Past exam of the biochemistry and Molecular Biology course of the University of Cambridge / 2025 / Paper 2 / Section B

The catalytic serine residue supplies a nucleophile: its side-chain oxygen attacks the carbonyl carbon of the beta-lactam ring. This generates a tetrahedral intermediate, followed by opening of the ring and formation of a covalent acyl-enzyme intermediate. Subsequent attack by activated water hydrolyzes that intermediate and regenerates the enzyme. The serine residue performs covalent catalysis and is regenerated, rather than being consumed as a reagent.

An appropriately positioned general base helps remove the proton from the serine residue hydroxyl, increasing its nucleophilicity. A lysine residue, such as Lys73 in the class A beta-lactamase catalytic network, is a plausible participant; its protonation state depends on its active-site environment and interactions with other residues. A glutamate residue, notably Glu166, also participates in proton transfer and activation of the water needed for deacylation. Efficient catalysis requires proton-transfer chemistry as well as positioning the attacking oxygen. The precise division of roles during acylation depends on the enzyme state, so the drawing alone cannot assign a complete mechanism to one residue.^[ref]

Bond c, the carbonyl-carbon–nitrogen bond within the four-membered beta-lactam ring, is cleaved. Hydrolysis opens the cyclic amide; it does not cut the side-chain amide or a carbon–carbon bond. The resulting open-ring product no longer has the intact beta-lactam pharmacophore required for its usual antibacterial action.

The compound imidazole resembles the metal-coordinating part of the histidine side chain and competes for binding to the nickel sites in immobilized metal affinity chromatography. At the lower concentration in the wash, it helps displace weakly or non-specifically bound proteins while the multiple contacts of the histidine tag retain the desired enzyme. At the higher elution concentration, competition is strong enough to displace the tagged enzyme. Low imidazole improves washing; high imidazole elutes the target. Remove or match the elution-buffer components before interpreting downstream assays, and verify purity independently of total protein concentration.

The dilution in the Bradford assay is $1000/5=200$. Read the fitted line rather than assuming it passes through the origin: the two absorbances correspond approximately to $4.3$ and $15\mu \mathrm{g}\mathrm{m}{\mathrm{L}}^{-1}$ in the assay mixture. ThereforeMultiplying by the $0.500\mathrm{mL}$ eluate volume gives about $0.43\mathrm{mg}$ of TEM-1 and $1.5\mathrm{mg}$ of TEM-52. Only TEM-52 meets the requirement.

Assuming the same recovery per culture volume, the required volumes areUse approximately $120\mathrm{mL}$ or more for TEM-1; the existing $50\mathrm{mL}$ is sufficient for TEM-52. Rounding the former up to $125\mathrm{mL}$ is sensible given graphical precision. Reading the lower standard as $4$–$4.5\mu \mathrm{g}\mathrm{m}{\mathrm{L}}^{-1}$ gives a corresponding requirement of roughly $110$–$125\mathrm{mL}$, not an exact integer volume. These estimates assume comparable dye response to bovine serum albumin, suitable buffer blanks and that recovered protein is the purified target; contaminants would inflate the inferred target yield.

For the Hanes-Woolf plot, the slope is $1/V_{\max }$, the vertical intercept is $K_m/V_{\max }$, and the horizontal intercept is $-K_m$. The lines have approximately equal slopes, $0.50\min \mu {\mathrm{M}}^{-1}$. The dashed TEM-1 line intercepts the vertical axis at about $13\min$, while the dotted TEM-52 line intercepts it at about $2.5\min$. ThusThe negative-axis intercepts independently give approximately $-26$ and $-5\mu \mathrm{M}$. Allow normal graph-reading uncertainty, especially at the smaller intercept. The limiting rates are similar; the Michaelis constant is substantially lower for TEM-52. A Michaelis constant is not generally a binding dissociation constant, and calculating a turnover number additionally requires the concentration of active enzyme.

