<figure><img src="image_23.png" /><figcaption>**Fig. 19.** Operating principle of the current para-hydrogen induced hyperpolarization (PHIP). A) (1) a alkyne-containing moiety is attached to the ¹³C-labeled metabolite of interest, (2) hyperpolarized para-hydrogen reacts with the alkyne function in an organic solvent, (3) the hyperpolarization is transferred to the ¹³C-nuclei by magnetic field-cycling and (4) the final metabolite of interest is extracted in water; B) Anaerobic glycolysis, a hallmark of cancer, is monitored by [1-¹³C]pyruvate reduction, which leads to [1-¹³C]lactate; C) Stacked one-dimensional ¹³C-NMR spectra of cancer cells in suspension upon supplementation with hyperpolarized [1-¹³C]pyruvate. Adapted from Cavallari et al. 2019 [1138].</figcaption></figure>

them appealing [1135]. Non-permeating compounds may also be studied using d-DNP combined with an online electroporator placed on the route between the polarizer and the analyzing magnet [1136]. To be more useful, this complicated arrangement would probably require improvements to cell survival upon electroporation, which reached only 50% according to the authors.

Analogously to d-DNP, para-hydrogen induced hyperpolarization (PHIP) is another method recently applied to monitor cellular metabolism in real time. Dihydrogen can be hyperpolarized at cryogenic temperatures (~30-70 K) at low cost compared to DNP. Hyperpolarization transfer to ¹³C-labeled metabolites requires execution of a suite of chemical reactions on precursors that can be hydrogenated, followed by magnetic field cycling and then extraction into water [1137] (Fig. 19). PHIP permitted characterization of hyperpolarized ¹³C-pyruvate to lactate conversion [1138,1139] and hyperpolarized ¹³C-fumarate to malate conversion [1140] in suspensions of cancer cells.

Although one might naively predict polarization enhancements in the order of ~10,000-50,000 fold, in practice d-DNP and PHIP have only been used to study metabolites in the millimolar range, far from the originally expected (sub)micromolar range. We consider this point further in Section 3. d-DNP is however a flexible method, which may theoretically be applied to many metabolites, even though they are not all equally amenable due to their different intrinsic relaxation times. However, d-DNP NMR is far from being a standard method for studying cellular metabolism. Many working on Metabolic Flux Analysis use chromatography-coupled Mass Spectrometry approaches, together with NMR spectroscopy of extracted material from quenched biological samples [39,762,798-800,1141]. d-DNP NMR is surely promising for metabolomics studies of biological extracts and fluids though, as shown by the recent ¹³C-NMR study at natural ¹³C-abundance of tomato extracts in less than 30 min per spectrum [1142]. Concerning our focus, i.e. real-time, non-destructive measurements on living cells, d-DNP NMR for metabolic studies offers interesting capabilities: we can imagine for example repeated injections of hyperpolarized material would permit surveying of the metabolic status of single samples over periods of days, provided that bioreactors are used to ensure cell viability. Such “high-end” equipment and expertise are of course not yet commonly available, but are becoming more and more accessible via established networks of NMR facilities.

## 2.4. Cellular structural biology

High resolution NMR is one of the few methods that allows structure determination at atomic-resolution of biological macromolecules. Compared to X-ray crystallography and cryo-electron microscopy, 3D structures determined by NMR spectroscopy only account for ~10% of all deposited structures in the Protein Data Bank [1143]. Nonetheless, NMR remains a very useful technique, particularly for studying smaller proteins that do not crystallize, as well as relatively weak interactions in solution; it has important capabilities in characterizing the conformational dynamics of many macromolecules (for instance, see [1144-1147]). NMR is thus extremely helpful in understanding the function of folded proteins, but it is indispensable for studying disordered proteins [1148-1158]. NMR is also useful to study cellular processes that occur at slower timescales than those of the internal motions, such as protein maturation events, cofactor binding, and post-translational modifications [1159-1166], all of which can contribute to regulation of protein function. These unique abilities for characterizing proteins at the atomic scale can be applied in complex native environments, notably by using ¹³C/¹⁵N isotope-filters.

Classical structure determination by solution NMR is applicable to relatively small, freely tumbling proteins [1143], while ssNMR is applicable to insoluble samples such as proteins embedded in intact membranes, fibrils, sediments or microcrystalline samples [1167]. Both approaches have been applied to determine the structure of proteins within intact cells or in native membranes, as discussed below. More recently, DNP-assisted ssNMR greatly improved the lower limits of sensitivity [1073,1074,1168-1171]. Currently, this technique necessitates low temperatures to be effective (~100 K or below) and is thus applied to frozen samples. It can provide structural information at atomic-resolution that complements cryo-electron microscopy data [1172,1173].

In-cell NMR has the ability to probe the deep ocean of macromolecular interactions occurring within the cell. These include the classical protein-protein interaction networks, the building blocks of all textbook cellular pathways, but also the complex macromolecular assemblies between proteins and nucleic acids phenomena, and finally the elusive “protein quinary structure”, a postulated self-organization of the cellular milieu itself occurring
through weak, diffusive interactions between all cellular
components.

Importantly, *in vitro* NMR studies of proteins commonly use high concentrations, approx. 0.1 to 1 mM. So far, in-cell NMR spectra of proteins have been recorded down to ~10 µM at the lowest, which gives access to less accurate structural information than *in vitro* NMR [1174]. However, higher concentrations would not necessarily be advisable: to put things into context, most endogenous proteins show cellular concentrations in the micromolar range, although the distribution is broad, from the picomolar to the millimolar range [1175-1177] (see Section 3.5.).

2.4.1. Solving structures of folded, soluble proteins in cells

¹³C and ¹⁵N isotope labeling, as well as ¹⁹F incorporation have been used since the 1970's to permit NMR isotope filtering to suppress the cellular background signals and study cellular walls or metabolites in living cells (see Sections 2.2. and 2.3.) [583,584,668,715]. This concept was later applied to proteins in cells. To the best of our knowledge, the first example reported ¹³C-histidine labeling and ¹³C-observation of hemoglobin in erythrocytes from mice fed with ¹³C-histidine [1178]. Isotope labeling of proteins was not then the standard approach that it has become since, and it took about twenty years before NMR measurements in complex media re-emerged as an interesting possibility. We can cite the ¹H-¹⁵N NMR spectra of GB1 in crude extracts in the 1990's, and, closer to our topic, the incorporation of 5-fluorotryptophan in yeast by Brindle and colleagues, which gave access to background-free ¹⁹F-NMR of enzymes in living cells [1179]. The approach became more popular once, in the early 2000's, Dötsch and colleagues used ¹⁵N-labeling of proteins overexpressed and then measured in *E. coli*, demonstrating the feasibility of the technique and the variety of the accessible information [1,1180-1182]. Cells could be grown initially in natural abundance media, and then transferred into ¹⁵N-enriched medium before inducing recombinant protein expression. Isotope labeling provided an effective filter to distinguish the protein signals from the cellular background, thus in principle allowing structural studies in complex biomolecular mixtures. This advantage was used to determine the structure of a ¹³C,¹⁵N-labeled protein from NOE-based restraints measured in a crude cell-extract stabilized with protease inhibitors [1183]. Intact cells, however, present several additional challenges. Above all, the 3D NMR experiments that are needed for resonance assignment and NOE-based structure calculation require long acquisition times, which are incompatible with the short lifetime of cells densely packed in the NMR tube. This and other limitations were overcome by Ito, Güntert and their coworkers, who demonstrated in 2009 that NOE-based *de novo* 3D protein structures could be obtained from NMR spectra recorded

from intact bacteria [1184]. Despite the huge methodological effort
[1185,1186], the in-cell structure of the small metalloprotein
TTHA1718 overexpressed in *E. coli* was overall superimposable to
that determined *in vitro*, except for small differences in the
heavy-metal binding loop attributed to the interaction with metal
ions in the bacterial cytosol. These authors further improved their
protocols to allow structural determination from lower levels of
expressed proteins in bacteria [1187] and in insect cells [1188].
The resulting structures of 5 proteins were mostly superimposable
on those determined *in vitro*, and the minor differences observed
were again attributed either to the smaller number of distance
restraints available from in-cell spectra, or to the interaction of
the protein with metal ions or other cofactors in the cytosol
(Fig. 20). This is consistent with the minute chemical shift differ-
ences observed by others in in-cell NMR spectra of small folded
proteins relative to *in vitro* values [1189-1192].

While in-cell NOE-based structure calculation remained a
tedious task, an alternative approach to obtain protein structures
from simple 2D ¹H-¹⁵N spectra with the aid of paramagnetic
restraints was proposed independently by three groups in 2016
[1193-1195]. In these studies, the small folded GB1 domain or
ubiquitin were tagged with lanthanide chelating tags and injected
into *Xenopus laevis* oocytes or HeLa cells. The paramagnetic tags
gave access to orientation and distance restraints (namely
pseudo-contact shifts, PCS, and residual dipolar couplings, RDCs)
for structural calculations. Similarly to the NOE-derived structures,
the structural model of GB1 obtained in both studies was almost
identical to the structure determined *in vitro*, and the minor differ-
ences were again attributed to weak interactions with the cellular
milieu.

PCSs are also complementary to ¹⁹F-labeling, which permits protein structural characterization in cells using 1D ¹⁹F-NMR, as shown by Li and his coworkers: i) they characterized *in vitro* the ¹⁹F-NMR signature of open and closed conformations of a Tb³⁺-chelated form of calmodulin containing ¹⁹F-tyrosine; ii) they microinjected this labelled protein into *X. laevis* oocytes with or without a binding peptide, and could monitor calmodulin conformations using ¹⁹F-NMR [1196].

The structures obtained to date in cellular settings suggest that the tertiary structure of soluble folded proteins is generally unchanged. This observation may be biased by the selected set of studied proteins, i.e. small, stable proteins yielding good quality spectra such as GB1, ubiquitin and, TTHA1718. Therefore, aside of the methodological achievements, it remains to be seen whether it will be worthwhile in future to determine protein structures in cells, if the same structures can be obtained *in vitro* with much less effort. Notably, an important conclusion of these studies is that minor differences in protein conformation or dynamics are often

<figure><img src="image_24.png" /><figcaption>**Fig. 20.** In-cell NMR structure of the small model protein GB1 in insect cells. Left: backbone amide chemical shift differences between purified and in-cell GB1 at 600 MHz; Right: Overlay of lowest energy structures of purified (red) and in-cell (blue) GB1. Adapted from [1188].</figcaption></figure>
correlated with chemical shift changes in the backbone resonances, meaning that chemical shift perturbation can help in the initial assessment of intracellular conformational changes at the single residue level, without needing a complete structural characterization.

### 2.4.2. Structural studies of membrane and cytoplasmic proteins using solid-state NMR

All the above approaches based on solution NMR are restricted to macromolecules that i) tumble freely in the cytosol, ii) are not significantly involved in diffuse interactions, iii) are not associated in large complexes, any of which cases would cause severe relaxation broadening of the signals. In parallel, solid-state NMR approaches have been developed that overcome these intrinsic limitations of solution NMR and allow the structural characterization of integral membrane proteins in intact cells or in native membranes, intracellular fibrils and – in frozen samples – soluble proteins inaccessible by solution NMR. DNP-assisted cellular ssNMR approaches have been further developed to overcome the low sensitivity associated with the use of small, fast spinning rotors.

NMR structural characterizations of membrane proteins in native cellular membranes were reported concomitantly by a handful of groups in the early 2010's [1197-1200]. Tian and colleagues overexpressed the transmembrane domain (TM) of the human LR11 protein in *E. coli* using uniform ¹³C,¹⁵N. Then, they lysed the bacteria and isolated the bacterial membranes by ultra-centrifugation and recorded ¹³C-¹³C correlation spectra, which revealed the characteristic chemical shifts of an α-helical conformation similar to that observed in DPC micelles [1198]. Similarly, Miao et al. applied ssNMR to investigate the structure of the M2 proton channel tetramer of influenza A embedded in *E. coli* membranes [1200]. The data obtained by NMR *in situ* was compared to that from the purified channel in reconstituted synthetic bilayer and revealed that, aside from small differences, the conformation of the histidine tetrad in the channel was essentially unperturbed. The same membrane extraction methodology was pursued by the groups of Oschkinat and Reif, but they took advantage of the higher sensitivity offered by the commercial DNP-ssNMR system recently developed at that time, which allowed spectra to be recorded using lower amounts of protein. Notably, they obtained structural information on a small bacterial protein in native membranes using GSTV-amino acid selective ¹³C/¹⁵N-labeling. ¹⁵N-edited 3D experiments allowed suppression of the lipid resonances. Here again, the authors could observe resonances at chemical shifts corresponding to α-helical conformations in native membranes.

The Baldus group further extended the above approach to whole bacterial cells [1201,1202]. Renault et al. applied DNP-ssNMR to analyze ¹³C,¹⁵N-labeled whole *E. coli* cells overexpressing the integral outer membrane protein PagL. By comparing the whole cell spectra with those obtained at higher resolution on isolated cell envelopes, they could identify resonances of PagL and other membrane-associated macromolecules (major lipoprotein, lipopolysaccharides, peptidoglycans and phospholipids). Using ssNMR at low temperatures, the same group characterized the major components of the bacterial membrane at even higher resolution in ¹³C,¹⁵N-labeled whole cells and cell envelopes. ¹³C resonance assignments obtained from purified samples were transferred to the native membranes with the aid of further 2D/3D experiments, allowing conformational analysis from ¹³C chemical shifts. While lipopolysaccharides and the peptidoglycan layer were rather mobile, PagL adopted the overall rigid fold seen in the crystal structure. However, several loops and the transmembrane segments 1-3 appeared to be sensitive to the effects of the native membrane environment.

