Past exam of the biochemistry course of the University of Cambridge / 2025 / Paper 1 / Section I

Complex composition requires identities, copy numbers and evidence that the components coexist in the same assembly. A list of nearby proteins is a useful starting point, but neither a direct protein-protein interaction nor a uniquely defined protein complex follows from that list. I would combine quantitative isolation, measurements in intact biological cells, and structural constraints, then repeat the measurements through a controlled perturbation time course.

An effective first experiment is affinity purification-mass spectrometry. Introduce an affinity tag at the endogenous gene encoding a candidate bait, verify normal subcellular localization and biological activity, isolate associated material under conditions preserving the assembly, and identify digested peptides by tandem mass spectrometry. Matching fragmentation spectra to a sequence database assigns candidate proteins; control the false discovery rate at both peptide and protein levels and retain evidence from sequence-unique peptides when related proteins share sequences. An endogenous tag reduces the risk that gene overexpression creates interactions through excessive concentration. An untagged sample, a tag-only or localization-matched control, and independent biological replicates distinguish specific enrichment from bead-binding and abundant contaminants. Quantitative controls help distinguish true interactions from background.^[ref]

Isolation has selective losses. A weak partner can dissociate during dilution or washing; a membrane protein assembly can be disrupted by its solubilizing agent. Compare extraction conditions, check recovery in the soluble and insoluble fractions, and use reciprocal co-immunoprecipitation from an independent subunit. Treatment with a nuclease tests whether the association depends on DNA or RNA, but removal after extraction can itself destabilize a real nucleic-acid-dependent assembly. Purification from mixed tagged and untagged cell populations, with distinguishable stable isotope labeling, can test whether partners exchange after cell disruption. A reciprocal enrichment supports coassociation; demonstrating a direct contact still requires additional evidence.

For stoichiometry, raw mass spectrometry intensity is insufficient because different peptides have different digestion and ionization efficiencies. Use calibrated peptide standards or carefully normalized quantitative estimates, ideally with several reference subunits. native mass spectrometry measures intact assembly masses and can distinguish candidate subunit counts, while controlled dissociation helps assign subcomplexes. Bound lipids, post-translational modifications, gas-phase losses and heterogeneous occupancy complicate assignment, so check the mass against the summed masses of candidate subunits. Size-exclusion chromatography coupled to multi-angle light scattering independently constrains assembly mass, although an unresolved mixture produces an average and the hydrodynamic size alone is not a molecular mass. Quantitative mass spectrometry of protein complexes can combine interaction strengths, abundance and stoichiometry.^[ref]

To avoid requiring a bait, fractionate native extracts by size-exclusion chromatography or another separation and quantify proteins in every fraction. Correlated elution profiles suggest membership of the same protein complex; comparison across an independent separation helps distinguish coincidence. This co-fractionation mass spectrometry can also reveal shifts from a large assembly into smaller forms after treatment. It may merge different assemblies sharing a subunit, and a failure to coelute does not exclude a transient interaction. chemical cross-linking mass spectrometry adds spatial information: identify pairs of linked residues, translate them into distance restraints appropriate to the reagent and residue accessibility, and ask whether a proposed assembly can satisfy them. A cross-link is a proximity constraint, not automatically a permanent interface; intermolecular assignment is especially difficult for identical subunits.

In living biological cells, proximity labeling captures an interaction neighbourhood before disruptive extraction. A bait fused to TurboID marks nearby proteins with biotin, allowing stringent enrichment and mass spectrometry. The original TurboID work demonstrated useful labeling over roughly ten minutes, which is much faster than older long-labeling implementations.^[ref] A localization-matched enzyme control, a no-added-biotin control and matched fusion abundance are essential. The labeling pulse integrates both spatial proximity and residence time; ten-minute labeling cannot resolve a seconds-long interaction sequence. Changes in ATP, biotin availability, enzyme activity or access to reactive residues can change the labeling signal independently of complex membership. A pulse followed by a chase does not make the covalent label disappear when the partner leaves.

