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Nuclei with the same resonance frequency in the specified environment. Equivalent chemical shifts do not automatically imply magnetic equivalence when couplings to other nuclei differ.

A chemical shift measures an NMR resonance position relative to a reference. Electronic environments alter local magnetic shielding, so shifts can report a changed molecular environment without uniquely specifying a structure.

Two sites are diastereotopic if replacing one or the other with the same test group gives diastereomers. In an achiral solvent they can have different chemical shifts, unlike a simple pair of symmetry-related sites. A rigid chiral metal chelate can make the two hydrogens of a linker methylene diastereotopic.

NMR of membrane proteins became practical through coordinated improvements in sample preparation, isotope labelling, relaxation control, instrumentation and solid-state methods. The obstacle is not simply that cell membranes are insoluble. Hydrophobic surfaces must remain shielded, while the protein must retain function and form a sufficiently homogeneous sample. In solution, the protein plus its detergent or lipid assembly tumbles slowly; rapid transverse relaxation then broadens resonances until assignments and structural restraints become difficult to obtain.

Recombinant expression and controlled isotope labelling allow selective observation of the protein against its environment. Uniform nitrogen-15 and carbon-13 labelling supports multidimensional assignments. deuteration reduces much proton–proton dipolar relaxation; reintroducing selected protons or labelled methyl groups retains sensitive reporters while limiting the relaxation network. selective methyl labelling is particularly useful for large complexes, but the sparse reporters do not automatically supply enough restraints for a unique complete structure. Exchangeable amide protons may also need back-exchange after growth in deuterated medium, and buried sites can exchange incompletely. Sample production, rather than the spectrometer alone, is therefore a central technical development.

The membrane mimic must balance biological fidelity and spectroscopic tractability. Detergent micelles shield hydrophobic surfaces and can produce relatively small assemblies; inappropriate detergents may alter packing, activity or oligomeric state. bicelles combine bilayer-forming lipids with components that stabilize their edges, and lipid bilayer nanodiscs use a scaffold to enclose a bilayer patch. Smaller, well-defined nanodiscs reduce the additional rotational burden. Franz Hagn and colleagues engineered optimized small nanodiscs and obtained high-resolution NMR information on the membrane protein OmpX, connecting scaffold design with a more tractable lipid-embedded sample.^[ref] No mimic is automatically native: compare ligand binding, transport or catalytic activity with an appropriate cell membrane preparation, and check concentration-dependent aggregation and oligomerization.

transverse relaxation-optimized spectroscopy selects a slowly relaxing component of a coupled spin system. For suitable nuclei and fields, dipolar and chemical shift anisotropy contributions partly cancel for one component while reinforcing others. Combined with deuteration, this preserves narrower lines in systems that would otherwise be inaccessible. The original TROSY experiments established the relaxation-interference principle rather than merely increasing signal by averaging longer.^[ref] Methyl-based variants exploit favourable methyl relaxation properties and selective labelling. Increasing magnetic field improves sensitivity and frequency dispersion, but also changes relaxation contributions; it is not correct to assume that every resonance becomes narrower at every higher field.

High-field magnets, stable radiofrequency electronics and cryogenic detection probes increase accessible sensitivity and resolution. Three-dimensional and higher-dimensional experiments spread overlapping signals and connect resonances through known covalent pathways. Once assignments are secure, nuclear Overhauser effects provide distance information, while residual dipolar couplings and paramagnetic relaxation enhancement can add orientational and longer-range constraints. chemical shift perturbations report changes in local environment on ligand binding, but do not by themselves determine a contact geometry. Each additional measurement restricts the structural model in a different way, which is particularly valuable when only a sparse set of membrane-protein resonances remains visible.

solid-state nuclear magnetic resonance provides a complementary route that does not require rapid overall tumbling. proteins can be studied in bilayers or other immobilized assemblies, using pulse sequences and sample orientation or magic angle spinning to control anisotropic interactions. The magic angle satisfies $3{\cos }^2\theta -1=0$, giving $\theta \simeq 54.74^{\circ }$. Spinning averages appropriate orientation-dependent terms, while decoupling, isotope dilution or deuteration and fast spinning reduce broadening from dense spin networks. Proton detection at very high spinning rates improves sensitivity; Loren B. Andreas and colleagues demonstrated structures of fully protonated proteins using fast-spinning proton-detected experiments, an important general methodological advance rather than a demonstration that every cell membrane sample is thereby solved.^[ref] dynamic nuclear polarization can additionally enhance signal through electron-to-nuclear polarization transfer, although radicals and cryogenic conditions can affect line widths and the conformational states observed.