Following these seminal works, (DNP-)ssNMR was slowly but increasingly adopted and can now be considered as the NMR technique of choice to investigate structural and dynamic features of proteins embedded in native membranes. When studying proteins in heterogeneous samples such as native membranes, the cryogenic temperatures required for DNP-ssNMR decrease the spectral resolution, due to the fact that flexible proteins are frozen in many different conformations. Despite this limitation, tailored amino acid selective labeling strategies allow assignment of residue pairs and secondary structure analysis, as shown by Kaplan et al. in an exemplary application of DNP-ssNMR to investigate the megadalton-sized bacterial type IV secretion system core complex [1173]. Yamamoto et al. evaluated the suitability of a similar approach, using low-temperature ssNMR and DNP to investigate cytochrome b5, a single-pass membrane protein harboring a large soluble domain, both in intact bacteria and in artificial membranes [1203]. Conversely, Shahid et al. showed that ssNMR at higher, closer-to-physiological temperature still enabled structural and dynamic information of a membrane-anchored domain of YadA, previously overexpressed in the isolated outer membrane of *E. coli* [1204]. In this case too, the transmembrane domain of YadA was found essentially in the same conformation as for the isolated microcrystalline protein, even though the more flexible regions exhibited a more dynamic behavior.

The supramolecular assembly of the KcsA channel was further investigated by Baldus, Weingarth and their colleagues in membranes reconstituted with *E. coli* lipids [1205]. Vissher et al. combined ¹³C-detected DNP-ssNMR with MD simulations to settle a debate about the opening mechanism of the channel, demonstrating that, in the membrane, channel gating and clustering are correlated phenomena, supporting the notion that the clustered channels can open in a concerted manner *in vivo*. An atomistic explanation of this phenomenon was given based on the observation that the gate opening/closing perturbed the structure of the N-terminal TM helix that contributes to the channel-channel interface and modulates clustering. The mechanism of KcsA gating in a reconstituted lipid bilayer was further elucidated by Jekhmane et al. through ¹H-ssNMR and MD simulations [1206]. Consistent with the above, mutations in a residue critical for gate selectivity were shown to cause marked changes in conformational dynamics. With the same approach, Pinto et al. investigated the bacterial β-barrel assembly machinery complex (BAM), a large multiprotein membrane complex required for the insertion of proteins into the outer membrane, in native *E. coli* membranes by ¹³C- and ¹H-detected ssNMR [1207]. Fractional deuteration and amino acid-selective ¹³C labeling strategies were employed to reduce the spectral complexity and provide structural information on the complex assembly in natural membranes. Overall, the results suggested that the various subdomains of BAM adopt the same folds observed in their detergent-solubilized complex. However, the large size and spectral complexity of the system prevented a detailed analysis at the spectral resolution reached by ssNMR. The complex assembly could be characterized in higher detail in synthetic lipid bilayers by ssNMR and DNP-ssNMR [1208].

The Weingarth group showed that (DNP-)ssNMR can provide precious atomistic insights into the mechanism of action of antibiotics bound to native bacterial membranes. For this purpose, Medeiros-Silva at al. combined high resolution ¹H-detected ssNMR in non-freezing conditions [1209] and DNP-enhanced ¹³C-detected ssNMR in frozen samples to determine the structure of the pore-forming antibiotic nisin bound to its target, the peptidoglycan precursor lipid II, in native membranes from the nisin-sensitive bacterium *Micrococcus flavus* [625]. The lipid II-nisin structure in the membranes was strikingly different from that determined in organic solvents, as the nisin-lipid II pore can form only in the membrane environment. In the pore structure, flexible hinge
regions were identified that allow nisin to adapt to the target membrane, suggesting that they could be mutated to improve nisin activity towards membranes from other pathogens. The same group used (DNP-)ssNMR to further study the mode of action of teixobactins, a recently discovered class of antibiotics highly active against several pathogenic bacteria, in native *Micrococcus flavus* membranes [626]. In that work, Shukla et al. showed that teixobactins form clusters on the membrane surface and solved the structure of the teixobactin-Lipid II complex. By comparing the complex in native membranes with that formed in different synthetic micelles, it was found that the binding affinity is strongly attenuated if membranes are anionic (as is the case for *M. flavus*), suggesting that changes in bacterial membrane composition modulate the efficacy of teixobactins.

Overall, the work by Baldus, Weingarth and other groups showed that ssNMR and DNP-ssNMR approaches can provide unique contributions to the characterization of membrane proteins embedded in native membranes. Furthermore, the insights into protein dynamics obtained by solid-state NMR perfectly complement the high-resolution structural information that nowadays can be obtained by cryo-electron tomography (cryo-ET). In explorative work by Baker et al., Baldus and Grünewald combined ssNMR and cryo-ET in a hybrid approach to investigate the structure and dynamics of *E. coli* YidC, a protein of the inner membrane responsible for inserting other inner membrane proteins into native cell envelopes [1210]. The same sample preparations were analyzed by ssNMR and cryo-ET, thereby allowing the integration of the results in the same structural model without introducing artifacts arising from sample differences. The correct morphology of cell envelope samples was assessed by cryo-EM, while ssNMR revealed conformational differences with respect to YidC reconstituted in micelles. Additionally, a structural model of the ribosome nascent chain in complex with YidC was obtained by cryo-ET, albeit at low resolution due to the low number of molecules. While this method has not been applied further, the recent progress of ssNMR and cryo-ET, in terms of spectral and spatial resolution, suggest that such hybrid strategies will be crucial in the future to provide a complete atomistic description of highly dynamic proteins, which are still challenging for cryo-EM/ET (see Section 4.1.).

The above studies show that samples of bacterial cells or isolated membranes are amenable to ssNMR studies, thanks in part to the adaptability of bacteria to autotomy and to the development of protocols for single-protein overexpression and selective isotope labeling schemes. Clearly, from the sample preparation standpoint, eukaryotic cells are more demanding, as the variety of expression and labeling protocols that is available for bacteria cannot be used. Furthermore, the increased size and complexity of eukaryotic cells imposes additional limitations in terms of increased spectral crowding and decreased concentration of the molecule of interest. Despite these difficulties, Kaplan et al. showed that ssNMR could be successfully applied to investigate natively abundant membrane proteins in plasma membrane vesicles extracted from human cells [1211]. For this purpose, they focused on the human epidermal growth factor receptor (EGFR), which was found to be highly abundant on the plasma membrane of A431 cells. In the purified vesicles, EGFR retained the correct orientation with respect to the asymmetric composition of the native membrane. The dynamics of the extracellular domain of EGFR isolated from isotope-labeled cells were characterized by room-temperature ssNMR and low-temperature DNP-ssNMR, revealing an unexpected conformational flexibility, which was greatly decreased upon binding to the epidermal growth factor. According to the authors, this would imply that a reduction in conformational entropy drives the EGF-dependent activation of EGFR by contributing to the free energy of EGFR dimerization.

Unlike membrane proteins, for which the 'native environment' can be preserved to a good extent by isolating the cellular membranes in which they reside, cytosolic and other intracellular proteins must be analyzed in intact cells to avoid dilution or alteration of the intracellular environment. Compared to isolated membranes, intact cells pose additional problems such as mechanical stability and dilution of the molecule of interest. Signal enhancement by DNP has been suggested to be necessary for obtaining sufficient intensity since the first in-cell ssNMR study by the Dötsch group in 2012 [1212]. In terms of applications, Narasimhan et al. demonstrated recently that intracellular soluble proteins, which would normally be studied by solution NMR, can be characterized at the structural level by DNP-ssNMR in intact human cells [1213]. In that work, an approach relying on cell electroporation for delivering isotope-labeled proteins (previously purified from recombinant bacterial expression) [1214] was chosen. This was applied to wild type ubiquitin, which is notoriously undetectable by conventional in-cell solution NMR due to extensive interactions with other cellular components (see Section 3.2.3.). Intracellular ¹³C,¹⁵N-labeled ubiquitin was selectively observed and shown to be correctly folded. 2D and 3D DNP-ssNMR experiments provided sufficient resolution to assign unambiguously several signals to protein residues, thereby suggesting that this method could pave the way for further structural studies in intact cells.

We expect that further methodological developments in the short term will focus on the optimization of cell sample preparation, and on the application of DNP to biological samples of increasing complexity, approaching that of real tissues. Concerning the latter, the Baldus group reported very recently the first pilot DNP-ssNMR study of isotopically labeled spheroids, which are 3D cell cultures increasingly employed as high-fidelity tumor model systems [1215]. In this latter work, Damman et al. established optimal conditions for *in vitro* spheroid growth and handling for DNP sample preparation. ¹H-¹³C INEPT spectra recorded on uniformly labeled cells showed different metabolic composition compared to that of the same cells cultured on monolayers. In addition, they showed that protein DNP-ssNMR was possible by observing ¹³C-¹³C correlations from an isotope-labeled EGFR-bound nano-body, thus opening the way towards structural biology studies in highly organized, tissue-like cell samples.

### 2.4.3. Disordered proteins: intracellular conformations, dynamics and interactions

The ability of NMR to probe protein conformational dynamics is of strong interest when studying the intracellular behavior of "intrinsically disordered" proteins or regions of proteins (IDPs, IDRs). By definition, these do not adopt a stable 3D structure by themselves, but can adopt ordered conformations upon binding [1149,1216,1217]. It may seem strange now, but the existence of disordered proteins was controversial twenty years ago, with some assuming that the absence of a stable fold was a simple matter of proteins waiting for a partner [1218]. However, the functionality of disordered regions in modulating binding kinetics and affinities of neighboring regions, exerting chaperoning activities, connecting multidomain proteins, transmitting cell signaling through post-translational modifications or triggering phase separation has now become clearer [1153,1219-1226]. NMR has contributed to the exploration of the physical properties of these objects, and, importantly, has allowed atomic-scale experimental observation of the existence of disordered states in cells.

How disordered proteins were affected by macromolecular crowding was one of the first questions that were tackled using in-cell NMR in the early 2000's [1227]. The highly complex intracellular environment can indeed affect the propensity of IDPs to adopt secondary or tertiary structures, to aggregate, or to modulate
their interaction with cellular partners or metal ions [6, 1150, 1228]. For studying these peculiar proteins, solution NMR at physiological temperatures is the technique of choice: freezing the cells would immobilize every molecule in a different conformation, preventing analysis of the conformational ensemble by ssNMR or single particle cryo-EM in most cases. Furthermore, the dynamic nature of IDPs/IDRs is advantageous for in-cell solution NMR studies: unlike stably folded proteins, where global tumbling dominates the correlation time of every residue, local dynamics at the level of individual residues in an unfolded peptide are mostly independent from each other. Therefore, while any interaction on any surface patch of a folded protein will affect global tumbling and thus cause line-broadening of all residues, only the residues directly involved in a given interaction will be affected in IDPs. This gives access to residue-level mapping of interactions even in the case of signal broadening beyond detection.

This effect was observed by Pielak and colleagues on FlgM, a polypeptide from *Salmonella enterica Typhimurium* that regulates flagellar synthesis [1227]. The bacterial environment caused signal broadening in the FlgM C-terminal half, which was attributed to a shift towards an alpha-helical conformation, that was previously observed *in vitro* [1229]. This conclusion was probably insecure at that time, because intermolecular interactions in cells would have had the same impact on FlgM NMR spectra, as was recognized later by Pielak [1230]. The opposite approach was exploited by the Shekhtman group to investigate the behavior of a prokaryotic ubiquitin-like protein (Pup) in the cellular environment. In *Mycobacterium tuberculosis*, Pup links covalently to substrate proteins and targets them to the proteasome by binding to mycobacterium proteasomal ATPase (Mpa). Maldonado et al. observed that, in the absence of Mpa, the protein remained fully unfolded in bacteria and was free from any other interactions [1231]. Using the so-called STINT-NMR approach employing mono- or dual- overexpression in *E. coli* of a pair of proteins of interest [1232,1233], the authors further investigated the interaction between Pup and Mpa in cells, and observed line broadening within the α-helical region (residues 21–51) and at the C-terminus (52–64), together with chemical shift perturbation in the C-terminal half of the α-helix, providing a mapping on Pup of the Pup:Mpa interaction [1231]. More extensive interaction was observed between Pup and the intact 1.2 MDa proteasome, leading to line broadening of most Pup signals. In later work, Majumder et al. re-analyzed the same interaction using singular value decomposition (SVD, see Section 3), in order to filter out spectral changes that were not directly caused by the Pup:proteasome interaction [1234]. They derived a more accurate set of interacting residues localized in the conserved region of the Pup α-helix, in agreement with X-ray data.

Among IDPs, α-Synuclein (α-Syn) has been by far the most studied. Its hallmark aggregates in Parkinson's patients (α-Syn is the primary component of Lewy Bodies) makes α-Syn a target of high biological and clinical relevance. In addition, α-Syn exhibits excellent stability at high concentrations and low secondary structure propensities *in vitro*. Hence, α-Syn is often used as a gold standard for developing NMR methods devoted to IDPs [396,397,1156,1157,1235-1240]. α-Syn is also found to bind lipid surfaces in cells, where the N-terminal residues 1-100 adopt an α-helical structure in their bound conformation [1241-1243]. In 2011, Bartels et al. reported that α-Syn purified from human cells adopted a tetrameric, α-helical conformation, but this interpretation was soon rejected by most of the community [1244,1245] prompting further investigation of the conformation of α-Syn in the cellular environment. The original hypothesis was said to be corroborated by a surprising interpretation of some α-Syn NMR spectra that supposedly indicated an α-helical conformation, maintained due to a milder purification strategy from the *E. coli*

cells used for recombinant expression [1246]. The Selenko group provided more definitive proof that the unfolded monomer is the major conformation of α-Syn both in bacteria and human cells, independently from the N-terminal acetylation. First, Binolfi et al. showed an exact superimposition of the NMR spectra from non-purified α-Syn in bacteria and all those recorded *in vitro* [1247]. This observation has been corroborated independently by multiple groups [1236,1248,1249]. Later, Theillet et al. performed a deeper investigation of the backbone dynamics of α-Syn in human cells by NMR and EPR [1214]. The authors developed a protein electrophoresis approach to deliver ¹⁵N-labeled recombinant α-Syn in a variety of human cell lines, both of non-neuronal and neuronal origin. Once present in the cytosol of these cells, α-Syn was quantitatively N-terminally acetylated in a post-translational fashion, as revealed by the chemical shifts of the N-terminal amino acids. This was in good correspondence with the stoichiometric acetylation of α-Syn previously shown in the human brain [1244,1245]. The whole in-cell NMR spectrum was clearly that of a disordered monomeric α-Syn in all cell lines. Residue-specific ¹⁵N NMR relaxation measurements were consistent with the high degree of conformational dynamics observed on IDPs *in vitro*, even though they were slowed down by an intracellular viscosity 1.5-fold higher than that in buffer. The same relaxation rates also indicated interactions with the cytosol, the residue-by-residue analysis revealing that the central region (the aggregation-prone non-amyloid-β component, NAC) was less exposed to the cellular environment than the N- and C-terminal segments. Finally, using Gd³⁺-loaded DOTA-derived cages linked by a thioester bond at different locations on the proteins, the authors measured intramolecular average distances by NMR and EPR spectroscopy, and could show that α-Syn adopted slightly more compact conformations in the cytosol than in buffer. All these results have been verified independently by Burrmann et al., using similar protocols, i.e. electrophoresis delivery in cultured human cells of recombinant, purified ¹⁵N-labeled α-Syn. They showed complementarity that the intracellular N-terminal interactions established by α-Syn in cells disappeared to a large extent upon silencing of Hsc70 and HSP90 [1250]. This chaperoning had a protective role on α-Syn binding to lipid membranes, notably those of mitochondria, and later α-Syn aggregation in cells.