Structural methods distinguish alternative interpretations of these measurements. cryo-electron microscopy can assign ordered subunits, estimate occupancy of resolved features, and separate some compositionally different particle classes. Flexible or small subunits may remain invisible despite being present. X-ray crystallography resolves stable interfaces but selects material that crystallizes. Förster resonance energy transfer between appropriately positioned labels tests nanometre-scale proximity in intact biological cells; single-particle tracking and fluorescence correlation spectroscopy can test comobility and exchange. Label placement, spectral cross-talk and accidental colocalization require controls. None of these techniques by itself reliably inventories every component.

The computational analysis should preserve these distinctions. Construct a protein interaction network with separate edge types for copurification, cross-linking and proximity labeling. Cluster reproducibly supported associations to propose modules, distinguish a stable core from variable accessory components, and integrate localization and measured protein abundance. Models such as AlphaFold 3 can propose interfaces for experimentally supported candidate components, but a confident predicted geometry does not demonstrate that an interaction occurs in the tested condition.^[ref] Test interfaces by site-directed mutagenesis, confirming normal protein folding, expression and localization before interpreting a lost interaction. Evaluate a proposed assembly against independently held-out cross-links or other measurements to reduce circular validation.

For perturbations, use vehicle controls, several doses and early-to-late sampling; randomize preparation and instrument order. Quantify input abundance alongside bait-normalized partner recovery. An illustrative relative recruitment statistic for partner $i$ at time $t$ iswhere $I^{\mathrm{IP}}$ is background-corrected recovered intensity, $b$ is the bait and $A_i$ is input abundance. This is an operational comparison under matched extraction and measurement response, not an absolute occupancy or dissociation constant. A doubled recovered partner signal with doubled input abundance and unchanged bait recovery gives $R_i=1$, illustrating why raw enrichment alone would mislead. Values near the detection limit need uncertainty bounds rather than unstable ratios.

A drug-induced loss of several core components could mean disassembly, degradation, changed localization or poor recovery. Distinguish them with input proteomics, microscopy and native fractionation, then test reversibility after washout and rescue by a drug-resistant target when feasible. For stress, monitor viability and extraction-resistant material; for disease-associated mutations, compare otherwise matched genetic backgrounds and rescue with the normal allele. post-translational modifications may recruit an accessory partner without changing the core. hydrogen-deuterium exchange mass spectrometry can then test interface exposure, but its altered exchange reports accessibility and structural dynamics rather than a new subunit by itself. A convincing remodeling claim is a reproducible change in assembly membership or occupancy, separated from changes in abundance, localization and assay recovery, and connected to a functional consequence.

Dynamic measurements explain which conformations a protein visits, how quickly it exchanges between them, and which transitions control biological activity. A static protein structure identifies possible contacts; protein dynamics can establish whether those contacts form before substrate binding, during enzyme turnover, or when a membrane protein switches between transport states. The useful comparison is therefore among conformational ensembles, transition rates and function under matched conditions.

Consider two interconverting conformations $A$ and $B$. For an ideal two-state system at thermal equilibrium, detailed balance gives $p_A{k}_{AB}=p_B{k}_{BA}$, soHere $G_A,G_B$ are molar Gibbs free energies, $R$ is the molar gas constant, and $T$ is temperature. Measuring only the population ratio cannot identify the exchange timescale: multiplying both rates by the same factor preserves that ratio. Conversely, a measured relaxation rate without populations does not determine the individual forward and reverse rates. Under this model, if $p_B=0.1$ and $k_{\mathrm{ex}}=100{\mathrm{s}}^{-1}$, then $k_{AB}=10{\mathrm{s}}^{-1}$ and $k_{BA}=90{\mathrm{s}}^{-1}$. These illustrative values are a consistency check, not measurements of a particular protein. A driven ATP-consuming cycle need not satisfy detailed balance around the entire cycle.

NMR spectroscopy is particularly powerful for a soluble globular protein. Residue-resolved chemical shifts report local environments; NMR relaxation dispersion measures exchange contributions to line broadening and can detect low-population states exchanging on microsecond-to-millisecond timescales. Different experiments probe faster local fluctuations or slower exchange. Analysis can estimate populations, rates and chemical shift differences, allowing a minor state's environment to be compared with a known ligand-bound state. dihydrofolate reductase provides a concrete example: relaxation-dispersion measurements identified excited conformations resembling states used elsewhere in its catalytic cycle.^[ref] This supports access to functionally relevant conformations before the corresponding chemical step; it does not establish that every detected motion accelerates the bond-making reaction.