The outcome is both structural and functional information. Spectra can distinguish conformations, map ligand responses and measure exchange even when no single static structure describes the system. Empty micelles or scaffold-only nanodiscs help identify environmental signals; unlabelled samples, independent assignments and distance-restraint validation guard against incorrect peaks and overfitting. Compare preparations and activity before and after long acquisitions, because aggregation or degradation can selectively remove signals. A missing peak may reflect exchange broadening or overlap, not an absent residue. The enabling combination is a functional and homogeneous cell membrane environment with an isotope and pulse-sequence strategy matched to the size and dynamics of the assembly; solution and solid-state NMR cover complementary regimes.

Biophysical measurements show that enzymes occupy enzyme conformational ensembles; establishing which motions control function requires linking those ensembles to particular kinetic steps. Motions range from picosecond side-chain fluctuations to microsecond–millisecond loop rearrangements and slower domain transitions. They can govern substrate access, productive alignment, chemistry or product release. These roles must be distinguished: the steady-state turnover rate $k_{\mathrm{cat}}$ describes the entire catalytic cycle and need not equal the rate of bond making or breaking.

crystallography identifies the structural framework and can reveal alternative loop positions or ligand-dependent states. Room-temperature measurements and time-resolved experiments can capture changes hidden by a single cryogenic model. Nevertheless, a displacement parameter combines motion, static disorder and modelling uncertainty, and a series of structures is not itself a trajectory or a rate measurement. Spectroscopic measurements in solution are particularly useful because they measure fluctuations and exchange while the enzyme remains in a defined chemical state.

nuclear magnetic resonance resolves dynamics at individual sites. longitudinal nuclear spin relaxation and transverse relaxation and heteronuclear nuclear Overhauser effects constrain fast fluctuations; a model-based protein NMR order parameter $S^2$ describes the amplitude of bond-vector motion after accounting for overall tumbling. NMR relaxation dispersion measures how exchange contributes to relaxation as refocusing conditions change. A two-state analysis can constrain an exchange rate $k_{\mathrm{ex}}=k_{AB}+k_{BA}$, state populations and a chemical shift difference. Measurements at multiple fields and temperatures help separate these parameters, but alternative models and insensitive time windows remain limitations. The inferred minor-state chemical shifts can be compared with separately characterized ligand-bound structures; one should not equate a dispersion curve with a fully resolved molecular movie.

In dihydrofolate reductase, relaxation measurements detected sparsely populated states related to other intermediates of the catalytic cycle, supporting a pre-existing landscape from which ligand binding can select conformations.^[ref] This connects conformational selection with enzymatic progression without excluding subsequent induced fit. In cyclophilin A, comparison of the free enzyme and catalytic conditions showed an intrinsic network of conformational exchange whose rates were associated with turnover.^[ref] Matching rates is informative but not sufficient to prove that a particular motion limits chemistry: the measured exchange may be a coupled consequence, or another step can occur on a similar timescale.

Kinetic methods provide the essential bridge. stopped-flow mixing follows a rapid fluorescence or absorbance response after substrate addition; rapid quench-flow terminates reactions at selected delays and measures chemical products. Binding, conformational change and chemical conversion can thus be fitted to competing kinetic schemes rather than all assigned to a single observed exponential. A pre-steady-state burst can reveal fast chemistry followed by slower release, although its amplitude also depends on the active enzyme fraction and mechanism. temperature jumps perturb an existing equilibrium on shorter timescales and expose relaxation back toward it. isotope substitution can help identify a chemical step, but changes in observed isotope effects must be interpreted within the complete kinetic scheme.