As previously explained, globular and unfolded regions react very differently to interactions in cells. Globular proteins soon become NMR invisible in the case of multiple cellular interactions or interactions with high molecular weight species, as observed for ubiquitin in *E. coli* [1191]. At the opposite extreme, only the interacting residues disappear in spectra of IDPs, the other segments remaining flexible and NMR visible, as observed with α-Syn [1236,1247-1249]. An interesting effect of this differential line broadening between globular and unfolded regions was reported by the Pielak group: they built a chimera of ubiquitin and α-Syn, and observed the disappearance of the NMR signal from ubiquitin but not from the α-Syn segment of the chimera [1248]. This ability to detect unfolded regions of protein constructs in cells was exploited by Pastore and colleagues [1251]. The authors compared the effect of the bacterial environment on recombinantly expressed bacterial (CyaY) and yeast (Yfh1) orthologs of human frataxin. While the globular domain of both proteins experienced intracellular interactions that caused broadening beyond detection, the unfolded N-terminal tail of Yfh1 was detected, indicating that an important population remained free from any interactions in *E. coli*.

The crowded cellular environment is expected to affect the dynamic behavior of IDRs. Among many implications, this effect may be critical for modulating the function of nucleoporins, which take part in the formation of the eukaryotic nuclear pore complex (NPC), and harbor long IDRs containing FG-rich repeats (FG-Nups)
that are responsible (together with non-FG Nups) for the selective permeability of the NPC towards macromolecules bound to specific transport factors (TFs) [1252,1253]. Hough et al. investigated at the residue-level the conformation and dynamics of two representative fragments of the FG-Nup Nsp1 in vitro and in bacteria [1254]. Both fragments were shown by analyzing ¹³C secondary chemical shifts to remain highly disordered in bacteria. In addition, intracellular ¹⁵N T₁ and T₂ relaxation measurements compared to in vitro data revealed that the regions adjacent to the FG repeats underwent interactions with the environment, while the spacer regions remained mostly unaffected. The interaction between Nup1 and a TF was further characterized in vitro, revealing a similar pattern of weak transient interactions between the FG repeats and several binding pockets on the TF, and suggesting that these constant interactions with the cellular milieu contribute to stabilizing the disordered state and preventing the aggregation of nucleoporins. Wall and Hough further investigated the conformational dynamics of Nsp1 in yeast cells [1255]. In addition to providing the first example of in-cell NMR in S. cerevisiae, the work showed that the bacterial and yeast cytosol interacted with the Nsp1 fragment FSFG-K in a slightly different fashion, reflecting the different composition of the two environments. Specifically, the bacterial milieu preferentially interacted with hydrophobic residues of Nsp1, while the yeast protein was more promiscuous and also interacted with hydrophilic residues.

In general, as seen in the studies cited above, the main source of information when investigating IDPs/IDRs in cells comes from changes in transverse relaxation rates. However, it is not always trivial to determine the underlying physical reasons, as increased apparent relaxation rates can be caused by decreased motions in the ns timescale (due to the interaction with macromolecules) but also by the presence of motions in the µs-ms timescale and/or by increased solvent exchange rates.

### 2.4.4. Protein maturation: metal binding, post-translational modifications and redox regulation

NMR chemical shifts are excellent reporters of changes in the chemical environment of amino acids such as those due to chemical modifications or non-covalent interactions. This holds true in cells and it allows the investigation of protein maturation events such as post-translational modifications, cofactor binding and redox regulation. These are often associated with changes in tertiary or quaternary structure, including sometimes the unfolded/folded equilibrium. In this Section, we present some examples of proteins, whose proper folding was shown to depend on metal binding or redox regulation, possibly catalyzed by dedicated chaperones. Protein stability in cells is a more global question, which we treat in a dedicated subsection (2.4.6.).

NMR spectroscopy has been shown to permit the detection and real-time monitoring of protein post-translational modifications (PTMs) [1160,1163-1166,1256-1258]. The power of this approach lies in the ability of heteronuclear NMR to distinguish multiple phosphorylation events at the single-residue level. This method is particularly adapted to the characterization of PTMs on IDPs. Kinetic profiles of each species can be determined, thus providing mechanistic insights [310,1156,1160,1256]: Are residues simultaneously or sequentially phosphorylated? Are the modifications mutually exclusive? In 2008, simultaneous publications from the Selenko and Lippens groups reported the direct in-cell NMR observation of residue-specific phosphorylation in intact X. laevis oocytes [1259,1260]. After microinjecting a ¹⁵N-labeled peptide derived from the viral SV40 large T antigen regulatory region, Selenko monitored the stepwise modification of SV40 adjacent target sites of casein kinase 2 (CK2) in intact X. laevis oocytes and in oocyte extracts using time-resolved ¹H-¹⁵N-edited NMR. Moreover, upon oocyte maturation induced by a progesterone treatment,

Cdk1 became activated and the simultaneous activity of Cdk1 and CK2 on the SV40 peptide was also monitored by real-time NMR. Similarly, Bodart et al. monitored the phosphorylation of the neuronal ¹⁵N-labeled Tau protein after injecting it into X. laevis oocytes. Using ¹H-¹⁵N-edited NMR, they detected many phosphorylation sites on the injected Tau in these oocytes, some of which may be due to PKA activity. In contrast, while using the same ¹H-¹⁵N-edited NMR approach, Zhang, Liu and their colleagues observed that Tau prephosphorylated by MARK2 was dephosphorylated upon delivery in mammalian cells (HEK-293T) [1261]. Pons and colleagues used ¹H-¹⁵N-edited NMR to observe multiple phosphorylation and dephosphorylation events on the recombinant ¹⁵N-labeled “unique domain” of the c-Src protein in intact X. laevis oocytes and extracts thereof [1262]: A single phosphorylation was observed in intact oocytes, whereas multiple events were observed after prolonged incubation in oocyte extracts; they also explored the PTMs cross-talks by adding various CDKs and PKA inhibitors in the extracts. Residue-specific methionine oxidation and oxidation repair [1263], acetylation [1214] and proteolysis [1161] have also been detected in live cells. A number of reports used NMR to monitor various PTMs in cell extracts, using the same principle of real-time ¹H-¹⁵N, ¹H-¹³C or ¹⁵N/¹³Cα-¹³CO edited NMR, which we will not discuss further here (see for example [20,1156,1256,1264-1269]). ³¹P-NMR for monitoring phosphorylation has poor sensitivity and resolution in comparison, but can sometimes be useful [1270-1273].

Phosphorylation can also be detected indirectly via spectral changes of a phospho-dependent binding partner. This rationale was used by Burz and Shekhtman to detect the phospho-dependent intracellular interaction between ubiquitin and two proteins containing ubiquitin-binding domains, namely STAM2 (Signal-transducing adaptor molecule) and Hrs (Hepatocyte growth factor regulated substrate): i) they transformed E. coli cells with plasmids encoding for STAM2, Hrs, ubiquitin, and the constitutively active tyrosine kinase Fyn, ii) they coexpressed STAM2 or Hrs (or chimera including both) and, optionally, Fyn in a natural abundance medium (99.6% ¹⁴N, 0.4% ¹⁵N) and later expressed ubiquitin in a ¹⁵N-labeled medium (the opposite scenario is also possible, see Section 3.5.4.); iii) they acquired ¹H-¹⁵N NMR spectra of ubiquitin in the presence of unphospho- or phospho- STAM2 and Hrs (with or without Fyn expression). Chemical shift perturbations in the ubiquitin spectrum upon interaction revealed how the interaction surface is affected by the substrate phosphorylation state, and notably that one of the two binding patches was lost upon STAM2 phosphorylation [1274]. A similarly indirect observation of the consequences of phosphorylation on protein interactions was provided by the Dötsch group [1275]. They investigated the intracellular behavior of the peptidyl-prolyl isomerase Pin1 in X. laevis oocytes as a function of its phosphorylation state: while the active, non-phosphorylated form of the WW domain of Pin1 generates very poor in-cell NMR spectra, the PKA-phosphorylated Pin1 or the WW domain in complex with a SMAD3-peptide permitted acquisition of good quality spectra with resolved signals after injection in oocytes. Hence, the authors concluded that in-cell NMR spectra of the active Pin1 suffered from the additive effects of enhanced conformational flexibility and multiple interactions with cellular components.

A change in the cysteine redox state may not be strictly speaking a post-translational modification, but it has similar consequences in terms of changes in chemical environments and thus chemical shifts. Hence, in-cell NMR has found a very interesting application in monitoring protein redox states in live cells, which is hardly achievable with any other technique. Moreover, NMR permits monitoring of the glutathione redox equilibrium in parallel. The equilibrium between the basic two cysteine forms, thiol or disulfide bond, is often accompanied by the proper folding of the
studied protein. In this case, not only do cysteines have different
chemical shifts in their two forms, but large chemical shifts are
also observed for all the amino acids from the domain folding upon
cysteine oxidation. This has been observed using in-cell NMR for
Mia40 [1276,1277], Cox17 [1278] and SOD1 [867,1279-1285].
Hence, two sets of resonances, i.e. those from the unfolded and
those from the folded conformations, can emerge in the same spec-
trum, and their relative peak intensities give access to the quantifi-
cation of the equilibrium. This showed that the intracellular redox
potential, measured according to the GSH/GSSG equilibrium, is not
strictly linked to the redox states of proteins as measured on
Cox17, Mia40 and SOD1 in E. coli and human cells [1278], or on
thioredoxin in human cells [475]. Chaperones are of course at least
partially responsible for this discrepancy between thermodynam-
ics and kinetic regulation. For example, the reduced, unfolded state
of Mia40, which must be maintained in the cytosol before being
directed to the mitochondria, is favored by an increased expression
of the redox-regulating proteins Grx1 or Trx1. Other cysteine reac-
tions can be monitored in cells, like the appearance of sulfinic acid
upon H₂O₂ treatment [1286] or the formation of covalent bonds
with drugs [1284].

Metal ion binding also affects protein chemical shifts. Similar to
the changes associated with cysteine disulfide bonds, metal bind-
ing is often necessary for adopting the native, folded conforma-
tions, which results in extensive chemical shift perturbations for
most amino acids in the domain of interest. Although in-cell
NMR studies focused on the chelation of paramagnetic metals by
proteins are technically feasible [1287], paramagnetic metals pro-
voke fast NMR relaxation and thus low signal intensities in their
vicinity, so that to the best of our knowledge no such studies have
been carried out (not counting the in-cell NMR of hemoglobin in
erythrocytes, whose NMR signals disappear upon deoxygenation
but not because of the chelated metal itself [1288]). Nevertheless,
Banci, Luchinat and colleagues provided a very interesting exam-
ple of the importance of studying metal binding by proteins in
cells: the metalloprotein copper, zinc superoxide dismutase (Cu,
Zn-SOD or SOD1), whose mutant variants in humans are linked
to the onset of familial Amyotrophic Lateral Sclerosis (fALS)
[1289]. In order to adopt its mature, enzymatically active confor-
mation, SOD1 must dimerize, bind one Zn²⁺ and one Cu¹/²⁺ per
protomer and establish an intramolecular disulfide bond [1290].
SOD1 is thus quite complex to study because of these multiple lay-
ers of maturation steps. Notably, while SOD1 readily binds two
Zn²⁺ per protomer in the absence of copper ions *in vitro*, this does
not occur in cells according to in-cell spectra: SOD1 remains in a E-
Zn-SOD1SH (one metal-chelating site empty, the other one Zn-
loaded) form in cells not supplemented with copper, thanks to
the cellular homeostasis of zinc [1279,1280]. The full maturation
of SOD1 requires the presence of higher levels of the Copper Chap-
erone for SOD (CCS), which is necessary for Cu⁺ insertion and is
also capable of favoring the intramolecular SOD1 cysteine oxida-
tion [1280]. In-cell NMR also allowed it to be proved that some
fALS-linked SOD1 mutations, which do not perturb the structure
of the final mature protein *in vitro*, prevented Zn²⁺ binding in cells
and caused the formation of unstructured species [1282]. Further-
more, CCS coexpression can stabilize the correct fold of these SOD1
mutants, avoiding the formation of aggregation-prone species,
thereby revealing that CCS is active as a molecular chaperone in
addition to its known metallochaperone activity [1282,1283].
The same authors have also used in-cell NMR to investigate the
binding of Cd²⁺ to intracellular SOD1 [1285], and that of copper
and zinc to the putative deglycase protein DJ-1 [1286]: although
it readily occurs *in vitro*, no sign of any such metal binding could
be detected in cells, according to the absence of any chemical shift
perturbations in the in-cell ¹H-¹⁵N NMR spectra of the two
proteins.

Altogether, in-cell NMR offers good capabilities to investigate
redox reactions and ion binding, which can be greatly perturbed
*in vitro* or upon cell lysis. Hence, in-cell NMR may be quite helpful
and complementary in the pharmacological pipeline to study the
impact of cellular stress or drug exposure on the proper folding
and redox state of some pathogenic protein targets [867].