The displayed separation between the two entrance landmarks increases from approximately $6.2$ to $7.7\text{A˚}$ in TEM-52, indicating a wider opening. The Gly238-to-Ser substitution can alter backbone packing and loop geometry; its effect is not simply the steric size of the new side chain. Increased accessibility or altered productive positioning could increase the effective association rate for penicillin G, producing a lower apparent Michaelis constant without changing the limiting catalytic step. In a simple Michaelis-Menten kinetics scheme, $K_m=(k_{-1}+k_{\mathrm{cat}})/k_1$, so increasing $k_1$ can lower $K_m$.

The structural change is consistent with improved access and the lower measured $K_m$, while similar $V_{\max }$ suggests similar limiting turnover under these conditions. It does not uniquely establish the mechanism or prove tighter equilibrium binding. TEM-52 contains additional substitutions, so a matched single-substitution series would be needed to isolate the contribution of residue 238. Structural work on TEM-52 independently supports the importance of remodeling access to its active site.^[ref]

The bulky substituents of ceftazidime can hinder productive entry into the more restricted TEM-1 active site. A wider entrance in TEM-52 could accommodate this substrate and allow its beta-lactam ring to be hydrolyzed. The prediction is reduced susceptibility of TEM-52-expressing bacteria compared with matched TEM-1-expressing bacteria. This is a mechanistic prediction, not a quantitative conclusion from the penicillin G data: entry alone does not guarantee efficient catalytic chemistry for a different substrate. Test ceftazidime hydrolysis directly and measure its minimum inhibitory concentration in an otherwise matched bacterial background, controlling enzyme expression, growth, drug entry and other resistance mechanisms.

The ADP addition containsDuring the initial stimulated interval, the plotted oxygen concentration falls from about $240$ to $207\mu \mathrm{M}$. The chamber volume after this addition is $1.900+0.050+0.050+0.010=2.010\mathrm{mL}$, givingThe conventional phosphorylation-to-oxidation ratio counts ATP formed per oxygen atom reduced, so its denominator is twice the number of oxygen molecules:This assumes the added ADP is phosphorylated and uses the displayed total oxygen loss over the pulse; it is an apparent experimental ratio. A separate correction for background respiration or proton leak would change the denominator and must be specified explicitly. Dividing by oxygen molecules without the factor of two would instead give an ATP:${\mathrm{O}}_2$ ratio near $3$.

Succinate is the most plausible substrate. Its oxidation by succinate dehydrogenase, or respiratory complex II, supplies electrons to the ubiquinone pool, bypassing respiratory complex I. Electron passage through respiratory complex III and cytochrome c oxidase supports a lower phosphorylation yield than entry through NADH at respiratory complex I, consistent with an apparent phosphorylation-to-oxidation ratio around $1.5$. The later involvement of a mitochondrial dicarboxylate carrier is additional evidence because succinate is one of its transported substrates. The ratio alone is not a chemically unique identification: other electron-entry routes and altered coupling can also produce a similar yield. Confirm by comparing defined substrates and sensitivity to a succinate dehydrogenase inhibitor.

Hexokinase consumes exported ATP in the reactionThe adenine nucleotide translocator imports regenerated ADP, permitting ATP synthase to use the proton-motive force. Increased proton return reduces the opposing gradient and allows faster activity of the electron transport chain and oxygen consumption: this is respiratory control. The first hexokinase additions therefore increase demand and respiration. By about the fourth addition, demand no longer limits the measured flux: the respiratory and transport system has reached its available capacity, so more hexokinase cannot increase it. This does not imply that hexokinase ceases functioning, and its zero incremental control at maximal demand does not make it dispensable for maintaining that demand.

