Techniques in Protein Biochemistry: Purify It, Then Prove It

Many of the clinical tests you will order as a physician run on the techniques in this chapter. The pregnancy test detects a hormone with an antibody. The troponin test that confirms a heart attack does the same. The classic confirmatory HIV test was a western blot. Strip away the brand names and the plastic cartridges, and what remains is a small set of methods for finding one protein in a crowd, pulling it out clean, and proving its identity. It starts with a problem of numbers.

A single human cell holds billions of individual protein molecules, but they are drawn from only about ten thousand distinct protein species. Across the whole body, alternative splicing and post-translational modification expand the roughly 20,000 genes into a proteome of more than a million distinct protein forms. The genome is the blueprint. The proteome is the building, lived in and constantly remodeled. To study one protein, you have to get it out of that building alone.

You cannot purify what you cannot find

Before the first spin, before the first column, you need an assay — a way to detect your protein in a tube where you cannot see it. For an enzyme, the assay is usually its own reaction. Lactate dehydrogenase converts pyruvate to lactate using the coenzyme NADH, which is consumed as the reaction runs; NADH absorbs light at 340 nm and lactate does not, so the absorbance falls, and the slope of that fall is the activity. The number that matters is specific activity — units of activity divided by milligrams of total protein. Picture a jar of mixed marbles: the count of red marbles is the activity, but the fraction that are red is the specific activity. A good step throws away other-colored marbles without losing red ones, so the fraction climbs. That single number guides every step that follows.

Four ways to separate a protein from its neighbors

First you have to get the proteins out of the cell. Break the cells open into a homogenate, then spin. Differential centrifugation uses force as a sieve. A gentle spin drops the heavy nuclei; crank it harder and mitochondria pellet; harder still and the smallest vesicles go down, leaving cytosol in the final supernatant. Each spin trades one mixture for two cleaner ones.

What centrifugation cannot finish, chromatography does, by exploiting four different properties of your protein, one at a time.

The first is solubility. Add ammonium sulfate and proteins begin to drop out of solution, but not all at once. Different proteins precipitate at different salt concentrations, so a carefully chosen cut leaves many contaminants behind. A round of dialysis through a semi-permeable membrane then washes the salt back out.

The second is size, and it runs backward from intuition. In gel filtration the column is packed with porous beads. Large proteins are too big to enter the pores, so they skip them and ride the short path between beads — and elute first. Small proteins wander into the pores, take the long way around, and come off last. It is the part students most often get backward.

The third is charge. An ion-exchange column carries fixed charges on its beads, and opposite charges stick. The rule follows from the protein’s pI: below its pI a protein is net positive and binds a negative (cation-exchange) resin; above its pI it is net negative and binds a positive (anion-exchange) resin. Raise the salt or shift the pH and the protein lets go.

The fourth, and most powerful, is binding affinity. Decorate the beads with something only your protein recognizes — a substrate analog, a cofactor, an antibody, a metal that grips a His-tag — and your target binds while everything else flows through. Then displace it with a flood of free ligand. So the choice is simple: if you know a binding partner, run affinity; if you know the size, gel filtration; if you know the pI, ion exchange; if you know nothing yet, salt it out and start narrowing. HPLC speeds any of these up — fine beads under high pressure, paid for with expensive plumbing.

Watch the numbers climb

Walk a crude lysate through five steps and record total protein, total activity, and specific activity at each one. Both totals fall: you lose protein, and you lose some enzyme along the way. Losing some target is the price of enrichment. The number that must rise, every step, is specific activity, because each step strips contaminant faster than it sacrifices target.

Run it and watch the table fill. Specific activity climbs from about 10 to 30,000 units/mg, a three-thousandfold purification. Yield, meanwhile, slides from 100% to roughly a third. The gel tells the same story in pictures: lane one is a smear of forty bands, and by the affinity step a single sharp band remains. That band is your protein, finally alone.

Reading the band: identity, sequence, mass

A pure band is not yet an answer. SDS-PAGE got you there by erasing every difference except mass: the detergent SDS coats the chain at roughly one molecule per two residues, giving every protein the same charge-to-mass ratio, so they sort by size alone and the small ones run fastest. Isoelectric focusing sorts the other way, letting proteins drift through a pH gradient until each stalls at the pH where its net charge is zero — its pI. Run focusing first, then SDS-PAGE at a right angle, and you get a 2D gel: every protein becomes a spot placed by both pI and mass. Compare a healthy proteome to a tumor’s, and a spot that shifts or brightens is a candidate biomarker.

When a protein has no enzymatic activity to assay — a hormone receptor, say — you reach for antibodies. An antibody-coated bead pulls its target out of a mixture (immunoprecipitation). Transfer an SDS-PAGE gel to a membrane, probe it with antibodies, and light up one band by chemiluminescence (the western blot), confirming both identity and size. Coat a plate instead and you have an ELISA, the workhorse of the clinic, with signal proportional to concentration. The same antibody-sandwich logic, in a strip-based lateral-flow format, is the engine inside the pregnancy test and the COVID rapid test.

Identity is not sequence. To read the order of amino acids, Edman degradation clips off the N-terminal residue, identifies it, and repeats. But each cycle is only about 98% efficient, so after fifty rounds you are down near a third of your signal, and the method stalls past roughly fifty residues. The modern answer is mass spectrometry. MALDI-TOF flicks ionized peptides down a flight tube where the heavier ones arrive later, weighing them to a fraction of a dalton. Cut your protein with trypsin, weigh the pieces, and match the mass list against a database — a peptide mass fingerprint. Push further with tandem MS, fragmenting one peptide and reading sequence directly from the spacing between peaks. Frederick Sanger spent twelve years reading the 51 amino acids of insulin by hand; today a 500-residue protein is sequenced in minutes.

What each method actually answers

Every method here answers one of five questions about your protein: is it present, how pure is it, did the last step help, how much did you keep, and is it still intact. Specific activity answers the first four at a glance — the slope you watch climb, step by step, from a smear to a single band. Mass spectrometry and Edman degradation answer the question the table cannot: which protein, exactly, read off one peptide mass or one residue at a time. Find it, enrich it, prove it. That is protein biochemistry, and now it is a few hours of work instead of twelve years.