2.4.5. Nucleic acids

DNA and RNA are difficult polymers to study by NMR because of
their large size, their relatively low chemical diversity (as com-
pared to proteins) and the difficulties of producing isotopically
labeled samples. Nonetheless, NMR spectroscopy is clearly a key
approach for describing a method of choice to describe their
dynamic structural ensembles, and has provided invaluable infor-
mation in the field [1291,1292]. The influences of the intracellular
milieu on nucleic acids are many: crowding and ion binding have
drastic effects on the adoption of canonical and non-canonical
structures [1293,1294].

Early studies took advantage of the 100% natural abundance of
the NMR-visible ³¹P isotope present in nucleic acids in many
viruses, which in the early 1980's notably allowed exploration of
DNA and RNA dynamics in viruses. ³¹P has large chemical shift ani-
sotropy values, which produce ~200 ppm wide ³¹P NMR signals
with powder patterns for ³¹P nuclei of limited mobility. At the
opposite extreme, isotropic motion progressively sharpens these
signals. This typical broad, powder pattern ³¹P NMR spectra
revealed immobilized nucleic acids in a single-stranded RNA plant
virus (Tomato bushy stunt virus, tobacco mosaic virus)
[1295,1296] and in the single-stranded DNA (ssDNA) filamentous
bacteriophages fd, Pf1 and M13 [1296-1299]. Relative restriction
was also reported from other RNA plant viruses (southern bean
mosaic virus, beladonna mottle virus, turnip yellow virus, cowpea
mosaic virus, alfafa mosaic virus), but internal, low-amplitude
motions of the RNA phosphates were detected on the nanosecond
timescale, yielding ~20 ppm wide ³¹P-NMR signals [1300-1304].
Pf1 DNA was found to be well-organized and uniformly oriented,
while fd DNA was not [1305]. Precise structural information
required ¹⁵N- and ¹³C-labeling of viruses, and their analysis by
magic-angle spinning and solid-state NMR. In the late 2000's,
¹⁵N-³¹P distance measurements could be realized for T4 bacterio-
phage (containing double-stranded DNA) and revealed a B-DNA
conformation [1306]. ¹³C/¹⁵N chemical shift assignments for T7
bacteriophage were also consistent with a highly ordered B-DNA
conformation with a C2'-endo sugar conformation, according to
the previously deposited chemical shifts for B-DNA structures
[1307]. Since the early 2010's, DNA conformations of ssDNA fd
and Pf1 bacteriophages have also been characterized using ¹³C-
chemical shift assignments, which were also consistent with B-
DNA, C2'-endo conformations, and an *anti* conformation of the gly-
cosidic bond, but a with a higher tendency for base-pairing in fd
[1308-1312]. Important proximities between lysine side chains
and DNA phosphate groups were also demonstrated in T4 and fd
phages but not in Pf1 [1306,1308-1312].

Nucleic acids structure has also been studied in eukaryotic cells
from the late 2000's onwards. Concentrated ¹⁵N-labeled DNA
oligonucleotides that can form G-quadruplexes were injected into
*X. laevis* oocytes and their ¹H-¹⁵N spectra provided direct proof that
G-quadruplex structures were stable in a cellular environment
[1313-1315]. Similar experiments were carried out using ¹⁹F-
NMR on RNA or DNA oligonucleotides containing a fluorinated
moiety : ¹⁹F chemical shifts were sufficiently sensitive to distin-
guish the G-quadruplex structures and yielded background-free
NMR spectra from frog oocytes [1316-1319]. The first observations
of DNA in human cells came only in 2018, using improved pore-
forming streptolysin O (SLO) [1320] or electroporation
[1321,1322] methods for intracellular delivery. SLO pore sealing
appears to suffer some leakage that makes cells that have been treated in this way more delicate to handle [1320,1323]. These advances have apparently unblocked the situation: in the last three years, there have been NMR studies in mammalian cells, which i) proved the stability of the DNA i-motif [1321,1324], ii) monitored DNA-ligand interactions [1325,1326], iii) showed the stability of a DNA-RNA hybrid G-quadruplex structure [1327], iv) investigated the Z-DNA conformation [1328], or v) detected the binding of a RNA riboswitch aptamer to ligands [1329]. Using DNP-enhanced MAS solid-state NMR on frozen cells, Petzold and colleagues could also observe an antisense oligonucleotide that had previously been electroporated into human cells, and whose phosphorothioate backbone shows peaks at chemical shifts that differ from the natural cellular DNA [1330]. Hence, even though these DNA oligonucleotides are not integrated in the native context of chromatin, their NMR characterization in cells is full of promise. NMR spectroscopy can report RNA chemical modifications at an atomic scale in cell extracts [1331]. Fluorination schemes for nucleic acids are regularly proposed that would facilitate in-cell NMR [1332,1333], also by allowing ¹³C-¹⁹F TROSY NMR of larger objects [1334]. Improved production protocols have emerged [1335] and Covid-19-linked research on ssRNA coronaviruses or on RNA-based vaccines will probably fuel further innovation in the field. Notably, methods to characterize cellular uptake and processing of RNA are needed [1336], among which NMR spectroscopy offers the usual advantages: it can be non-destructive and provides an atomic-scale information.

### 2.4.6. Protein folding, quinary structure and large assemblies

Cellular environments can affect protein folding in multiple ways. At the outset, we want to stress the fact that folding free energy or folding thermodynamics are meaningful terms only for experiments on isolated systems at equilibrium. Hence, the concept of folding free energy applies usually to an isolated system [comprising a set of purified proteins and buffer] under constant solvent conditions. In living cells, one should rather limit the analysis to the simple concept of folded/unfolded populations.

Macromolecular crowding causes excluded volume effects that can promote folding stabilization, while attractive and repulsive interactions between the crowding agents and a given protein can reinforce or weaken a protein fold [1337-1340]. Hence, a fifth level of protein structure emerges from its interactions with the ensemble of cellular species, which is bona fide a structural organization category, a "quinary structure" [1341], in addition to being a driving force of proteins' structural behavior (Fig. 21). In-cell NMR clearly has a role to play in studying this integrative phenomenon.

Early observations had shown that different proteins can establish what could be called different densities of intracellular interactions, as revealed by the variable NMR-detectability of intracellular proteins. In the late 1990's, Brindle and colleagues reported that two yeast enzymes showed NMR spectra consistent with a ~2 times higher viscosity in yeast cells than in buffer, while two other proteins were not NMR visible at all in intact cells, although they became detectable upon cell lysis [1179,1343]. The broadening beyond detection in the latter two cases revealed lower average molecular tumbling rates, which in turn showed that the apparent local viscosity measured for the two NMR-visible enzymes was not universal. This information was obtained by recording ¹⁹F-NMR spectra of yeast cells supplemented with 5-fluorotryptophan and overexpressing the proteins of interest. Overall, this revealed differential intracellular motional restrictions. In the early 2010's, several studies attempted to understand this effect. Gierasch and colleagues measured the intracellular ¹H-¹⁵N TROSY effect on overexpressed GB1 in live *E. coli* to extrapolate an intracellular viscosity, which was only 2-3 times higher than that of aqueous buffers [1191]. ¹⁹F-labeling strategies were used by Pielak, Li and colleagues to deconvolute viscosity and cellular interactions in *E. coli* expressing recombinant GB1 and ubiquitin. While GB1 was only weakly involved in cellular interactions, ubiquitin experienced important transient, unspecific binding in the heterologous bacterial cytosol [1344]. Using ¹⁵N-edited NMR, Gierasch and colleagues reported the same cellular interactions on small model proteins NmerA and ubiquitin expressed in *E. coli*, and could show that a triple mutant ubiquitin-L8A-I44A-V70A provided much bet-

<figure><img src="image_25.png" /><figcaption>**Fig. 21.** Representation of the impact of quinary interactions, i.e. the interactions of a protein with its native cellular environment, which are proposed to create a fifth level of protein structure. This scheme represents the possible phenomena specific to the intracellular milieu that may affect protein stability. Unlike purified proteins in aqueous buffers (shown in a), high macromolecular concentrations as shown in b) can produce crowding, which may favor protein compactness and folding, while both specific and non-specific interactions with other cellular components may either stabilize or destabilize a protein's fold, or provoke phase separation, oligomerization or even aggregation. Adapted from Gopan et al. 2021 [1342].</figcaption></figure>
ter spectra in cell lysates [1191]. Hydrophobic patches at the surface appeared to favor unspecific interactions in cells.

Hence, intermolecular interactions were mainly responsible for a certain motional restriction in cells, which provoked the broadening beyond detection of some globular proteins. The impact of these cellular interactions on protein stability had to be investigated.

Let us first extend the picture by considering also intracellular electrostatic interactions. In parallel, Crowley et al. employed a combination of in-cell NMR spectroscopy and SEC analysis to determine the origin of the diffuse interactions of cytochrome c in bacterial cells, which prevented NMR detection [1345]. By analyzing the SEC elution profile of CytC in crude cell lysates, in the presence of increasing amounts of salt, CytC was shown to interact electrostatically with negatively charged components, through a positively charged Lys/Arg-rich area on the protein surface. With the same approach, the Crowley group showed that electrostatic interactions mediated by the Lys/Arg-rich HIV-1 Tat peptide fused to GB1 induced 'stickiness' in cells and in lysates [1346], and later showed by ¹H-¹⁵N and ¹⁹F in-cell NMR that these electrostatic interactions increased gradually as a function of the length of a poly-Arg tail fused to GB1 [1347], thereby suggesting a general, non-specific interaction with negatively charged cellular components. The Banci group showed that proteins undergo both diffuse and partner-mediated interactions, depending on the composition of the cellular environment. Barbieri et al. analyzed a series of surface mutants of human profilin in the cytosol of bacterial and human cells [1348]. Profilin is a small soluble actin-binding protein that is also involved in several functional interactions with proteins harboring poly-proline regions, and with phosphoinositides. In bacteria, which lack the functional partners of profilin, a freely-tumbling protein could be obtained by mutating just the surface Lys/Arg residues, which were responsible for the diffuse electrostatic interactions, whereas in human cells the free protein was observed only after all three functional interactions were abolished by introducing mutations on specific surface residues. Both Tat-GB1 and profilin also experienced interactions in the bacterial lysate and, remarkably, it was observed that treatment with Mg²⁺ and RNase abolished the interactions, suggesting that a common cause of NMR line broadening was the interaction with bacterial RNA [1346,1348].

In the mid-2010's, Danielsson, Oliveberg and colleagues examined the effects of both hydrophobic and electrostatic quinary interactions on the NMR spectral features in the cytoplasm of *E. coli*. Mu et al. applied in-cell NMR to estimate the intracellular rotational diffusion of three evolutionarily unrelated proteins, bacterial TTHA, human HAH1, and human SOD1 β-barrel [1349]. Surface mutations were introduced on each protein to evaluate the electrostatic dependence of rotational diffusion in cells. While the WT proteins had different rotational diffusion rates, analysis of the mutants revealed that all three proteins showed a common diffusion dependence on net charge density, surface hydrophobicity and electric dipole moment, suggesting that intracellular macromolecular interactions follow a simple set of physicochemical rules. To decode these rules, Leeb et al. analyzed the intracellular T₁ and T₂ relaxation measured for the above set of proteins and charge mutants introduced in the cytoplasm of human cells, and found that they are inconsistent both with a simplistic 'increased viscosity' model [1350] and with simple exchange broadening. Instead, it fits within a theoretical framework where the anomalous T₁ increase and T₂ decrease arise from the formation of transient complexes with intracellular macromolecules, in fast exchange with the free protein. Good agreement with the experimental data was obtained when considering a proteome-like distribution of particle sizes. Fitting of relaxation data with the above model allowed estimation of the fractional population of

the bound state of each protein. Proteins with a less negative net charge were found to have a higher bound state fraction, while a plot of the bound population as a function of net charge revealed the 'basal' interaction propensity of each protein, independent of its net charge.

Complementary observations have been reported by the Shekhtman group. When they first applied in-cell NMR to observe a protein overexpressed in *P. pastoris*, they observed that different metabolic states triggered a change in localization of overexpressed yeast ubiquitin [1192]. When methanol was used as the carbon source, ubiquitin was uniformly distributed in the cytosol and was detected by ¹H-¹⁵N in-cell NMR, whereas when a mixture of dextrose/methanol was used ubiquitin was mainly localized in intracellular vesicles or granules, and was undetectable by in-cell NMR. That finding pointed to a strong connection between cellular metabolism and quinary interactions. The same group further investigated the molecular origins of quinary interactions. Majumder et al. showed that RNAs are an important component of protein quinary interactions both in bacteria and eukaryotic cells by measuring the efficiency of cross-correlated relaxation-induced polarization transfer in ¹H-¹⁵N CRINEPT-HMQC-TROSY spectra [1351]. This approach allowed an estimation of the average size of the complexes occurring between a set of globular proteins and intracellular components. By comparing the intracellular behavior with proteins alone or in the presence of RNA *in vitro*, the authors concluded that RNA is a key component of quinary interactions. The group further investigated the metabolic dependence of quinary interactions previously observed in yeast [1192] and observed by confocal microscopy an increased colocalization between ubiquitin and RNA when *P. pastoris* was grown in a mixed dextrose/methanol medium, together with a change in the total cellular RNA content and its size distribution [1352]. In later works, Shekhtman and coworkers showed that proteins can interact with ribosomes. DeMott et al. investigated such interactions *in vitro* and in cells and observed that the enzymes adenylate kinase, dihydrofolate reductase and thymidylate synthase could bind intact ribosomes with micromolar affinity, and that such interactions modulated their enzymatic activity [1353]. GFP also interacted with ribosomes, leading to decreased translational diffusion. In later work, the group developed an NMR bioreactor and applied it to monitor in real time the behavior of intracellular thioredoxin in response to cell treatment with antibiotics binding to the small ribosomal subunit [1354]. Treatment with these antibiotics resulted in an increase of quinary interactions between thioredoxin and mRNA, further strengthening the notion that, for some proteins, quinary interactions are strongly modulated by the intracellular content of mRNA, rRNA and by the assembly state of ribosomes. Such interactions could explain the dependence of protein rotational diffusion on the intracellular metabolic state previously observed in yeast by Bertrand et al. when changing growth media [1192]. Consistently, Sugiki et al. observed a marked change in protein rotational diffusion in bacteria when inducing GB1 expression for different times and at different optical densities [1355], suggesting that metabolic changes affect the extent of quinary interactions also in *E. coli*, possibly through a change in RNA and ribosome content. Other hypotheses might also be appropriate, such as the metabolite content, the metabolic state and the capacities of the chaperone ensemble to bind to the folded and unfolded forms of the overexpressed protein, or even to catalyze their proper folding.

