Question 1 of 6: Polyacrylamide Gel Electrophoresis (PAGE)

Question 1: Polyacrylamide Gel Electrophoresis (PAGE) (20 marks)

Question text not reproduced: the examination questions are © Engineers and Geoscientists BC. Open the official past paper (linked at the top of this page) to read the question, then follow the worked solution below.

(a) The Premise Behind PAGE

Polyacrylamide gel electrophoresis separates a mixture of protein molecules by driving them, under an applied electric field, through the pores of a crosslinked polyacrylamide gel matrix. The gel behaves as a molecular sieve: the density of crosslinking (set by the acrylamide/bis-acrylamide percentage) fixes an effective pore size, so a protein's rate of migration toward the anode depends on how easily it threads through that mesh. In the SDS-PAGE form of the technique, every polypeptide is coated with a uniform negative charge and unfolded into a similar rod-like shape (see part c), so pore resistance — and hence migration distance — becomes a function of chain length (molecular weight) alone, and larger proteins migrate more slowly than smaller ones. Comparing migration distance to a set of molecular-weight standards run in an adjacent lane therefore lets the analyst estimate an unknown protein's size, assess purity (number of bands), and confirm subunit composition.

(b) Silver Stain vs. Coomassie Brilliant Blue R250

A purified preparation is, by definition, a concentrated single-species (or few-species) sample, so the protein is present in the microgram range readily loaded per lane — well above the detection floor of Coomassie Brilliant Blue R250 (roughly 0.1–1 µg per band). Coomassie is the correct choice here: it is a simple one-step organic-dye stain, inexpensive, fast, and reasonably linear with protein mass over its working range, so band intensity can be used semi-quantitatively to judge purity and relative abundance. Silver staining is roughly 10–100× more sensitive (down to low-nanogram amounts) but is a multi-step, more expensive, more time-consuming protocol that is also less quantitatively linear and more prone to lane-to-lane variability — properties that matter for detecting trace contaminants or very low-abundance species, not for visualizing a purified prep that is already loaded at microgram levels. Silver stain would only be preferred here if the goal were to hunt for faint, low-abundance contaminant bands underneath the dominant IgG bands that Coomassie might miss.

(c) Why SDS Is Integral to PAGE

Sodium dodecyl sulfate is an anionic detergent that binds along the length of a denatured polypeptide backbone at a roughly constant mass ratio (~1.4 g SDS per g protein). This does two things simultaneously: it coats every protein with a large, uniform negative charge that swamps the protein's own intrinsic (variable) charge, and, combined with heat, it unfolds the polypeptide into an extended, rod-like conformation that erases shape differences between proteins. With charge and shape standardized across the whole sample, electrophoretic mobility through the polyacrylamide sieve is governed almost entirely by chain length (molecular weight) — which is exactly what turns PAGE into a reliable size-separation and size-estimation tool. Without SDS (native PAGE), migration reflects an uncontrolled combination of size, shape, and native charge, which cannot be interpreted as a simple molecular-weight ladder.

(d) Two Bands: Preparation Steps and What They Reveal

An intact IgG antibody is a Y-shaped heterotetramer held together by inter-chain disulfide bonds: two identical heavy chains (~50 kDa each) and two identical light chains (~25 kDa each), for an intact mass of roughly 150 kDa. To obtain exactly two dark bands in one lane, the sample was run under reducing, denaturing SDS-PAGE: SDS plus a reducing agent (β-mercaptoethanol or DTT) and heat. The reducing agent cleaves the inter-chain (and intra-chain) disulfide bonds holding the four subunits together, while SDS coats and linearizes each freed chain. Because the two heavy chains are identical to each other, and the two light chains are identical to each other, the dissociated antibody resolves into exactly two distinct size classes — a heavy-chain band near ~50 kDa and a light-chain band near ~25 kDa — rather than four separate bands or a single intact-antibody band.

The fact that only two clean bands appear, at positions consistent with canonical heavy- and light-chain sizes, tells the analyst that the starting purified material was a single, homogeneous IgG species with the expected canonical two-heavy/two-light architecture — not a mixture of different antibody clones or isotypes (which would add extra heavy- or light-chain size variants), and not a degraded or proteolytically clipped preparation (which would add smaller fragment bands). It confirms both the purity and the structural integrity of the purified antibody.

(e) An Alternative Protocol Giving a Different Banding Pattern

Running the identical purified sample under non-reducing SDS-PAGE (SDS present, but no β-mercaptoethanol or DTT, and no heat-driven disulfide disruption) would give a markedly different result: the inter-chain disulfide bonds stay intact, so the antibody denatures and unfolds under SDS but remains covalently linked as one ~150 kDa species, producing a single band near 150 kDa (Lane A in Fig. 1) instead of the two bands seen under reducing conditions. Other protocols that would also change the pattern include limited proteolytic digestion (e.g., papain, which cleaves IgG into two Fab fragments and one Fc fragment before reducing SDS-PAGE, giving additional ~25 kDa Fab/Fc-related bands distinct from the intact heavy/light pattern) or running the sample as native PAGE (no SDS at all), where migration would instead depend on the antibody's native folded size, shape, and net surface charge rather than chain length alone.