Single-molecule Förster resonance energy transfer adds information lost in population averages. A donor–acceptor pair reports a distance-sensitive efficiency,where the characteristic distance $R_0$ depends on the dyes and their photophysical environment. Trajectories can reveal state dwell times, heterogeneous pathways and ordering of domain transitions relative to a turnover reporter. Labels must preserve activity; donor-only and acceptor controls, corrections for detection efficiencies, and tests for blinking or bleaching are necessary. A FRET change can arise from dye orientation or local quenching, so corroborating distances or structural data are required before assigning a specific domain movement. A mean efficiency also cannot simply be converted into the mean molecular distance because the relation is nonlinear.

hydrogen-deuterium exchange mass spectrometry maps changes in protection and solvent accessibility across an enzyme, usefully locating regions altered by substrate binding or mutation. Exchange also depends on hydrogen bonding, local opening equilibria and chemical conditions, so it is not a direct universal clock for internal motion. molecular dynamics can connect observed sites to candidate atomic pathways and predict further tests. Its force field, sampling and treatment of chemistry limit inference; a conventional classical trajectory does not demonstrate bond-breaking kinetics by itself.

Causal perturbation is stronger than correlation when coupled to these measurements. In the DHFR “dynamic knockout” study, the N23PP/S148A variant retained a closely similar active site ground-state structure but lost much active site millisecond exchange and showed slower hydride transfer. The S148A comparison was especially informative: it also altered the large closed-to-occluded transition yet retained more exchange and much faster chemistry.^[ref] This argues against identifying the gross loop transition itself as the obligatory chemical event. Controls must still examine stability, binding, active site geometry and altered electrostatic or solvation ensembles: a mutation is not an isolated dial for “motion”.

A coherent mechanistic account therefore combines structural states, populations, exchange rates, independently resolved chemical kinetics and perturbations that make successful predictions. In an activated-rate description $k\sim (k_{B}T/h)\exp (-\Delta G^{‡}/RT)$, changing access to productive arrangements can alter the relevant free-energy barrier or pre-equilibrium population. Motions do not provide a new energy source outside thermal fluctuations and chemical free energy. Internal motions can gate access, organize reactive configurations and permit release; which role matters is an experimental question for each enzyme and each step.

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.

The product is 1,4-dimethoxy-2,5-bis(2-methylbutan-2-yl)benzene. Each new substituent is $-\mathrm{C}(\mathrm{C}{\mathrm{H}}_3{)}_2\mathrm{C}{\mathrm{H}}_2\mathrm{C}{\mathrm{H}}_3$, a tert-amyl group, rather than the unrearranged secondary alkyl group of the alcohol. The two substituents are related by the symmetry of the ring.

The proton assignments are:

The integrals add to 30 H. The unusually shielded terminal ethyl methyl signal is compatible with an aryl-containing environment; its NMR quartet–NMR triplet partner and integral are the decisive assignment evidence. The molecular formula and degree of unsaturation, $\mathrm{DBE}=4$, fit a benzene ring with saturated side chains.

The anionic nitrogen donor replaces iodide to giveHere Cp denotes ${\eta }^5$ cyclopentadienyl and $\kappa N$ specifies attachment through the ring nitrogen of the pyrrolyl anion, ${\mathrm{C}}_4{\mathrm{H}}_4{\mathrm{N}}^{-}$. Fe(II), $d^6$, receives six electrons from Cp, four from the two CO ligands and two from an N-bound pyrrolyl ligand: $6+6+4+2=18$.

Two coupled CO stretching modes, approximately symmetric and antisymmetric combinations, account for the two strong carbonyl-region absorptions. The bent two-CO arrangement of this piano-stool complex permits both combinations to be infrared active. They are not evidence for two different chemical products.

Heating expels two CO molecules and changes the binding of the pyrrolyl ligand from ${\eta }^1$ to ${\eta }^5$, giving the sandwich azaferrocene,The four electrons lost with CO are replaced by the four additional electrons donated on changing the hapticity of the pyrrolyl ligand. Thus $B$ is also Fe(II), $d^6$, and 18-electron: $6+6+6=18$. Each complex has one time-averaged Cp proton environment, two pairs of protons from the pyrrolyl ligand, one adjacent to N and one farther from N. Hence three proton environments, with integrals $5:2:2$, fit both structures. For proton nuclear magnetic resonance, three signals means three chemical shift environments; the individual resonances can still show coupling.