What is the impact of all these intracellular interactions on protein stability? The Pielak group has extensively studied the effect of crowding agents on protein folding, and turned to in-cell measurement in the early 2010's. In 2011, they observed that the bacterial cytosol could not force the folding of a protein carrying mutations, destabilizing it by only 1 kcal/mol [1356]. This implied that the stabilizing effect of volume exclusion was compensated by
destabilizing interactions with the crowding agents. To quantify
these effects at the residue level, Monteith & Pielak investigated
the effects of transient opening of the folded structure in the small
model protein GB1 in bacteria, which system is well-suited to H/D
exchange measurements by NMR. By placing bacteria overexpress-
ing GB1 in D₂O over periods of hours, and by measuring H-amide
peak intensities after quenching the exchange (at low pH/Temp)
they could quantify H/D exchange in a residue-specific manner.
They analyzed the impact of the cellular milieu on the free energy
of fold opening and concluded that GB1 is stabilized in bacteria by
about 0.75 kcal/mol, whereas crowding agents in vitro may have
the opposite effect [1357]. However, after realizing that the intra-
cellular pH was much lower than expected, they concluded in a
correction that GB1 was actually destabilized by about 1 kcal/mol
[1358]. Using ¹⁹F-labeling and ¹⁹F-detected NMR, they observed
similar destabilization for a GB1 mutant in a later paper [470],
and almost no effect on a small SH3 domain [1359], always in
E. coli. In the next paper, using measurements of H/D exchange
in bacteria, they showed that surface mutations of GB1 were more
stabilizing in cells than in buffer [1360]. They noticed that this cel-
lular stabilization was comparable to that from a classical protein-
protein interaction, thereby putting at the same level functional
interactions with specific partners and the non-specific cytosolic
interactions. Again using H/D exchange in bacteria placed in D₂O,
Pielak and colleagues explored the effect of intracellular pH
[470,1361] and observed stronger destabilization of GB1 in acidic
conditions. This led them to the conclusion that low pH would
cause an accumulation of positive charges on cellular proteins,
which, in turn, would exert increased numbers of destabilizing
interactions with the negatively charged GB1. Finally, they
observed a destabilization of ¹⁹F-labeled SH3 in bacteria submitted
to osmotic shock, which could be compensated by addition of gly-
cine betaine [1362]. We must acknowledge the fact that there are
daunting questions in characterizing quinary interactions of
heterologous proteins in variable bacterial environments. It is dif-
ficult to know whether the bacterial proteome, the metabolome,
chaperone activity, the amount of unfolded cellular species, etc...
are constant after the cells have been transferred from H₂O into
D₂O for periods of hours. It is still more difficult to think that it
is the case in low pH or high salt conditions. All of these conditions
might stabilize/destabilize the unfolded/folded states of the pro-
tein of interest.

The contribution of electrostatic interactions to folding stability and quinary structure in living cells was also investigated using solution NMR by the Oliveberg group. Danielsson et al. investigated the thermal unfolding of the SOD1 β-barrel, a synthetic model protein showing a simple two-state unfolding equilibrium *in vitro* [1363], which is amenable to intracellular delivery [1364,1365]. By integrating the volume of the crosspeaks corresponding either to the folded or to the unfolded species, the authors compared the temperature dependence of the “folding free energy” of SOD1 β-barrel purified *in vitro*, overexpressed in *E. coli* and delivered by electroporation in human adenocarcinoma cells [1365]. The concept of folding free energy should be taken with caution here, as discussed below at the end of this section. Consistent with what has been observed for other globular proteins in bacteria, SOD1 β-barrel was markedly destabilized in the cytosol of both bacteria and human cells, leading to a decreased melting temperature and, notably, moving both cold and heat unfolding into the physiological regime. Also in this case, the effect was attributed to interactions with cellular components that stabilize the denatured state of the protein. In most organisms, the intracellular environment behaves like a polyanionic system due to the average net-charge of the macromolecules being negative [1366]. Based on this assumption, Sörensen et al. investigated *in vitro* the destabilization of SOD1 β-barrel using polyacetate ions and

observed that, similar to what is observed in cells, the apparent
SOD1 destabilization originated from favorable interactions
between polyacetate and the unfolded state of the protein
[1367]. More recently, Iwakawa et al. studied the effect of a nega-
tively charged crowding agent, lysozyme, on SOD1 β-barrel struc-
ture and oligomerization. Lysozyme slowed down the protein
oligomerization, while NMR analysis of the monomer revealed that
the interactions between lysozyme and SOD1 stabilized the latter
in alternative excited state [1368].

Overall, we have seen that the intracellular environment can
have disruptive effects on the NMR spectrum of a macromolecule.
Indeed, the increased viscosity of the cytosol, the interactions with
other cellular components and the possible alteration of the fold-
ing/unfolding equilibria result in signal broadening. This broaden-
ing commonly renders many signals undetectable when studying
macromolecules by solution in-cell NMR. We discuss this more
thoroughly and provide orders of magnitude concerning protein
size, viscosity, multiple interactions and relaxation effects in
Section 3.

We want to give a quick cautious note to conclude: varying
temperature, pH or salt concentrations can have strong effects on
cells, their native contents, and their capacity to maintain home-
ostasis, and all these can affect the studied protein. As far as in-
cell studies are concerned, we recommend care using the term
"folding free energy". If they are healthy, living cells are anyway
not systems that let their components evolve towards thermody-
namic equilibria. Examples have been reported showing that cellu-
lar stress or cell growth phase can have an impact on intracellular
NMR spectra [1355,1369]. The same comments apply to other
methods using thermal profiling followed by mass-spectrometry
[1370-1375] or in-cell FRET readouts [1342,1375-1378]. Nonethe-
less, NMR experiments can reveal crowding and viscosity effects,
intracellular interactions and their impact on folding and binding
populations. Such knowledge will certainly help refine perceptions
of protein structure and therapeutic strategies.

2.5. Cellular drug assays

Drug research pipelines have incorporated NMR spectroscopy
fruitfully at several steps of drug development [1379-1381]. Here
again, the versatility of NMR permits a broad range of applications,
including fragment-based approaches [1382,1383], competition or
affinity assays [1384], structure-based design [1385,1386], enzy-
matic reaction monitoring [1387], metabolic footprinting
[40,1388], among others. All these possibilities have also been
applied in cells [19,22]. The benefits of studying drug:target inter-
actions directly in cells are evident: the simultaneous assessment
of drug membrane permeability, cellular availability, drug:protein
binding and eventually dose-dependent binding curves *in situ* is
clearly appealing. Moreover, the experiments are not necessarily
costly or convoluted, they are even quite simple in some cases as
we show below.

More common approaches than in-cell NMR techniques exist to
monitor ligand impact on cellular activities. These are discussed in
Section 4.

2.5.1. *In-cell drug screening through binding*

Different strategies have been used to develop in-cell NMR drug
screening protocols. We distinguish three categories: observing
either the target, the candidate drug compounds, or a reporter of
the target activity.

Shekhtman and colleagues have used the selective observation
of a ¹⁵N-labeled protein target, focusing on the inhibition of het-
erodimeric protein:protein interactions [1389]. We discuss details
of the corresponding protocols for sequential protein expression in
Section 3 (subsection “Protein:protein interactions”). The protein
target is observed using a ¹⁵N-editing pulse sequence, so that the ¹⁴N cellular content is rendered “NMR-invisible”. The NMR spectra (usually 2D ¹H-¹⁵N correlation spectra) or the protein of interest are different in the presence and absence of the interaction with the protein partner. Hence, drug screens have been performed to identify inhibitors of the interactions between i) FKBP and FRB, two components of the mTOR pathway controlling notably T-cell activation [1390], and ii) the mycobacterial ATPase MPa and the ubiquitin-like protein Pup [1391]. The compounds identified later showed inhibiting activities in yeast and mycobacteria. Luchinat, Banci and their coworkers also recently carried out an in-cell screen on human carbonic anhydrases CA1 and CA2 in cultured human HEK293T cells [1392]. The study was facilitated by the fact that CA1/CA2 histidine side chain amide protons yield NMR signal in a remote, usually empty region of ¹H-NMR spectra, between 11 and 16 ppm. Drug binding was detected by CA1/CA2-histidine peak shifting in one-dimensional ¹H-NMR spectra. In principle, about 100 compounds may be screened per week using such a setup, in combination with the matrix approach employed by Shekhtman. Although it requires prior cell transfection and overexpression of the target protein, this approach permits detection of the binding of a compound to the enzymatic site of a human protein target in a human cellular background. Even better, it can report the dissociation of a compound from its target, revealing possible engagements with off-target cellular entities [1393].

Enzyme activity reporters have been shown to be promising instruments for in-cell NMR drug screening. Breeze, Hu and their coworkers have for example used meropenem, an antibiotic degraded by the New Delhi metallo-β-lactamase subclass 1 (NDM-1), an infamous bacterial drug resistance enzyme. They first established real-time monitoring of meropenem hydrolysis by NDM-1 overexpressed in live E.coli: methyl NMR signals of intact and degraded meropenem are not the same (Fig. 22), and can be used as reporters of progress of the reaction using one-dimensional ¹H-NMR spectroscopy [1394]. Next, these cells were grown in 96-well plates, exposed to a collection of compounds and NDM-1 inhibition assessed by integrating NMR signals of intact and decomposed meropenem after quenching the reaction [1395]. The authors calculated that their protocol would permit screening of about 10,000 compounds per week. Garau, Dalvit and their coworkers used a similar approach, but this time by recording ¹⁹F-NMR one-dimensional spectra of ARN1203, a fluorinated analog of the cannabinoid neurotransmitter anandamine. Its ¹⁹F-moiety has different chemical shifts before and after ARN1203

degradation by the fatty acid amide hydrolase (FAAH). The corresponding NMR peaks proved to be valuable probes of FAAH activity in situ, as shown using membrane extracts of HeLa cells overexpressing this protein [1396]. HEK293 cells overexpressing FAAH in 24-well plates were later used to screen compounds [1397]. This approach is appealing: it relies on simple, sensitive, background free ¹⁹F-NMR spectroscopy [1387,1398] of the supernatant pipetted from the culture plates. Moreover, it allows an approximate cell toxicity assay by using ¹H-NMR for the quantification of lactate production [1397].

As shown in subsection 2.3, “Metabolic activities” (see above), a number of molecules probing specific cellular activities using an NMR-readout have been developed. Many of them may be useful instruments for in-cell NMR drug screening. In particular, other cleavable ¹⁹F-reporters have been published to detect specific enzyme activities in cells [1399,1400]. MRI ¹⁹F-reporters for detecting enzymatic activities have also been developed: they contain a paramagnetic moiety, which destroys the NMR signal of the ¹⁹F moiety as long as the reporter is intact; upon cleavage by the enzyme of interest, the two moieties are separated and the ¹⁹F NMR signal becomes observable [137,1401,1402]. These have often been tested in cells [1403-1407] and some of them may be used for drug screening. Other ¹⁹F-, ¹³C- or ¹⁵N-labeled reporters of kinase activity [1408,1409] or methionine sulfoxide reductase [1410] may also be useful for example.

To the best of our knowledge, ligand-based NMR detection has not been used to achieve drug screening in live cells. The most closely-related study that we found was a Saturation Transfer Difference (STD-NMR) screening study of Rademacher and Peters on norovirus virus-like particles (VLPs, auto-assembled particles made of viral capsid proteins) [1411]. The usual NMR screening methods, like STD-NMR, waterLOGSY, or relaxation weighted spectroscopy detect weak interactions showing affinities in the micro-to milli-molar range and fast association/dissociation [1386,1412]. Hence, they are likely to generate high rates of false positives, even when recording experiments with and without expression of the protein target: weak interactions can be detected with cellular content, which itself is not perfectly reproducible and constant. Nikolaev, Allain and their coworkers have for example detected significant numbers of weak interactions between metabolites (notably those containing aromatic rings) and common E. coli proteins [1413,1414]. This implies that the metabolic states should be very similar between cell aliquots to avoid different competition levels between screened drugs and metabolites. These ligand-based NMR

<figure><img src="image_26.png" /><figcaption>**Fig. 22.** ¹H-NMR analysis of antibiotic degradation by living bacteria: times series of spectra can be recorded, which report notably the evolution of antibiotics in the supernatant in real-time. A) One-dimensional ¹H spectra showing the progressive hydrolysis of Meropenem by purified NDM-1 drug-resistance enzyme, B) in presence of living cells of a standard E. coli strain and C) in presence of an E. coli strain expressing NDM-1. D) Time-dependent evolution ¹H-NMR signal intensities of various antibiotics in presence of E. coli cells expressing NDM-1.</figcaption></figure>
detection methods (STD-NMR, TRNOE, waterLOGSY,...) are thus
more adapted to “on-cell NMR” studies, which we detail below.

Other methods using paramagnetic tagging of protein targets [1415,1416] may be better adapted to these kind of assays: they may permit a more specific signal broadening of compounds binding to a defined protein carrying a paramagnetic center. We would not be surprised to see such approaches being developed in the coming years.

2.5.2. *Drug modes of action and cell penetrance*

The present subsection presents two aspects of drug treatment, which relate to i) the effects of drugs on the overall profiling of cellular composition, and ii) the detection of NMR signals revealing intracellular drug molecules, hence drug penetrance in cells. To some extent, this subsection covers studies that were also presented in Section 2.2.2., which treated the characterization of peptidoglycan and its interaction with antibiotics, and Section 2.3., which focused on monitoring metabolism, sometimes in presence of drugs. Hence, we will not repeat here explanation of the principles of NMR readout used in these studies.