At low ATP demand, the proton-motive force is high and much of the remaining oxygen consumption supports proton leak. Increasing leak conductance then creates a substantial extra demand for proton pumping and strongly changes respiratory flux. At high phosphorylation demand, proton return through ATP synthase is large, the proton-motive force is lower, and leak contributes a much smaller part of total proton return. Control of oxygen consumption shifts toward substrate supply and the respiratory machinery, so the leak flux control coefficient becomes small. The plot's value near zero does not mean that the physical leak current vanishes; the separate maximal-rate table likewise gives a small nonzero coefficient, $0.04$.

The mitochondrial dicarboxylate carrier is an inner-membrane transport protein that exchanges suitable dicarboxylates, including succinate and malate, with counter-substrates such as phosphate. In this experiment, it supplies external succinate to the matrix side of respiratory complex II; the inner mitochondrial membrane does not simply allow unrestricted diffusion of the charged substrate. Without adequate uptake, substrate delivery would restrict respiration even if the respiratory enzymes remained intact. Its essentiality is substrate- and preparation-dependent: it is not the obligatory entry route for every possible respiratory fuel.

The reported coefficients add toFor a complete set of independently scaled activities controlling a steady-state flux, the flux summation theorem gives ${\sum }_i{C}_i^J=1$, where $C_i^J=\partial \ln J/\partial \ln E_i$. One way to see this is to scale all reaction capacities by the same factor: steady-state concentrations can remain the same while all fluxes scale by that factor, so the summed logarithmic sensitivities equal one. The listed processes therefore leave a net coefficient of about $0.14$ unaccounted for under the model assumptions. Measurement uncertainty, imperfect inhibitor specificity or incompatible measurements could also contribute to the discrepancy. Control is distributed; neither the sum nor the largest individual coefficient identifies a single universal rate-limiting step.

Important unlisted candidates include ATP synthase, the mitochondrial phosphate carrier, and succinate dehydrogenase itself, which supplies electrons from succinate to ubiquinone. Availability or transfer of ubiquinone and cytochrome c, and associated transport or metabolite constraints, may also need explicit treatment depending on how the pathway is grouped. ATP synthesis, phosphate supply and substrate oxidation have not all been individually represented. Respiratory complex I is absent from the table but is not required for the direct forward route from succinate into the ubiquinone pool; its omission alone does not account for missing control of that route.

Titrate the omitted activities using small perturbations around the same steady-state condition, measuring oxygen consumption and calculating local slopes rather than assigning control from complete inhibition. For example, suitable ATP synthase, succinate dehydrogenase and carrier perturbations could test their contributions, with independent checks of inhibitor selectivity and activity loss. Measure ATP production alongside oxygen consumption, as well as mitochondrial membrane potential, to separate a change in phosphorylation efficiency from a change in electron flux. Check chamber oxygen availability, mitochondrial integrity, substrate abundance and assay reproducibility.

Repeat the control measurements across ADP demands and with defined alternative substrates. Carefully titrated FCCP can test respiratory capacity when proton-gradient backpressure is relieved, but the uncoupled state is a different operating condition and does not itself reveal the coupled-state coefficients. Fit the measured local sensitivities with uncertainty and test whether the expanded set satisfies the flux summation theorem. This distinguishes omitted control from technical inconsistency and shows whether the distribution changes with demand.

Culture each patient and control sample under matched conditions, including equivalent growth-factor withdrawal where appropriate, then prepare paired unstimulated and PDGF-stimulated samples with the same stimulation time. Rapidly lyse on ice with suitable detergents and protease and phosphatase inhibitors. Measure protein concentration and load equal amounts after SDS-PAGE sample preparation. Separate the proteins with molecular-mass markers, transfer them to a suitable membrane, verify transfer and block non-specific binding. Incubate with the Ras-PST25 primary antibody, wash, add a detectable secondary antibody, wash again and record the signal within its linear range. Preserve the original phosphorylation state and compare equal, verifiably transferred inputs. Include a total-protein loading measure, appropriate antibody controls and independent biological replicates; use matched exposures rather than independently rescaling lanes.

