Polyacrylamide Gel Electrophoresis. It is an analytical technique used to separate proteins based on their molecular weight.

Principle:

• SDS is an anionic detergent that binds to proteins in a constant ratio (approximately 1.4 g SDS per gram of protein).
• SDS denatures proteins and coats them with a uniform negative charge, masking their intrinsic charge.
• This ensures that the charge-to-mass ratio becomes nearly identical for all proteins.
• When an electric field is applied, proteins migrate toward the anode (+).
• The polyacrylamide gel acts as a molecular sieve; smaller proteins move faster through the pores, while larger proteins move slower.
• Thus, separation is achieved purely on the basis of molecular size (weight).

SDS plays several critical roles in PAGE:

• Denaturation: SDS disrupts non-covalent interactions (hydrogen bonds, hydrophobic interactions), unfolding proteins into linear polypeptide chains.
• Uniform Negative Charge: SDS binds proteins at a constant ratio (~1.4 g SDS/g protein), imparting a net negative charge proportional to the protein's length.
• Masking Intrinsic Charge: The large negative charge from SDS overrides the protein's natural charge, so separation depends only on size.
• Constant Charge-to-Mass Ratio: This ensures all proteins migrate toward the anode at speeds determined by molecular weight, not charge.

Without SDS, proteins would separate based on both charge and size, making molecular weight determination impossible.

Qualitative analysis answers "which proteins are present?" while quantitative analysis answers "how much protein is present?".

SDS-PAGE uses a discontinuous gel system with two layers:

1. Stacking Gel (Upper Gel):

• Low acrylamide concentration (~4-5%), giving large pores.
• pH ~6.8 (Tris-HCl buffer).
• Function: Concentrates all proteins into a thin, sharp band before they enter the resolving gel, ensuring they start migration from the same point.

2. Resolving Gel (Lower Gel):

• Higher acrylamide concentration (~8-15%), giving smaller pores.
• pH ~8.8 (Tris-HCl buffer).
• Function: Actually separates proteins based on molecular weight through the sieving effect.

The difference in pH and pore size between the two gels creates the stacking effect, resulting in sharper, better-resolved bands.

Polyacrylamide gel is formed by the polymerization of two monomers:

• Acrylamide – forms long polymer chains.
• Bis-acrylamide (N,N'-methylenebisacrylamide) – acts as a cross-linker to form a mesh-like network.

Polymerization Mechanism:

• It is a free-radical polymerization reaction.
• APS (Ammonium Persulfate): Provides free radicals that initiate polymerization.
• TEMED (N,N,N',N'-Tetramethylethylenediamine): Acts as a catalyst that accelerates the formation of free radicals from APS.

Role Summary:

• APS = Initiator (source of free radicals).
• TEMED = Catalyst (stabilizes and accelerates free radical formation).

The pore size of the gel is controlled by the ratio of acrylamide to bis-acrylamide and the total acrylamide concentration.

The molecular weight of an unknown protein is determined by comparing its migration with molecular weight markers (standards) of known sizes.

Procedure:

1. Run the unknown sample alongside a protein ladder (marker) of known molecular weights.
2. Measure the migration distance of each protein band from the top of the resolving gel.
3. Calculate the relative mobility (R_f) for each band:
   
4. Plot a standard curve of (y-axis) versus (x-axis) using the marker proteins.
5. This produces a linear relationship.
6. Determine the of the unknown protein and read the corresponding from the curve.
7. Take the antilog to obtain the molecular weight.

Key relation: There is an inverse linear relationship between and migration distance — smaller proteins migrate farther.

Relative Mobility () is the ratio of the distance traveled by a protein band to the distance traveled by the tracking dye (dye front).

Importance:

• is a normalized, dimensionless value that allows comparison between different gels.
• It corrects for variations in gel length and running time.
• It is used to construct the standard curve ( vs ) for molecular weight determination.
• values range from 0 to 1, where smaller proteins have higher (migrate farther).

Thus, provides a reproducible measure of protein migration essential for accurate molecular weight estimation.

After electrophoresis, protein bands must be stained for visualization. Common methods include:

1. Coomassie Brilliant Blue Staining:

• Most common method; binds to proteins electrostatically.
• Sensitivity: ~50-100 ng of protein.
• Simple, inexpensive, and reproducible.

2. Silver Staining:

• Uses silver ions that bind to proteins and are reduced to metallic silver.
• Sensitivity: ~1-5 ng (very high, ~100x more sensitive than Coomassie).
• More complex and less reproducible for quantification.

