Spectroscopy for biomolecules

Why it matters

Before spending weeks on a crystal structure, a biophysicist wants quick answers: Is my protein folded? Did the mutation change its structure? Does it unfold at 50 °C or 70 °C? Does the ligand bind? Spectroscopy answers these questions in minutes to hours, using small amounts of protein. The three workhorse methods are circular dichroism (CD), fluorescence and nuclear magnetic resonance (NMR). Each sees something different, so the skill is choosing the right one for the question.

Absorbance first

Proteins absorb UV light at 280 nm (mainly from tryptophan and tyrosine side chains) and strongly below 230 nm (the peptide backbone). The Beer–Lambert law, A = ε · c · l, turns A280 into a concentration using the protein's extinction coefficient ε, which can be calculated from its sequence. Nearly every spectroscopy experiment starts with an accurate concentration.

Circular dichroism: secondary structure

Chiral molecules absorb left- and right-circularly polarised light slightly differently. In the far-UV (about 190–250 nm), the protein backbone's arrangement gives characteristic CD spectra:

• Alpha helix: negative bands at about 222 nm and 208 nm, positive near 193 nm.
• Beta sheet: a negative band near 216–218 nm and a positive band near 195 nm.
• Disordered: a strong negative band near 200 nm.

Uses:

• Confirm a purified or mutant protein is folded and estimate its secondary structure content.
• Thermal melts: follow the 222 nm signal while heating to find the melting temperature Tm, where half the protein is unfolded.
• Compare buffers or formulations for stability.

Limits: CD gives an average over the whole molecule, not residue-level detail. Buffers that absorb strongly in the far UV (high chloride, imidazole) spoil the signal. Near-UV CD (250–320 nm) reports on the environment of aromatic side chains and so on tertiary structure.

Fluorescence: sensitive reporters

Fluorescence measures light re-emitted after absorption, and it is very sensitive (micromolar to nanomolar protein).

• Intrinsic tryptophan fluorescence: excite at about 295 nm; emission peaks near 330–340 nm when Trp is buried in a nonpolar core, and shifts toward 350 nm (red shift) when the protein unfolds and Trp meets water. Ligand binding can quench or enhance it.
• Extrinsic dyes: dyes such as SYPRO Orange fluoresce when they bind exposed hydrophobic patches, which is the basis of differential scanning fluorimetry (DSF, thermal shift), a fast way to screen buffers or ligands in plate readers. Thioflavin T reports amyloid fibrils.
• Fluorescence anisotropy/polarisation: a small labelled ligand tumbles fast (low anisotropy) until it binds a big protein (high anisotropy), giving binding curves.
• FRET: energy transfer between two dyes reports distances of roughly 2–10 nm, used for conformational changes and interactions.

Limits: you see the reporter, not the whole protein. Inner-filter effects at high absorbance and photobleaching can distort results.

NMR: atomic detail in solution

NMR detects nuclei with spin (¹H, ¹⁵N, ¹³C) in a strong magnetic field. For proteins, the standard experiment is the ¹H–¹⁵N HSQC, which shows roughly one peak per backbone amide, a "fingerprint" of the folded protein.

Uses:

• Check whether a protein is folded (well-dispersed peaks) or disordered (peaks bunched together).
• Map binding sites: peaks that move (chemical shift perturbation) when a ligand is added identify the residues involved, even for weak binders.
• Study dynamics and disordered proteins, which crystallography and cryo-EM struggle with.
• Determine full structures of small proteins (typically below about 30–40 kDa).

Limits: needs isotope-labelled protein (grown on ¹⁵N/¹³C media), relatively high concentrations (often 0.1–1 mM), expensive high-field instruments, and specialist training. Large proteins give broad, overlapping peaks.

Choosing a method

Question                                          First choice
Is it folded? What secondary structure?           far-UV CD
At what temperature does it unfold? (one protein) CD thermal melt
Screen 96 buffers or ligands for stability        DSF (thermal shift, fluorescence)
Does a labelled peptide bind? What Kd?            fluorescence anisotropy
Which residues does a weak fragment touch?        NMR (HSQC chemical shift mapping)
Dynamics or disorder at residue level             NMR

Worked example

A lab makes a point mutant of an enzyme and the activity drops tenfold. Is the mutant misfolded, or is the mutation in the active site?

1. Far-UV CD of wild type and mutant: identical spectra with minima at 208 and 222 nm, so the overall helical fold is intact.
2. CD melt: Tm falls from 62 °C to 58 °C, a modest destabilisation, not unfolding at assay temperature (25 °C).
3. Conclusion: the mutant is folded; the activity loss is more likely due to the active site. If the lab had ¹⁵N-labelled protein, an HSQC overlay would show which residues shifted.

Safety

Instruments use intense UV lamps, lasers and (for NMR) very strong magnetic fields that are dangerous to people with implants and to anything ferromagnetic nearby. Operate them only after your facility's training.

Common mistakes

• Using CD buffers with high chloride or imidazole and blaming the protein for noise.
• Reading a DSF Tm as an absolute stability value. It is excellent for comparison, but depends on the dye and heating rate.
• Expecting NMR structures of large complexes. That is cryo-EM territory.

Recap

• CD: secondary structure and thermal stability.
• Fluorescence: sensitive reporters of environment, binding, distance and stability screens.
• NMR: residue-level binding, dynamics and small-protein structures, at higher cost.

Try it

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