Exploring Biotechnology with GFP (Part 2) Quiz Questions

Size-exclusion Chromatography

This technique separates molecules by their size in solution (hydrodynamic radius), not by chemical binding. The column is packed with porous beads, and the pores have a defined size range.

Large molecules are too big to enter the pores. They travel only through the spaces between beads, take the shortest path, and elute first.

Small molecules diffuse in and out of the pores. That detour makes their path longer and slower, so they elute later.

Intermediate molecules enter some pores partially and elute in between.

Nothing sticks to the matrix, so the buffer just carries everything through, and the order of elution is roughly the reverse of the order of size. In the lab, GFP (~27 kDa) is separated from larger and smaller contaminants in the cell lysate, and you track it by its glow under long-wave UV. The matrix is swelled beforehand because dry beads have collapsed pores. Hydration opens them to their working size and keeps the bed from changing volume (and cracking) after packing.

470 nm

GFP's chromophore (the Ser-Tyr-Gly tripeptide at residues 65-67) absorbs blue light. Wild-type GFP has a major absorption peak near 395 nm and a smaller one around 475 nm, and it then emits green light at about 509 nm. Reading at this wavelength has two advantages:

It is specific to GFP. Most proteins absorb at 280 nm (from Trp and Tyr), and nucleic acids at 260 nm, so measuring at 280 would pick up everything in your fractions. Almost nothing else in the lysate absorbs at this wavelength.

Absorbance at this wavelength reports on properly folded, fluorescent GFP, since the chromophore only forms and functions when the protein folds correctly. Excitation with light at this wavelength results in stable fluorescence.

Beer-Lambert Law

It relates how much light a solution absorbs to the concentration of the absorbing species: A = ε · l · c

A = absorbance (unitless)

ε = molar extinction coefficient (M⁻¹cm⁻¹), a property of the molecule at a given wavelength (the manual gives 7000 for GFP at 470 nm)

l = path length of light through the sample (cm)

c = concentration (M)

SDS-PAGE (Sodium Dodecyl Sulfate [SDS]-Polyacrylamide Gel Electrophoresis)

This separates proteins by molecular weight only.

Denaturation and uniform charge: SDS unfolds proteins and coats them with negative charge in proportion to their length, so every protein has about the same charge-to-mass ratio. Their natural charge and shape no longer matter.

The gel is a molecular mesh: Polyacrylamide forms a cross-linked network of pores. The acrylamide percentage sets the pore size.

Electrophoresis: when voltage is applied, the negatively charged proteins migrate toward the positive electrode (red, bottom). Small proteins thread through the mesh easily and travel far, while large proteins are slowed down and stay near the top.

Result: The distance migrated is roughly linear with the log of molecular weight, so you plot migration distance (or Rf) against MW on a semilog graph using the standards, then read off the unknowns.

SDS (sodium dodecyl sulfate), 2-Mercaptoethanol (BME or β-ME), and Loading Dye

What is in the denaturing solution?

SDS (sodium dodecyl sulfate)

An anionic detergent that disrupts non-covalent interactions, unfolds the protein, and coats it with a negative charge (about 1.4 g SDS per g protein). This makes migration depend on size alone.

2-Mercaptoethanol (BME or β-ME)

A reducing agent that breaks disulfide bonds between cysteines. This separates protein subunits and fully linearizes the protein, so it doesn't run as a compact, partly folded structure.

Loading Dye

This typically contains a tracking dye (bromophenol blue, the blue band you watch) and a dense component like glycerol or sucrose. The density makes the sample sink into the well instead of diffusing into the running buffer. The dye is small and runs ahead of the proteins, so it shows you when to stop the run (the manual says when it's a couple of centimeters from the bottom).

Staining

Fixation: The staining solution typically contains an alcohol (like methanol or ethanol) and an acid (like acetic acid). This mixture denatures the separated proteins and precipitates them, trapping them firmly in place so they cannot diffuse out of the gel.

Electrostatic and Hydrophobic Binding: Under the acidic conditions of the stain, Coomassie dye molecules become negatively charged (anionic). They bind tightly to positively charged basic amino acids (such as arginine, lysine, and histidine) through electrostatic interactions, as well as hydrophobic residues via van der Waals forces.

Color Change: When the dye binds to proteins, its absorption spectrum shifts, turning the dye from a dull reddish-brown (free solution) to an intense, visible blue complex.

Destaining

The dye also soaks into the gel itself, giving a uniform blue background. It is only loosely held there, whereas protein-bound dye is held tightly.

When you replace the stain with a dye-free destain solution (similar to methanol/acetic acid), unbound dye diffuses out into the solution, while dye bound to protein stays put. Changing the solution repeatedly keeps the concentration gradient steep so the background clears faster, and the paper towel in the tray soaks up free dye.

The bands then stand out against a clear background. Leaving the gel in destain too long will eventually pull dye off the bands too, which is why the manual says not to over-stain or over-destain.