The healthy sample establishes the basal and normal stimulated patterns for the same cell type and handling conditions. Sample A contains bands around $96$ and $30\mathrm{kDa}$ in both conditions; PDGF additionally induces bands near $73$, $62$ and $44$–$45\mathrm{kDa}$. These are compatible, respectively, with stimulated Raf, Akt and unresolved MEK/ERK signals, given the supplied masses. Ligand-dependent appearance of those bands is the normal comparison for suspected constitutive signaling. Their identities remain provisional until confirmed with target-specific reagents. The unchanged bands are useful internal observations but cannot be assumed to be validated loading controls merely because their intensities look similar.

The most informative difference is phosphorylation without PDGF, relative to sample A. The following are candidate interpretations of the supplied Western blot, not definitive clinical diagnoses or protein identifications:

A single broad phospho-antibody cannot reliably distinguish the closely sized MEK and ERK bands or determine whether any phosphorylation site is activating. Clinical assessment and genetic and functional confirmation remain necessary.

Use a total-protein stain or validated constitutive loading control, and measure total amounts of the candidate signaling proteins alongside their site-specific phosphorylated forms. This distinguishes increased phosphorylation fraction from increased protein abundance. Include a no-primary or secondary-only control and a phosphatase-treated aliquot to test phospho-dependent detection. Confirm candidate identities using validated target-specific phospho-antibodies, suitable positive controls and, where feasible, depletion or inhibition of the relevant target. Repeat the paired time course with biological replicates and non-saturated imaging. Matching culture state, stimulus dose and sampling time is essential because an ordinary difference in response kinetics could otherwise mimic constitutive activation.

An activating Ras lesion is a parsimonious candidate, because Ras lies upstream of the branch to Raf and the branch to phosphoinositide 3-kinase. Persistent GTP-bound Ras can therefore produce both basal patterns through a single change. A lesion restricted to MEK would not directly explain the simultaneous Akt signal. The blot does not uniquely locate the mutation: a constitutively active upstream receptor tyrosine kinase or altered regulation of Ras could also activate both branches. Test the Ras nucleotide state, sequence relevant genes, and determine which targeted perturbation suppresses both outputs.

mTOR has the supplied molecular mass of about $259\mathrm{kDa}$, so optimize the separation and transfer for a much larger protein. Use a suitably low-percentage or gradient SDS-PAGE gel and high-mass markers, and verify efficient transfer with a protocol appropriate for large proteins, such as optimized wet transfer. Probe with a validated phospho-mTOR reagent and measure total mTOR in parallel. The broad Ras-PST25 reagent cannot be assumed to recognize a useful site on mTOR merely because it recognizes other pathway proteins. Include a positive pathway control and a suitable inhibition or phosphatase control; a missing band on the existing blot could reflect poor separation or transfer rather than absent mTOR phosphorylation.

Analyze the corresponding region from a parallel preparative SDS-PAGE gel, digest the recovered proteins and identify peptides by liquid chromatography-mass spectrometry. The lane and gel region should match the $35\mathrm{kDa}$ signal, with an adjacent-background or control-sample region processed in parallel. Because several proteins can comigrate, validate the candidate against the actual phospho-band, for example by immunoprecipitating the candidate and testing Ras-PST25 reactivity, using a target-specific Western blot, or showing disappearance of the band after specific depletion. Phosphopeptide identification provides additional evidence; mass alone or detecting a common contaminant in the excised region is insufficient.

PDGFR is a receptor tyrosine kinase; its key ligand-induced autophosphorylation is on tyrosine residues. Ras-PST25 recognizes phosphorylated serine residues and threonine residues, so it is not the appropriate reagent to detect the characteristic activated PDGFR phosphotyrosines. Use a phosphotyrosine or validated phospho-PDGFR antibody, ideally alongside total PDGFR. The larger receptor may also require a different gel and transfer range, but changing the size range alone would not correct the epitope mismatch.