The mechanisms of action of antibiotics have been studied on intact cell walls from the early days of NMR in biology, and on whole cells from the 2000's. Early solution NMR observations had revealed that the peptidoglycan (PG) residue ¹⁵N-D-Ala of the Gram-positive *B. licheniformis* became more mobile upon exposure to vancomycin [582]. Lower cross-linking in PG was later observed in the presence of penicillin G or benzylpenicillin, using ¹⁵N CPMAS solid state NMR of the isolated cell walls of *A. viridans* labeled with ¹⁵N-Lys [584,1417]. Because the PG of *B. subtilis* does not contain any lysine, the incorporation of L-[2-¹³C-¹⁵N]-Asp was used to record ¹⁵N-REDOR-filtered ¹³C-solid state NMR spectra and to evaluate the decrease of PG cross-linking index upon cephalosporin treatment [595,596]. In the 2000's and 2010's, combinations of ¹³C- and ¹⁵N-amino acid supplementation were used to execute isotopic ¹³C or ¹⁵N-REDOR-filters followed by ¹⁵N- or ¹³C-NMR

detection, which allowed quantification of PG cross-links on bacte-
rial cells exposed to various antibiotics (notably vancomycin,
amphomycin, plusbacin, oritavancin on S. aureus and E. faecium)
(Fig. 23) [578,597-601,610,615,619]. The effects of antibiotics on
D-Ala incorporation in the surface teichoic acid polymer have also
been scrutinized by solid state NMR of Gram-positive bacteria
[589,598,599,615,620,621]. Lately, Cegelski and coworkers have
also reported solid-state NMR ¹³C and ¹⁵N spectral signatures of
whole bacterial cells at natural abundance, which allow the rela-
tive quantification of PG and WTA, thus permitting a rapid evalua-
tion of the mechanism of action of antibiotics [579,1418]. Similar
applications may emerge for the analysis of bacterial or fungal
extracellular matrices [1419,1420] or mammalian cells, whose
whole-cell solid-state NMR spectra show visible changes depend-
ing on their metabolic state [1421,1422] or upon protein synthesis
inhibition for example [1423]. The level of accuracy on the meta-
bolic state provided by ssNMR is however rather limited and prob-
ably less adapted to the characterization of minute metabolic
changes in mammalian cells. HR-MAS NMR is more likely to pro-
vide useful insights in such situations (see Sections 2.3.5. and
3.3.4.). It has indeed been used to study the effects of cell exposure
to various compounds like chemotherapeutic agents [1008-1014]
or pathway activators [990,1015,1016]. Unless it is performed at
low spinning rate, HR-MAS does affect cell viability and does not
permit real-time NMR monitoring of cells upon drug treatment.

The drug mechanisms can also be characterized on a structural
level. We described in-cell studies aiming at mapping drug candi-
date epitopes in Section 2.6. Concerning proven drugs and their
tridimensional structures in situ, we found mostly reports on
antibiotics. Since the 2000's, using similar solid-state REDOR
NMR spectroscopy, Schaefer and his coworkers progressively
revealed the structural details of drug binding in bacterial cell
walls. The presence of ¹⁹F nuclei in antibiotic molecules has been
helpful, because it allows ¹³C-¹⁹F or ¹⁵N-¹⁹F REDOR measurements
for example. Hence, coupled with ¹³C- or ¹⁵N-amino acid specific

<figure><img src="image_27.png" /><figcaption>**Fig. 23.** Example of an application of solid-state NMR (ssNMR) to whole bacterial cells (*S. aureus*) to characterize the effects of antibiotics: here, vancomycin and oritavancin inhibit the incorporation of D-Ala in wall teichoic acid (WTA). **A)** The cells are grown in a medium supplemented with L-[1-¹³C]Lys and L-[D[¹⁵N]-Ala, which are incorporated in the peptidoglycan (PG) and wall teichoic acid; only the ¹⁵N-NMR signals of constrained species are visible using solid-state NMR techniques. **B)** ¹³C-ssNMR spectra using REDOR methods showing either (lower) ¹³C-labeled species, here mostly L-[1-¹³C]Lys, or (upper) ¹³C-labeled species bound to ¹⁵N-labeled species, i.e. L-[1-¹³C]Lys-[D[¹⁵N]-Ala]. **C)** Close-up view of ¹⁵N-NMR spectra of whole *S. aureus* treated either with no antibiotic, with vancomycin or with oritavancin. **D)** ¹⁵N-ssNMR spectra obtained using REDOR methods showing either (lower) ¹⁵N-labeled species, here mostly [¹⁵N]Ala, or (upper) ¹⁵N-labeled species bound to ¹³C-labeled species, i.e. D-[¹⁵N]Ala-[L-[1-¹³C]-Lys]. Adapted from Singh et al. 2017 [621].</figcaption></figure>
labeling, this allows combined isotope editing and internuclear distance measurements for those rare events where antibiotic molecules bind to PG precursors [578,589,597,600,607-609,619,1424]. Other ssNMR studies have been carried out to solve the structures of nisin or teixobactins binding to the bacterial Lipid-II in native membranes, using recently developed ¹H-detected ssNMR techniques and equipment (see Section 2.4.2.) [577,625,626].

An important aspect of drug activity is the ability to penetrate cells. In many cases, the process starts with entry of a cell-penetrant pro-drug followed by its intracellular chemical modification to generate the final active compound. NMR spectroscopy has interesting capabilities in this regard, because of its atomic scale resolution combined with real-time measurements. These were explored by Lippens and colleagues in the early 2000's, to monitor the pro-drug ethionamide penetration and activation in mycobacteria using HR-MAS NMR [1023,1024]. We have seen earlier that NMR is very well suited to characterize diffusion coefficients, and Lippens and coworkers used this ability to distinguish intracellular from extracellular molecules by employing diffusion filters within pulse sequences to separate signals. This provided a definitive proof and time-monitoring of the localization of the prodrug. More recently, Luchinat, Banci and their coworkers tracked drug penetration by observing the NMR signal displacements of a protein target, the human carbonic anhydrase 2 (CA2) [867,1392,1393]. After its overexpression in cultured, transfected cells, this protein provides ¹H-NMR signals in empty windows of the spectrum, because of its catalytic histidines, whose side chain imidazoles generate slowly exchanging NH resonances at 11-16 ppm. These resonances shift upon binding to drugs, thus reporting notably drug cell penetration. This approach may be adapted to a number of protein targets, but it requires of course finding convenient, well-observable NMR signals that report drug-protein binding. Hence, it will probably require finding a specific amino acid isotope labeling adapted for every novel target.

## 2.6. Epitope mapping, structure-activity relationship

Structure-activity relationship (SAR) studies refer to hit or lead compound optimization, which has been a recognized application of NMR to structural biology for about 25 years [1425-1427]. Extensive details about the typical methods dedicated to SAR by NMR (STD-NMR, TRNOE, waterLOGSY, and others) can be found in these latter citations, and will not be discussed further here; specific examples of applications to cellular samples were discussed in Sections 3.4.2 and 3.4.3.

### 2.6.1. Virus receptors

We will first mention NMR studies of virus:ligand interactions. We will not take part in the debate of whether viruses are really cells or live organisms, which probably depends very much on the chosen definition of life [1428,1429]. However, virus:ligand NMR investigations are included here, because they represent an important body of work in the development of on-cell NMR studies. NMR spectroscopy can indeed help to map the epitopes of ligands binding surface proteins of viruses. This is of interest because many viruses attach to cellular surface glycans to favor their entry into appropriate host cells, which also often implies the existence of a glycan-cleaving enzyme for the later release of replicated virus particles [1430-1432]. Hence, characterizing these recognition events can help to understand viral infection mechanisms, including possible inter-species or inter-individual transmission, and to design antiviral therapeutics [1433-1437].

In 1992, the first publication that we found in this field reported the use of intact H3N2 influenza virus at 2-4 g/L and the observation of ¹H-NMR signals of sialoside oligosaccharides upon titration from 1 to 20 g/L [1438]. Binding affinities in the millimolar range

were extracted and the virus preferences for the various surface oligosaccharides were discussed. This approach required high quantities of virus and of glycans and was likely to provide information only on weak affinity interactions. The situation was improved by the exploitation of Saturation-Transfer Difference NMR (STD-NMR) spectroscopy, a concept already exploited in the late 1970s' [1439-1441] that can provide a spectroscopic footprint of binding epitopes (a number of dedicated reviews have been published on this subject, see [1384,1442,1443]). Soon after being recognized as an efficient drug discovery method in the late 1990's [1412], STD-NMR was used to map the epitope of an antiviral compound binding to human rhinovirus particles [1444]. These results were obtained using a virus concentration of 20 nM, i.e. 170 mg/L, and antiviral compounds at 120 µM. Similar STD-NMR studies described glycan preferences of virus-like particles, i.e. assemblies of viral structural proteins that do not contain any viral genetic material. The molecular basis of glycan epitope recognition was described at the atomic-scale for various influenza serotypes [1445-1447], rabbit hemorrhagic disease virus [1448], bovine norovirus [1449], rhesus rotavirus [1450], and human norovirus [1411,1451,1452].

Virus-like particles have also been used to detail the interaction between peptide inhibitors and hepatitis B virus [1453], and between fusion inhibitors and influenza hemagglutinin [1454]. In the last 5 years, investigators reported studies using whole virus particles to understand the adaptation of various influenza serotypes to different glycans and thus different species [1455,1456], to verify the proper binding of fluorescent probes to enteroviruses binding [1457], and to map the epitopes of compounds inhibiting the entry of the enterovirus EV71 [1458].

Recently, Vasile and colleagues published STD-NMR studies using transfected mammalian cell lines stably expressing functional influenza hemagglutinin on their membranes: using 10⁷ cells and avoiding purification steps, they could achieve a rapid epitope mapping of various ligands [1459,1460]. The wealth of structural information from all these STD studies would be difficult to describe fully. The corresponding results may be seen as overlapping with those of mass-spectrometry, glycan arrays or crystallography. However, results from isolated viral receptors can often be unreliable [1461,1462], crystal structures of protein:glycan complexes are often limited to the resolved description of very few glycan residues, and studies on viral particles clearly provide a useful support to the *in vitro* toolbox [1463].

### 2.6.2. Mammalian membrane proteins

Mammalian cell transfection has also been useful to characterize the binding epitopes of drug candidates targeting membrane proteins by NMR spectroscopy *in situ*. Membrane proteins represent an abundant reservoir of drug targets [1464], but difficulties with their purification and stability for *in vitro* studies often make their study extremely problematic.

In this context, STD-NMR has the capacity to provide ligand mapping without the classical purification steps. Jimenez-Barbero and colleagues published a proof of concept in 2005, where they showed that mannan STD-NMR signals were detected in the presence of K562 cells transfected with the immune system lectin-like receptor DC-SIGN, but not with mock-transfected cells [1465]. This approach was applied to a human Golgi resident nucleotide sugar transporter (called CST) expressed in *Pichia pastoris*: STD signals of CMP-sialic acid were recorded in the presence of Golgi-enriched fractions of cells either expressing CST or not [1466], and, in a subsequent study, expressing CST mutants that affect binding or saturation transfer [1467]. In the absence of 3D structural data, this provided a primary mapping of the interaction both on the ligand and on the membrane-protein. Similar experiments were carried
out on GDP-mannose and the GDP-mannose transporter of *Aspergillus niger* [1468].

The extremely important drug-target family of G-Protein Coupled Receptors (GPCRs) has also been studied by on-cell STD-NMR spectroscopy, confirming *in situ* binding of ligands on i) insect cells expressing and exposing human cannabinoid receptors (the NMR samples containing 5.10⁶ cells/mL exposing the receptor at about 100 nM) [1469]; ii) membrane fractions of mammalian cells expressing the human sweet receptor and an inactive mutant [1470,1471], the chemokine receptor CXCR4 [1472], and the thiol odorant receptors MOR244-3 [1473] and OR2T11 [1474] (the NMR samples containing about 10 nM of the relevant GPCRs). These studies often report STD signals even without expression of the receptor, probably because of unspecified interactions of the tested hydrophobic ligands. Spectroscopists have thus used Saturation Transfer Double-Difference (STDD-NMR) by subtracting the STD signals obtained in presence and absence of receptor expression. Naturally, this compromises sensitivity. However, such experiments provided direct evidence of GPCR binding, and led to an epitope mapping. This approach is highly complementary to the more common luciferase GloSensor assays, which report end-point, indirect evaluations of GPCR activation through cAMP production [1472,1474].

Another issue comes from the tendency of adherent mammalian cell lines to aggregate and settle, which often forces the use of suspension cells or membrane fractions. Jimenez-Barbero and colleagues proposed a solution: they introduced 1 million cells in a 4 mm diameter cylindrical sample container, conventionally called a “rotor”, and spun them between 1 and 4 kHz using a HR-MAS probe (see Sections 2.2.3. on LPS, 2.3.5. on lipids and 3.4.3. for specific discussion of HR-MAS). This caused cell sedimentation and the formation of a homogeneous layer on the rotor walls, which permitted STDD-NMR measurements for a glucose co-transporter and some ligands [1475].

This sedimentation issue is one of the reasons that pushed spectroscopists to use slow-settling platelets for a number of early on-cell STDD studies. Another convenient aspect when using platelets is their very high content of integrin αIIbβ3 (1-5.10⁴ per cell, about 50% of membrane proteins [1476]), which is required for platelet aggregation. Combined with a low cytoplasmic volume, platelets were thus ideal for early characterizations of ligand binding to intact mammalian cells by NMR (integrin αIIbβ3 was at a concentration of about 600 nM in the NMR samples) [1476]). Because native ligands of integrins are short peptides containing an RGD motif, resolving their receptor-bound conformations was of considerable interest. Another NMR method known for a long time, named TRNOE, had been developed for such applications: it detects the large negative NOE of slow tumbling species, which reveals a footprint of the bound-ligand conformation (via interatomic distances) even though it is detected on the free ligand population [1412,1477]. Applied to platelets, the measured TRNOE allowed the bound conformations of two cyclic RGD-containing peptides to be solved [1478]. TRNOE spectroscopy was later used to characterize the interaction between RGD-containing peptides and the integrin αvβ3 expressed at high levels by certain cancer cell lines (generating nM range concentrations in the NMR samples) [1479-1484], or with the integrin αvβ1 [1485,1486]. These studies led to design of RGD-containing peptidomimetics showing improved affinity and specificity for the studied integrins. Recently, STD epitope mapping was carried out for the large protein osteopontin binding to integrins, using mammalian cells entrapped in a methylcellulose hydrogel [1487], which seems to us a promising approach.