This distinction helps test conformational selection against induced fit. If an unbound protein already samples a binding-competent conformation and the ligand stabilizes it, conformational selection contributes. If substantial rearrangement follows initial binding, induced fit contributes. Both can occur in the same mechanism. Measure binding and structural signals over a range of ligand concentrations, include independently measured exchange rates, and fit explicit kinetic models. A matching chemical shift alone does not establish pathway flux, and different models can fit a single relaxation trace. Combining equilibrium populations, rapid-mixing kinetics and site-directed mutagenesis can constrain which route dominates under the tested conditions.

single-molecule Förster resonance energy transfer reveals heterogeneity concealed by ensemble averaging. For a calibrated label pair, the ideal distance dependence iswhere $R_0$ is the Förster distance. Distinct efficiencies and transitions between them can reveal domain closure, opening and dwell times; $r=R_0$ gives $E=1/2$. A distribution of efficiencies is not automatically a distribution of structural states: dye orientation, blinking, label flexibility and photobleaching also contribute. Use donor-only and acceptor-only samples, alternating excitation where appropriate, activity measurements after labeling, and more than one label geometry. For a single memoryless exit with rate $k$, the dwell survival is $e^{-kt}$ and the mean dwell is $1/k$; missed fast events and multiple hidden states violate that simple interpretation.

These measurements connect protein dynamics to allostery. A ligand at one site can change the populations or exchange barriers at a distant functional site, even if the mean protein structure changes little. Measure both sites, compare their thermodynamic coupling and test whether an interface mutation changes coupling while preserving protein folding. hydrogen-deuterium exchange mass spectrometry identifies changes in protection across a protein and is useful for large assemblies. Interpretation needs matched pH and temperature, correction for exchange during analysis, and recognition that protection combines solvent access, hydrogen bonds and structural opening. A protected region can be an interface or a stabilized internal region; it is not direct proof of contact.

For membrane proteins, the environment is part of the experiment. micelles of detergents, lipid bilayer nanodiscs and lipid vesicles can stabilize different conformational ensembles. NMR spectroscopy, electron paramagnetic resonance distance measurements and single-molecule Förster resonance energy transfer can compare cytoplasmic domain movements in these environments. In the beta-2 adrenergic receptor, combined fluorine NMR spectroscopy and double electron-electron resonance showed that agonist occupancy did not fully stabilize one active conformation; interaction with a G protein or a mimetic further favored the active state.^[ref] This explains how ligands with similar binding can differ in signaling efficacy: binding occupancy and population of a productive signaling conformation are distinct quantities.

Ion channels provide an unusually direct connection between molecular switching and function. Patch clamp records individual openings and closures, permitting measurement of conductance, open probability, dwell times and responses to membrane potential or ligand. For $N$ identical independent channels with single-channel current $i$ and open probability $P_o$, the mean current is $Ni{P}_o$. A changed whole-cell current could therefore reflect channel number, conductance or gating; single-channel measurements help separate them. Correlating a conformational probe with electrical activity can assign a structural motion to opening, but a multi-state gating model is usually more realistic than simply equating all structural fluctuations with current-producing transitions. Control membrane integrity, recording bandwidth, label effects and the relevant electrochemical gradient.

For a carrier protein, structural dynamics tests alternating access: the substrate-binding region becomes accessible successively from opposite sides without maintaining an open pore across the lipid bilayer. Measure conformational exchange together with uptake into lipid vesicles, varying substrate, ATP or the driving electrochemical gradient. A conformation trapped by a mutation may be an on-pathway intermediate, an off-pathway dead end or a misfolded state; compare binding, transport and the driving-energy reaction. cryo-electron microscopy classification supplies candidate structural states, but particle-class fractions can be biased by preparation and classification and do not directly provide transition rates. molecular dynamics can suggest connecting routes and transient interactions; predictions must reproduce observables and are limited by sampling and the physical model.

The strongest functional inference comes from perturbing a particular transition and measuring the resulting activity. Preserve abundance, localization, protein folding and binding where possible, then ask whether the measured rate change predicts the functional change. Slow reaction product release may determine enzyme turnover even when chemical conversion is fast; a membrane protein can bind a substrate normally while failing to complete its transport cycle. Biophysical dynamics turns a structural catalogue into a mechanism when state populations, transition rates and activity constrain the same model.