3. Fluorescent Staining (e.g., SYPRO Ruby):

• Uses fluorescent dyes detected under UV.
• Sensitivity: ~1-2 ng, with a wide linear range good for quantification.

4. Zinc/Copper (Negative) Staining:

• Stains the gel background, leaving protein bands clear.

Sensitivity Order: Fluorescent ≈ Silver > Coomassie.
Coomassie is preferred for quantitative analysis due to its linear response, while silver is preferred for detecting trace proteins.

Reducing agents such as β-mercaptoethanol (β-ME) and Dithiothreitol (DTT) are added to the sample buffer to ensure complete denaturation.

Role:

• They break disulfide bonds (-S-S-) between and within polypeptide chains.
• This separates proteins into their individual subunits (monomers).
• Ensures proteins are fully linearized so that separation depends only on molecular weight.

Importance:

• Without reducing agents, multi-subunit proteins would remain linked, showing incorrect (higher) molecular weights.
• Reducing conditions provide the true subunit molecular weight.

Reducing vs Non-reducing SDS-PAGE:

• Reducing conditions: Disulfide bonds broken → subunits separated.
• Non-reducing conditions: Disulfide bonds intact → native multimeric structure partly retained.

The SDS-PAGE procedure involves the following steps:

1. Gel Casting:

• Prepare and pour the resolving gel, allow it to polymerize, then pour the stacking gel with a comb to form wells.

2. Sample Preparation:

• Mix protein sample with sample loading buffer (containing SDS, β-mercaptoethanol/DTT, glycerol, bromophenol blue).
• Boil at 95-100°C for 5 minutes to denature proteins.

3. Loading:

• Load samples and a molecular weight marker into the wells.

4. Electrophoresis:

• Fill tank with running buffer (Tris-glycine-SDS).
• Apply a constant voltage/current; proteins migrate toward the anode.

5. Staining:

• After the dye front reaches the bottom, remove the gel and stain with Coomassie Brilliant Blue.

6. Destaining:

• Remove excess dye to reveal distinct protein bands.

7. Analysis:

• Analyze band positions for qualitative identification and band intensity for quantitative estimation.

The sample loading buffer (also called Laemmli buffer) prepares proteins for electrophoresis. Its components and functions are:

• SDS: Denatures proteins and imparts uniform negative charge.
• β-Mercaptoethanol / DTT: Reduces disulfide bonds to separate subunits.
• Glycerol: Increases sample density so it sinks into the wells.
• Bromophenol Blue: A tracking dye that indicates the migration front during the run.
• Tris-HCl (pH 6.8): Maintains proper pH for the stacking process.

Function:

• Ensures proteins are fully denatured and linearized.
• Facilitates easy loading into wells.
• Allows visual monitoring of electrophoresis progress.

Before loading, the sample is typically heated at 95°C to complete denaturation.

Densitometry is a technique used to quantify proteins based on the intensity of stained bands on a gel.

Principle:

• The intensity (optical density) of a protein band is proportional to the amount of protein present (within the linear range of the stain).

Procedure:

1. Capture a digital image of the stained gel using a scanner or gel documentation system.
2. Use image analysis software (e.g., ImageJ, Quantity One) to measure the integrated density (area × intensity) of each band.
3. Run known standards of different concentrations alongside samples.
4. Construct a standard curve of band intensity vs known protein concentration.
5. Determine the unknown protein's concentration by comparing its band intensity to the standard curve.

Applications:

• Comparing relative protein expression levels.
• Estimating purity and concentration.

Limitation: Accuracy depends on staining uniformity and the linear range of the dye.

When to prefer:

• Native PAGE: When protein activity or native conformation must be preserved (e.g., enzyme assays, protein-protein interactions).
• SDS-PAGE: When molecular weight determination, purity checking, or subunit analysis is required.

Several problems may occur during SDS-PAGE:

1. Smiling (Distorted) Bands:

• Cause: Uneven heating, high voltage.
• Remedy: Reduce voltage, ensure proper cooling.

2. Streaking / Smearing:

• Cause: Protein degradation, overloading, insoluble aggregates.
• Remedy: Use fresh sample, reduce load, ensure complete solubilization.

3. Fuzzy / Diffuse Bands:

• Cause: Incomplete polymerization, old buffers, protein degradation.
• Remedy: Use fresh reagents, add protease inhibitors.