The bridged, sterically hindered biphenyl framework is twisted and does not rapidly interconvert the two faces of each methylene group on the NMR timescale. Label one hydrogen on each CH2 $H_A$ and the other $H_B$. Replacing $H_A$ versus $H_B$ by a test substituent gives diastereomeric environments, so these hydrogens are diastereotopic and need not have equal chemical shifts even in an achiral solvent. Molecular symmetry relates the two $H_A$ sites to each other and the two $H_B$ sites to each other.

Each methylene hydrogen undergoes geminal coupling to its partner, giving the two doublets, each accounting for two hydrogens overall. Their separation is $0.21$ ppm; at operating frequency ${\nu }_0$ in MHz it corresponds to $0.21{\nu }_0$ Hz. This separation is not the geminal coupling constant, which must be measured from the line spacing within a doublet. If the shift separation is not large compared with that coupling, an AB pattern with second-order distortion is expected rather than ideal first-order doublets. A racemic pair of twisted conformations still has these signals: enantiomers have the same spectrum in an achiral medium, while the two hydrogen environments within each molecule remain different.

The diagnostic feature is the large vicinal coupling characteristic of an axial–axial relationship in a chair. The td patterns for $H_a$ and $H_c$ contain two large couplings to neighbouring axial hydrogens, so both are axial. The ddd patterns for $H_b$ and $H_d$ also include a large ring vicinal coupling, with the smaller values reflecting the other ring hydrogen and the side chain. The assignments areThe chemical shifts alone do not determine this assignment. In particular, three unequal splittings of a methine are not a geminal-coupling pattern.

For an intact complex in the usual fast-conformational-averaging regime, counting distinct chemical shifts rather than individual split lines gives $C:9$, $D:11$, $E:10$ signals.

In $C$, the uncoordinated pyridinium arm permits the paired positions of a monosubstituted cyclopentadienyl ligand to average: its four ring hydrogens give two sets, $2+2$. The linker $\mathrm{C}{\mathrm{H}}_2$ gives one set, the four inequivalent pyridinium C–H positions give four, its N–H gives one, and the two symmetry-related bound acetonitrile methyl groups give one. Hence $2+1+4+1+1=9$. The exchangeable N–H resonance may be broadened or absent experimentally; nine is the structural count before exchange removes it.

In the rigid cis-dicarbonyl chelates $D$ and $E$, the different pyridyl-N and fourth-leg donors destroy the operation that related the two sides of the substituted ring. Each of the four Cp hydrogens is distinct, and the two linker hydrogens are diastereotopic. Together with four pyridine C–H sites, this gives $4+2+4=10$. The remaining bound acetonitrile adds one methyl signal to $D$, giving 11; $E$ has none, giving 10. The two enantiomers of a racemate have identical spectra in an achiral solvent and do not double these counts. This prediction assumes resolved resonances and no rapid chelate opening that averages inequivalent sites; variable-temperature proton nuclear magnetic resonance would test that assumption.

The vinyl ether has an important resonance contributorDonation from oxygen into the conjugated system increases electron density at the terminal unsaturated carbon, helping explain its unusually shielded ${\delta }_C=87.4$ ppm and its terminal protons near 4.1–4.2 ppm. The directly oxygen-bound unsaturated carbon is strongly deshielded at $151.1$ ppm; its proton at $6.480$ ppm also experiences the oxygen substituent and alkene magnetic anisotropy. These trends are consistent with the whole delocalized electronic structure; a chemical shift is not simply a meter for an atom's partial atomic charge.

For the saturated part, the stronger local deshielding by oxygen places $\mathrm{OC}{\mathrm{H}}_2$ at ${\delta }_C=67.8$, ${\delta }_H\simeq 3.948$, while $\mathrm{C}{\mathrm{H}}_2\mathrm{Cl}$ lies at $41.9$ and $3.708$ ppm. The two terminal vinylic protons have different geometrical relationships to the oxygen substituent and therefore different chemical shifts as well as different vicinal couplings.