In the last 5 years, two remarkable on-cell STD-NMR studies led to the progressive design of high-affinity ligands of the B-cell receptor Siglec-2 [1488] and of CXCR4 [1489]. STD-NMR or TRNOEs

also confirmed the binding of some peptides to the vascular endothelial growth factor receptors VEGFRs [1490,1491] or the angiogenesis associated CD13 [1492], and permitted mapping at a residue-scale of the Bombesin peptide interaction with the GPCR Gastrin Releasing Peptide Receptor (GRP-R) [1493].

Finally, another interesting approach was explored by Izadi-Pruneyre and colleagues, aimed at providing direct experimental evidence of drug binding to a putative membrane protein target [1494]. A series of candidate drugs against *Mycobacterium tuberculosis* had been discovered, and the QcrB subunit of the cytochrome bc₁-aa₃ complex had been identified as its target from spontaneous mutant strains. STD-NMR experiments were performed on live bacteria cells, expressing either the wild type or the resistant mutant of the QcrB subunit, exposed to one of these new drugs: i) STD signals from the drug were observed only in the presence of the wild-type QcrB, therefore confirming the identity of the drug target; ii) the relative intensities of the STD signals gave clues about the chemical moieties that were the most engaged in binding pockets. This method is perfectly complementary to the chemical and genetic approaches that led to discovery of the drug series, and thus sets a stronger basis for the future drug developments.

### 3. In-cell NMR theoretical and practical considerations

#### 3.1. A historical aside: the beginnings of NMR of biological material

In this section, we sketch a brief historical summary of early NMR work on biological tissues and cells. Many initial misinterpretations were resolved only later as the subject progressed. We have included a discussion of these here, in part because we feel they have historical interest, but also because some of the incorrect theories described below were still cited and re-employed by non-specialists after they had been disproved.

##### 3.1.1. *Early controversies in NMR studies of water (1950-70's), and what can be learned from them*

As early as 1950, Shaw, Palmer and Elsken carried out NMR studies on water in biological material such as apple, potato and maple wood to establish a method for water content quantification [1495]. The NMR signal was measured by low resolution nuclear magnetic absorption at that time, and the authors soon discovered that the correlation between measured intensities and water content was not perfect [1496]. They realized for instance that “soluble solids” such as carbohydrates, proteins or fatty acids were contributing to the NMR signal from apple tissues or in milk, whereas starch in potato tissues made negligible contributions [1497-1500]. Hence, to be accurate, NMR quantification of water required a prior estimation of these soluble species. They noticed however that these solutes were causing broader linewidths for the observed signal, and suggested that characterizing these linewidths might provide a method to investigate the interaction between water and the non-aqueous species [1495-1497]. They also pointed out that T1 relaxation became faster as the viscosity increased in sucrose and gelatin solutions [1499], in agreement with earlier results from Bloembergen, Purcell and Pound [1501]. Faster T1 relaxation had also been reported in various rabbit tissues and in yeast cells by Odeblad and Lindström in 1955 [1502]. These T1 rates would later be connected explicitly to molecular correlation times and the nature of the surrounding spin lattice. Odeblad and Lindström also quantified the water exchange rate in red blood cells by measuring their remaining water ¹H signal after incubation in D₂O. The authors acknowledged however the progressive appearance of deoxygenated hemoglobin, whose paramagnetism would cause faster relaxation times and lower apparent signal strength, as shown earlier independently by Bloch,
Bloembergen et al., and Odeblad et al. [1503-1505]. Hence, in 1956, these first studies had already established the observables and the parameters that would be scrutinized in cellular studies in the decades that followed: signal intensity, linewidth and relaxation times could be related to molecular composition, chemical exchange, viscosity, chemical environment and the effects of paramagnetic species.

The next two decades of NMR research on water in living samples were nevertheless controversial, at least in part because all of these parameters can influence all of these NMR observables. Different interpretations as to what was the dominant parameter causing intensities to be missing could easily emerge and be argued. Hence, the field became involved in two major controversies. The first one has its origin in a model, that in retrospect seems surprising, from Gilbert Ling, who disputed the existence of cellular cation pumps and their function in regulating transmembrane cation concentration gradients: these would have cost too much energy to the cell in his view. Instead, he had proposed since the 1950's an "association-induction hypothesis" leading to the model of a "polarized-oriented multilayer" of structured cellular water and Na⁺/K⁺ ions in cells [1506-1508]. This sounds strange now that cation pumps are well-established, that venom and drugs targeting them are known, and that several Nobel prizes have been awarded to researchers in the field. In fact, it did already sound awkward to a number of contemporaries, but during the 1960's some experimental results were published that were consistent with this controversial theory [74,1509-1515]. These papers reported line broadening and relaxation rates of the water NMR signal in animal tissues, notably upon muscle or nerve excitation. The results were interpreted in the framework of a two-phase model, with one minor "liquid" population and one major "solid" or "ice-like state" population of water molecules. These interpretations were also supported by other NMR experiments on Na⁺ and K⁺, but, as discussed in a later section, these latter experiments proved to have an incorrect interpretation of "invisible" signal. Among others, Cope, Hazlewood and Damadian became involved in the debate [1516], which became more complex once Damadian and others reported variations of T1 relaxation rates in tumors and in normal tissues in the 1970's, again in highly influential journals [1517-1520]. Damadian adopted the model of Ling [1517,1521], which provided a framework to describe the loss of water structure as a hallmark of cancer cells, and his findings led him to promote NMR as a method to detect tumors. While his support of the model of Ling proved to be mistaken, Damadian's recognition of the possibility of using relaxation properties to distinguish tumors from healthy tissue encouraged the development of MRI (see [1522] for review). Another positive outcome was that the controversies, which were hotly debated (see [1521]), pushed the community to propose better explanations during the 1970's. To summarize these results briefly, with hindsight the incorrect interpretations proved to arise from i) an inability to detect the early, rapidly relaxing components of NMR signals, which in turn resulted from using materials and equipment that required delays between excitation and acquisition, and/or ii) the use of imperfect models [197,272].

Indeed, physicists working in the field determined several possible mechanisms that would affect T1 and T2 relaxation rates of water. Water T1 relaxation rates in cells and tissues were found to be correlated with water content [1523-1526], which varies, for example, between different organs. Thus, water T1 and T2 appeared to be an ambiguous parameter for diagnosis in the mid-1970's (a good review appears in [1526]). Intracellular T2 relaxation could be interpreted in terms of multiexponential regimes, and accelerated notably i) by chemical exchange between bulk water and water molecules in the hydration layer of macromolecules, ii) by the diffusion of water molecules in magnetic field

gradients generated by heterogeneous magnetic susceptibility, iii) by the exchange between intracellular and extracellular media, iv) by the progressive appearance of paramagnetic species due to hemoglobin deoxygenation and the release of reactive oxygen species in tissue samples [73,1527-1534]. The physics of these interactions was not straightforward, and the quantification of their effects remains an active subject for study, notably to achieve a good understanding of Blood Oxygenation Level Dependent (BOLD) MRI (see Section 3.2. and the following references [79,1535-1537]). Fortunately, the introduction of injectable paramagnetic agents contributed substantially to the success of MRI, by strongly affecting water NMR relaxation rates and thus generating simpler T1- or T2-weighted imaging contrast.

### 3.1.2. The case of quadrupolar nuclei

Quadrupolar nuclei require careful manipulation during NMR experiments. Their fast relaxation is accelerated by dense and heterogeneous cellular environments. We will not discuss the theoretical background to this, which would be well beyond the scope of this review (see for instance [263,1538,1539]). Instead, we present this as a historical discussion, complementary to the previous subsection on water.

We start by discussing the impact that ³⁹K NMR studies made on the understanding of intracellular K⁺ dynamics. Similarly to what happened with ²³Na, ³⁹K NMR spectroscopy started with the observation by Cope and Damadian in 1970 of limited "NMR-visibility" of the intracellular pool of ³⁹K⁺: only 30-40% of the expected signal could be observed in cells by NMR [269]. In the late 1960's, a fundamental debate on the nature of the cellular solvent took place: this low signal was interpreted as an argument in favor of limited ³⁹K dynamics in cells, and thus in favor of the existence of an organized, semi-crystalline intracellular solvent. The tenacity (see [1521]) of its champions had long-lasting effects on the debate in the literature. This model was however not compatible with diffusion and electrical conductivity measurements [270]. This motivated some NMR spectroscopists to try to better understand and characterize ionic dynamics, which in turn implies trying to better understand and characterize quadrupolar relaxation, since these ions are quadrupolar. As a result, ³⁹K NMR signal intensity and relaxation were thus quantified in multiple systems and correlated with the relaxation theory of quadrupolar nuclei. ³⁹K studies showed that K⁺ was freely mobile in vacuoles of pea stem, which are very aqueous in character [193,194]; low percentages of immobilized K⁺ were estimated in frog muscle [271] and halobacteria [1540]; 100% ³⁹K signal, i.e. 100% free K⁺, was observed in duck erythrocytes, which contain nuclei and mitochondria, and in mammalian erythrocytes, which do not [218].

Theoretical frameworks had been provided as early as 1970, and through the 1970's by multiple groups: in a cellular environment where ²³Na⁺ and ³⁹K⁺ interact even very weakly with macromolecular assemblies, their spin 3/2 and quadrupolar nature would result in the appearance of a fast-relaxing component affecting 60% of the total expected NMR signal [196,270,1541-1545]. This fast-relaxing signal could be detected only with advanced equipment. Consistent with this, Shporer and Civan had shown experimentally from 1972 that ²³Na⁺ and ³⁹K⁺ showed multiple T2 relaxation components in non-homogeneous samples where these ions could interact transiently, including frog muscles and halobacteria [195,196,271,1540]. Although this framework became consistent and had good predictive power [202,336,1546], the controversy continued during the 1980's, and different groups regularly reported low ³⁹K⁺ intensities equivalent to ~40% of the expected signal in yeast [200], in rat hearts [225,1547], and in E. coli [276]. The experimental proofs were broadly agreed only in the 1990's, showing that the observation of the fast and slow components of the intracellular ³⁹K signal required a number of advances in
<figure><img src="image_28.png" /><figcaption>**Fig. 24.** Schematic representation of an erythrocyte in a B₀ magnetic field, and of the phenomena influencing NMR relaxation of water. The gray lines indicate the magnetic field lines generated by the difference between the magnetic susceptibilities of the intra- and extra-cellular media. These magnetic field gradients are intrinsically lower inside the cells, but cause basal peak broadening. Signal linewidth is also affected by faster relaxation due to i) diffusion of water molecules through magnetic field gradients (the resonance frequency of every molecule varies during acquisition, yielding coherence losses), ii) interactions with diamagnetic proteins (albumin or oxygenated hemoglobin), iii) proximity with paramagnetic compounds (like deoxy-hemoglobin), and iv) exchange between sample compartments (intra- and extra-cellular). All these phenomena can affect every molecule observable by NMR in cellular samples. Adapted from Li & van Zijl 2020 NMR Biomed [1536].</figcaption></figure>

equipment: advanced probes and sensitivity, magnetic field homogeneity, faster electronics to record the NMR signal with the shortest delays after pulses and improved baseline correction so as not to cancel the broad, fast relaxing component of ³⁹K even in the presence of significant baseline roll [272,273]. The different mitochondrial content in tissues, and later the ribosome content in *E. coli* [1548] were established as important contributors to ³⁹K interactions and thus of cellular ³⁹K NMR relaxation.

An interesting note: in cells, both T1 and T2 are bi-exponential, having a fast-relaxing component, while the extracellular ³⁹K is likely to show a mono-exponential slow-relaxing relaxation in absence of transient interactions with macromolecules [278,280,1549]. When this slow component of T1 is known, the extracellular signal can be removed using an inversion-recovery block of the appropriate duration (~-ln(0.5*T1)). The fast-relaxing component can then be selectively excited and observed as has been shown in some ³⁹K studies [278,280,1549].

## 3.2. Line-broadening and observability

NMR signals are broadened in living samples. It is worthwhile detailing the sources of this line-broadening if we want to understand how to counteract them. We discuss the main contributions below, i.e. viscosity effects, heterogeneous magnetic susceptibility, compartmental exchange, and multiple interactions (see Fig. 24). These take place in spatial dimensions in the range of 1 to 20 micrometers for prokaryotes to mammalian cells, respectively. For each, the subcellular compartments are at least 10 times smaller. The most studied system has been erythrocytes (red blood cells), which can be schematized as disks of 4 µm radius and 2

µm thickness. Mammalian erythrocytes are enucleated, while avian erythrocytes do have a nucleus and functional mitochondria, which has been useful for comparisons in NMR studies. Larger cells exist, such as frog oocytes that are about 1 millimeter across and also have internal subcellular compartments of exceptional sizes, up to hundreds of micrometers.

In this section, we will try to provide helpful numbers and concepts to understand qualitatively the driving forces of NMR line-broadening in samples containing cells. This is necessary for understanding the current limits of the technique, and thus predicting windows of opportunities in terms of applications, as well as imagining solutions to overcome these limitations.

### 3.2.1. Intra- and extra-cellular mobilities of molecules

Before evaluating the NMR-related phenomena that affect line-broadening in samples containing living cells, we have to determine an important driving factor, namely intra- and extra-cellular apparent viscosities. This is of course more a matter of concern for solution NMR than solid-state NMR. Another difficulty is to determine the molecular diffusion regimes, at the NMR T2 time-scale, i.e. in the timeframe between 1 to 10 milliseconds: are we dealing with free diffusion or restricted molecular motion? In this paragraph, we consider the ideal situation where i) only intermolecular steric repulsion affects intra- and extra-cellular molecular mobilities, and ii) translocation is negligible in comparison to intra- and extra-cellular diffusion. The average membrane permeability for small compounds is indeed low [1550] (Fig. 25). Our assumption is thus somehow realistic in most cases, although it suffers plenty of exceptions: the translocation of various small molecular species is of course accelerated by many dedicated membrane channel proteins.