4. No Bands / Faint Bands:

• Cause: Insufficient protein, poor staining.
• Remedy: Increase protein amount, use sensitive stain (silver).

5. Skewed / Slanted Bands:

• Cause: Uneven gel polymerization, air bubbles.
• Remedy: Pour gel carefully, avoid bubbles.

6. Vertical Streaking:

• Cause: Precipitated or aggregated proteins.
• Remedy: Centrifuge sample before loading.

The discontinuous buffer system (Laemmli system) uses different buffers and pH values to concentrate proteins into sharp bands.

Components:

• Stacking gel buffer: Tris-HCl, pH 6.8.
• Resolving gel buffer: Tris-HCl, pH 8.8.
• Running buffer: Tris-glycine, pH 8.3.

The Stacking Effect (Mechanism):

1. In the stacking gel (pH 6.8), glycine exists mainly as a neutral zwitterion with low mobility (trailing ion).
2. Chloride ions have high mobility (leading ion).
3. Proteins have intermediate mobility and get sandwiched between the fast chloride ions and slow glycine ions.
4. This creates a moving boundary that compresses proteins into a very thin, concentrated stack.
5. Upon entering the resolving gel (pH 8.8), glycine becomes fully ionized and highly mobile, overtaking the proteins.
6. Proteins then separate purely by size in the resolving gel.

Result: All proteins start migration from the same sharp starting line, producing well-resolved, sharp bands.

Molecular weight markers, also called protein ladders or standards, are mixtures of proteins with known molecular weights run alongside the samples.

Types:

• Pre-stained markers: Already colored, allowing visualization during the run and transfer.
• Unstained markers: Require staining after the run; often more precise.

Why Essential:

• Serve as a reference to estimate the molecular weight of unknown proteins.
• Used to construct the standard curve ( vs ).
• Help monitor the progress and quality of electrophoresis.
• Allow verification of proper separation and resolution.

Without markers, it would be impossible to determine the size of an unknown protein or confirm that separation occurred correctly.

SDS-PAGE is a powerful tool for assessing protein purity through qualitative analysis.

Method:

1. Load the purified protein sample onto the gel alongside markers.
2. After electrophoresis and staining, examine the number of bands.

Interpretation:

• Pure protein: Shows a single sharp band at the expected molecular weight.
• Impure protein: Shows multiple bands, indicating contaminants or degradation products.
• The relative intensity of the target band vs contaminant bands indicates the degree of purity.

Applications:

• Monitoring purification steps (e.g., before and after chromatography).
• Confirming homogeneity of a protein preparation.
• Detecting proteolytic degradation.

Quantitative purity can be estimated by densitometry, calculating the percentage of the target band relative to total protein.

Destaining is the process of removing excess/unbound stain from the gel after staining, so that protein bands become clearly visible against a clear background.

Process (for Coomassie staining):

• The stained gel is immersed in a destaining solution typically containing:
  • Methanol/Ethanol (~40%)
  • Glacial acetic acid (~10%)
  • Distilled water
• The gel is gently agitated and the solution is changed periodically.
• Dye bound to protein bands remains, while dye in the gel background diffuses out.

Why Necessary:

• The entire gel absorbs stain initially, masking the protein bands.
• Destaining removes background staining, increasing contrast.
• Enables clear visualization and accurate densitometric analysis.

Note: Rapid destaining can be aided by including a piece of foam or tissue that absorbs the released dye.

The acrylamide concentration in the resolving gel determines the pore size and thus the range of protein sizes that can be effectively separated.

Relationship:

• Higher acrylamide % → smaller pores → better resolution of small proteins (low MW).
• Lower acrylamide % → larger pores → better resolution of large proteins (high MW).

Typical Gel Concentrations and Separation Ranges:

Gradient Gels:

• A gel with a gradient of acrylamide (e.g., 4-20%) allows separation of a wide range of protein sizes in a single gel.

Significance: Choosing the correct acrylamide concentration is essential for achieving optimal resolution of the proteins of interest.

Step 1: Calculate the Relative Mobility ().

Step 2: Substitute into the standard curve equation.

Step 3: Solve for MW by taking the antilog.

Result: The molecular weight of the protein is approximately 44.7 kDa (assuming MW expressed in kDa).

Note: The exact units depend on how the standard curve was calibrated. This demonstrates how and a standard curve are used together to estimate molecular weight.