The carbon spectra are A → 5, B → 8, C → 6, and D → 7. Number each acyclic acid or ester chain from its carbonyl carbon. The assignments below account for every carbon-13 NMR environment.

For A, both ketone carbonyl carbons have $\delta =208.3$ ppm and all four methylene carbons have $\delta =36.5$ ppm. Molecular symmetry and rapid conformational averaging explain why six carbons give only two resonances.

For B, written $\mathrm{HOOC}-\mathrm{CH}=\mathrm{CH}-\mathrm{CH}=\mathrm{CH}-\mathrm{C}{\mathrm{H}}_3$, assign $\delta ({\mathrm{C}}_1)=172.8$, $\delta ({\mathrm{C}}_2)=118.1$, $\delta ({\mathrm{C}}_3)=147.4$, $\delta ({\mathrm{C}}_4)=129.7$, $\delta ({\mathrm{C}}_5)=140.8$, and $\delta ({\mathrm{C}}_6)=18.1$ ppm. The alternation among vinylic chemical shifts reflects polarization of the conjugated chain, especially the deshielded carbon beta to the carboxyl group.

For C, written $\mathrm{HOOC}-\mathrm{C}\equiv \mathrm{C}-\mathrm{C}{\mathrm{H}}_2-\mathrm{C}{\mathrm{H}}_2-\mathrm{C}{\mathrm{H}}_3$, assign the carbonyl at $158.5$, the adjacent sp carbon at $72.9$, the other sp carbon at $92.6$, and the terminal methyl at $13.4$ ppm. The two methylene resonances are $20.7$ and $21.0$ ppm. A plausible assignment is ${\mathrm{C}}_4=20.7$ and ${\mathrm{C}}_5=21.0$ ppm, but their ordering is tentative from the supplied shift list alone; a correlation experiment would resolve it.

For D, written $\mathrm{C}{\mathrm{H}}_3\mathrm{CH}=\mathrm{CHC}(=\mathrm{O})\mathrm{OCH}=\mathrm{C}{\mathrm{H}}_2$, the carbonyl is at $163.2$ ppm. The crotonyl beta-CH, alpha-CH and methyl carbons are at $148.0$, $121.7$ and $18.2$ ppm, respectively. The O-bound vinyl CH is at $141.3$ ppm and its terminal $\mathrm{C}{\mathrm{H}}_2$ is at $97.4$ ppm. The two sides of the ester oxygen therefore produce quite different vinylic environments.

Use the proton labels in the original drawing below. Let $a$ denote the three methyl hydrogens, $b$ the crotonyl beta-CH, $c$ the crotonyl alpha-CH, $d$ the O-bound vinyl CH, and $e,f$ the terminal vinyl hydrogens trans and cis to $d$, respectively.The integrals sum to eight hydrogens. Multiplet centres give the chemical shifts; a peak label at one edge of a multiplet is not its centre. The terminal vinyl hydrogens are chemically nonequivalent because one is cis and the other trans to the O-bound vinyl hydrogen. Each crotonyl CH couples to the other and to the methyl group; the resolved long-range interaction makes the methyl a doublet of doublets rather than a simple doublet.

The assignments are ${\mathrm{H}}_a$: $6.787$ ppm, $1$H, doublet of quartets; ${\mathrm{H}}_b$: $6.496$ ppm, $1$H, doublet of quartets; ${\mathrm{H}}_c$: $4.281$ ppm, $2$H, quartet; ${\mathrm{H}}_d$: $1.331$ ppm, $3$H, triplet. The chemical shift of a multiplet is its centre, rather than its most downfield line.

Each vinylic proton has scalar spin-spin coupling to the other vinylic proton and to three equivalent spin-$\frac{1}{2}$ fluorine nuclei. The first interaction produces a doublet, and the second a $1:3:3:1$ quartet: eight lines overall, partly interleaved for ${\mathrm{H}}_a$. The two chemical shifts differ because the vinylic sites have different substituents.

The ethoxy $\mathrm{OC}{\mathrm{H}}_2$ protons are deshielded by oxygen and split into a $1:3:3:1$ quartet by the three methyl protons. The methyl protons are split into a $1:2:1$ triplet by the two equivalent methylene protons. The relative NMR signal integrals are $1:1:2:3$.

The six carbon environments are assigned as follows, with chemical shifts at the centres of the fluorine-split multiplets:The supplied carbon-13 spectrum is interpreted as proton-decoupled NMR, as indicated by its unsplit ethoxy signals. Proton decoupling removes C–H splitting but leaves C–F scalar spin-spin coupling. Three equivalent spin-$\frac{1}{2}$ fluorine nuclei give each resolved quartet its $1:3:3:1$ intensity pattern. The carbonyl's much smaller four-bond C–F interaction is unresolved in the plotted spectrum; “singlet” describes what is observed, rather than proving an exactly zero interaction.

The wide quartet is centred at $(127.48+116.74)/2=122.11$ ppm. The other two centres are $(132.04+130.63)/2=131.335$ and $(129.09+128.85)/2=128.97$ ppm. The signal near $77$ ppm is the deuterated chloroform solvent, not another carbon of the analyte. The structure labels skip ${\mathrm{C}}_i$: the ethoxy methylene is ${\mathrm{C}}_j$.

One fluorine-19 NMR resonance, integrating to three equivalent fluorine nuclei, appears as a doublet of doublets with $J=6.6$ and $1.8\mathrm{Hz}$. This predicts four principal lines of equal intensity, from coupling to ${\mathrm{H}}_a$ and ${\mathrm{H}}_b$, respectively. Relative to the unknown resonance centre, their offsets are $\pm 4.2$ and $\pm 2.4\mathrm{Hz}$.

Equivalent fluorine nuclei do not split one another. This prediction assumes ordinary proton-coupled acquisition; proton decoupling would collapse these four principal lines to a singlet. The proton and carbon spectra do not determine the numerical fluorine chemical shift. Weak natural-abundance carbon-13 satellites could accompany the principal pattern.

Define the retard ratio using the kinetic energy at the analyser entrance before the deliberate retardation: $R=K_a/E_p$. Initially $R=111.4/100.0=1.114$. With the original excitation, a carbon photoelectron arrives with $K_{a,C}=515.0-284.8-4.4=225.8\mathrm{eV}$. ThusThe new deliberate retardation is $225.8-202.69=23.11\mathrm{eV}$. For fixed slits, angular acceptance and hemisphere radii, the geometrical fractional energy width is approximately constant, so the absolute analyser width grows in proportion to pass energy: about 2.03 times poorer absolute energy resolution. Counts may improve, but closely separated chemical shifts become harder to resolve. An alternative definition $(K_a-E_p)/E_p$ gives 0.114 rather than 1.114 for the same physical setting and leads to the same final pass energy when used consistently.

Use two-dimensional NOESY, or ROESY where molecular tumbling makes NOESY less favorable, to look for through-space correlations between proton sites on the two ends that are brought together in the closed conformation. Assign resonances first and compare predicted ring contacts with an open reference or model.

The nuclear Overhauser effect involves NMR cross-relaxation through magnetic dipole–dipole coupling. Its strong approximate $r^{-6}$ dependence makes it sensitive to short interproton distances. A NOESY cross-peak therefore reports spatial proximity, including between sites far apart along the covalent skeleton; it is not a NMR scalar coupling transmitted along that skeleton.

Run a dilution series: end-to-end contacts that persist at concentrations where intermolecular association is negligible support intramolecular ring closure. Hydrogen-bonded NH chemical shifts and diffusion measurements can support the assignment, but neither alone proves ring topology. A flexible open chain can transiently bring ends together, and intermolecular contacts can also give NOEs. Several geometry-consistent cross-peaks, concentration behavior and controlled mixing times make the case much stronger than one peak.

nuclear magnetic resonance of the spin-one-half isotope ${}^{29}\mathrm{Si}$. Equivalent silicon sites give the same isotropic chemical shift, while condensation and silanol substitution can distinguish environments. Line heights alone are not quantitative populations.