Water self-diffusion coefficients are about 1.5x10⁻⁹ and 3x10⁻⁹ m².s⁻¹ at 283 K and 310 K, respectively [1551], while diffusion coefficients of small metabolites in pure water are of the same order of magnitude, e.g. 1.05, 0.91 and 0.67 x10⁻⁹ m².s⁻¹ at 298 K for glycine, alanine and glucose, respectively [1552]. If we neglect the fraction of water molecules and metabolites that solvate or interact with macromolecules, small molecules (<1 nm diameter) experience intracellular viscosities about 1.5 times that of pure water [1553]. Hence, in one millisecond the Brownian motion of water and glucose would lead to a three-dimensional mean-square displacement of about 3.5 and 2 micrometers, respectively (11 and 6 µm after 10 ms). We can thus consider that small molecules undergo restricted motions in bacteria, and in mammalian cells too, if we take into account the meshwork of subcellular organelles. It is less evident for the extracellular medium, depending on cell density: a packed cell pellet will exert a certain restriction of the mobility of small molecules, whereas cell at densities below ~5% v/v may not.

Concerning larger molecules such as proteins, intracellular viscosity depends on molecular weight, but also on the diffusion time scale. Regular size proteins (hydrodynamic radius < 5 nm or molecular weight below 200 kDa) experience local viscosities in mammalian cells of about 1.5 to 5 times that in water, and in *E. coli* about 8 to 30 times that in water [1553]. To give some orders of magnitudes, GFP₁, GFP₃ and GFP₅ (28, 84 and 140 kDa) have immediate diffusion coefficients of 0.09, 0.05 and 0.0035 x10⁻⁹ m².s⁻¹ in water, and 0.07, 0.03 and 0.015 x10⁻⁹ m².s⁻¹ in cultured human cells, respectively [1554]. These coefficients hold true for a few milliseconds, but, on a timescale of about 100 ms, decrease to 0.03, 0.01 and 0.005 x10⁻⁹ m².s⁻¹ in the same cells. Intracellular obstacles such as organelles, cytoskeleton and macromolecules are responsible for such anomalous diffusion. Hence, GFP₁ following a Brownian motion in mammalian cultured cells will show a mean square displacement of about 0.6 µm in 1 ms. In *E. coli*, the same GFP₁ can move about 0.3 µm in 1 ms. The cytosol of *E. coli*
<figure><img src="image_29.png" /><figcaption>**Fig. 25.** Indicative, average membrane permeability ($P_m$, vertical axis) of small substances (listed according to their molecular weight and electric charge) for model lipid bilayers. Straight lines link the substances to their $P_m$. $P_m$, corresponds to the membrane permeability in a model of passive membrane diffusion where $J = -P_m \times \Delta c$, with $J$ the transmembrane flux density, and $\Delta c$ the transmembrane concentration gradient. Adapted from Hannesschlaeger et al. Chem Rev 2019 [1550].</figcaption></figure>

also has an equal impact on non-folded, disordered proteins. Using NMR, Waudby et al. reported interesting numbers for non-folded α-synuclein (radius of gyration ~4 nm [1555]) recombinantly expressed in *E. coli* cells: α-synuclein motion is strongly restricted in the millisecond timescale, its root mean square displacement being constant at about 0.35 µm for any evolution time between 10 and 20 ms, close to the 0.5 µm in width of an *E. coli* cell [104].

Local nanoscale viscosities are similar among cultured mammalian cell lines [1556] and through the cell cycle [1557], except of course for proteins engaged in complexes that vary with the phases of the cell cycle [1558]. Local viscosities are similar in the cytosol and in the nucleus for proteins of average size [1559,1560]. However, important differences have been measured recently at the sub-micrometer scale: a single protein of 28 kDa can show diffusion coefficients of $0.025 \times 10^{-9} \text{ m}^2 \cdot \text{s}^{-1}$ in the cytosol except in the few hundreds of nanometers close to actin cytoskeleton bundles, where its diffusion drops by a factor two; moreover, its diffusion coefficients in the nucleus were measured to be about 0.02 and $0.007 \times 10^{-9} \text{ m}^2 \cdot \text{s}^{-1}$ outside and inside nucleoli, respectively [1561]. The same study revealed an intriguing feature: by adding positive net charges to the tracked fluorescent protein, diffusion coefficients dropped progressively to zero. Similar observations have been reported in *E. coli*, where protein-protein and protein-lipid but not protein-nucleic acid interactions play significant roles [1562]. As for small molecules, the apparent extracellular viscosity for proteins is not much affected by the presence of serum or even by the extracellular matrix (ECM) in the case of mammalian multicellular spheroids and tumors: it has been measured recently to be 1.2 to 2 times larger than in pure water for proteins below 200 kDa, depending on the cell line [1563]. However, in this last case, the ECM shows typical correlation lengths of about 20-40 nm that to some extent restrict extracellular protein diffusion.

Altogether, the intracellular and extracellular environment of mammalian cells exert a 1- to 5-fold increase in apparent local viscosity for both small and large molecules. Bacterial cells have more impact on macromolecules, up to 20-30 fold. These reported local viscosities have roughly proportional consequences on translational and rotational diffusion (anomalous effects can appear at

the micrometer scale for translational diffusion, and the effects on rotational diffusion are thus theoretically always a bit smaller than the local viscosity measured by translational diffusion) [1553,1564]. We can translate these increases in viscosity into correlation times or apparent molecular weights: the latter would be proportional to viscosity, using the Stokes law and homogeneous spheres as a model for proteins [1565]. Hence, the experimental, apparent viscosities can already be helpful for the NMR spectroscopist: as far as solution NMR is concerned, rapid calculations show that, based on predicted correlation times, folded proteins of over 25 kDa will always be difficult to observe in *E. coli*, while those below 50 kDa in mammalian cells may be within range for producing useful spectra. Once more, this corresponds to the theoretical case of molecules that do not interact with any cellular component.

Finally, organelles and cytoskeleton establish a sub-micrometer meshwork, where small molecules as well as proteins experience somehow an intracellular restricted motion on the NMR T2 timescale of one millisecond. Extracellular metabolites also experience restricted motion in cell pellets. This will be of importance for the next section.

### 3.2.2. Inhomogeneous magnetic susceptibility

Living material is organized into spatial structures, which causes sample inhomogeneity. This has consequences at different levels, the first being in terms of local magnetic susceptibilities $\chi_{\text{loc}}$. Chemical shifts are proportional to the local magnetic susceptibility $B_{\text{loc}} \propto \mu_0(1+\chi_{\text{loc}})H_0$ (where $H_0$ the spectrometer magnetic field strength), provided we consider the ideal situation where a molecule does not diffuse during the NMR measurement. Magnetic susceptibilities of cells and subcellular components have been reported, but information about this is relatively scarce in the literature, to the best of our knowledge. We have gathered the experimental numbers that we could find in Table 1.

The most studied cells are erythrocytes (red blood cells), which are mostly filled with hemoglobin at about 5 mM. Hence, erythrocytes show magnetic susceptibility variations due to dia- and para-magnetism of oxygenated and deoxygenated hemoglobin, respectively. Their contribution to NMR signal broadening is difficult to disentangle from those due to intracellular interactions and the chemical environment. The only clear reports -to the best of our knowledge- used variable angle spinning to separate them: magnetic susceptibility differences between the buffer and the cytosol of CO-treated (to generate stable paramagnetic hemoglobin), pelleted erythrocytes (mammalian, i.e. enucleated) generate chemical shift differences of 0.1-0.2 ppm (Fig. 26) [1566-1568] that vary in a linear fashion with the intracellular hemoglobin concentration.

When they do not contain any paramagnetic center, proteins seem to have a common average magnetic susceptibility, which is about 1 ppm more diamagnetic than that of water [1569,1570]. Fatty acids are about 1 ppm less diamagnetic than water, and we can expect similar values for tri-acylglycerides present in intracellular lipid droplets. Cholesterol is also found in lipid droplets, and in contrast is more diamagnetic than water. Among phospholipids, we found experimental information only for dipalmitoylphosphatidylcholine (DPPC), for which susceptibility values surprisingly different from those of fatty acids were reported [1571]. Phospholipids are mainly present in mono- or bilayers such as those in bicelles, whose anisotropy of magnetic susceptibility is on the order of magnitude of 0.5 ppm [1571-1574]. The information on DNA magnetic susceptibility may be interpreted with caution: it appears to depend on the degree of solvation and on the interactions with metals or ammonium groups [1575-1579], to comprise dia- and para-magnetic components over multiple scales and thus to vary with the magnetic field [1580]. In our view, the intracellular magnetic susceptibility of
the cell nucleus is thus quite uncertain. Its measurement has been recently reported by Tao and colleagues in a study that resulted in surprising values, also revealing the cytosol to be paramagnetic [1581]. The authors argue that the cytosol hosts organelles that generate paramagnetic reactive oxygen species such as peroxysomes and mitochondria (whose integrity should probably be verified). These results should be carefully considered: the presence of paramagnetic metal ions in cell culture media and cytosol has been regularly reported by EPR [1582].

Overall, cells and their culture medium represent inhomogeneous samples containing compartments, whose intrinsic magnetic susceptibilities vary in the range of 1 ppm. There is no direct correspondence between local magnetic susceptibility and local chemical shifts: the local magnetic fields are continuous functions of space, and the local susceptibilities have non-negligible impacts on magnetic field gradients on a micrometer scale.

Every local magnetic field generates a local chemical shift. In the case of living samples, the sum of all the local environments generates chemical shift distributions. It is however not straightforward to predict local susceptibility values. Magnetic susceptibility variations cause perturbations of the local magnetic field and Larmor frequency, which is a continuous function of space. Hence, calculating the local chemical shifts in heterogenous samples has long been a matter of concern. Using a model of theoretical round avian erythrocyte cells ($\chi_{\text{nucleus}}=-8.9 \times 10^{-6}$, $\chi_{\text{cytoplasm}}=-8.86 \times 10^{-6}$, $\chi_{\text{medium}}=-8.82 \times 10^{-6}$), Kuchel and colleagues calculated that magnetic field gradients could reach about 2 G/cm (at a spectrometer field of 9.4 T) in the micrometer surrounding the nuclear and plasma membrane, i.e. about 1-2 Hz between molecules at the surface of the nucleus and far away in the cytosol [1594,1595]. The consequences were about 20 times larger for erythrocytes that would contain paramagnetic deoxyhemoglobin ($\chi_{\text{nucleus}}=-8.9 \times 10^{-6}$, $\chi_{\text{cytoplasm}}=-7.9 \times 10^{-6}$, $\chi_{\text{medium}}=-8.82 \times 10^{-6}$). Later calculations by Gillis et al. gave about 2-3-fold larger values using more realistic shapes for mammalian paramagnetic erythrocytes (Fig. 27) [1596]. Based on experimental reports, Jensen and Chandra calculated root mean square magnitudes of field inhomogeneity of 0.5 ppm in erythrocyte suspensions and 0.2 ppm in grey matter [1597]. These field variations clearly depend on cell geometry and are half as large for spherical cells [1596,1598]. Given the sus-

ceptibility values reported from cells or cell extracts (Table 1), the chemical shift distribution in the cytosol of a round cell is expected to be similar to that in deoxy-erythrocytes, i.e. 0.1-0.2 ppm. This corresponds to the chemical shift distribution due to the presence of nucleus, cytosol and extracellular medium that have susceptibility differences of about 1 ppm.

The variations of local magnetic fields are intrinsically larger outside of the considered compartment, e.g. the nucleus, the internal organelles or the cell. However, the local fields are a few percent (Fig. 27). For large magnetic susceptibility differences between intra- and extra-cellular media (e.g. for paramagnetic deoxygenated erythrocytes), the resulting heterogeneity in magnetic susceptibility can grow until a certain cell density and then progressively decrease for packed cell samples [1536,1599]. Altogether, we can consider that intra- and extra-cellular molecules are equally affected by the resulting chemical shift variations of 0.1-0.2 ppm.

At a more local level, we can also take into account the magnetic susceptibility differences imposed by the sharp transition between aqueous intra- and extra-cellular media and lipids in membranes or droplets. Using a simplistic model of a magnetic dipole moment and its associated magnetic field imposed by a spherical cell, Wind and Hu calculated that chemical shift distributions of lipids (or other molecules) in membranes may be as much as 0.6 ppm due to their variable orientations in the field and the local differences in magnetic susceptibility [636]. Lipid $^1\text{H}$ line-widths in cellular samples are usually 0.1-0.2 ppm. A large fraction of these lipids is contained in lipid droplets, which are approximately round objects: spheres generate a homogeneous internal magnetic field, but also local inhomogeneities at their external surface. Frog oocytes show large experimental lipid linewidths of about 0.3 ppm, which may be due to their large asymmetric protein/lipoprotein (yolk) or lipid compartments, but also to the adsorbed microscopic air bubbles that are commonly found on the oocyte surface.

In addition, we can consider the chemical shift averaging that molecular diffusion would cause, which may act to reduce the apparent chemical shift distributions described above. We have seen that metabolites diffuse at rates that would allow them to move micrometers per millisecond, i.e. more than a whole E. coli cell length, or a tenth of a mammalian cultured cell. Proteins move

<table><caption>**Table 1**

Volume magnetic susceptibility at room temperature (SI, dimensionless; divide by $4\pi$ for converting to the CGS system); d: density.</caption><thead><tr><th>Material</th><th>χ·10<sup>6</sup> (SI units, dimensionless)</th><th>references</th><th>Material</th><th>χ·10<sup>6</sup> (SI units, dimensionless)</th><th>Reference</th></tr></thead><tbody><tr><td>H<sub>2</sub>O</td><td>-9.05</td><td>[1583]</td><td>Deoxygenated erythrocytes</td><td>-6.1</td><td>[1584]</td></tr><tr><td>D<sub>2</sub>O</td><td>-8.8 (-9.0 in [1585])</td><td>[1552]</td><td>CO-treated RBC (intracellular diamagnetic hemoglobin ~5 mM ~350 g/L)</td><td>-9.25/-9.4</td><td>[1566,1584]</td></tr><tr><td>H<sub>2</sub>O+[NaCl]=150 mM (d=1.05 g/mL)</td><td>-8.95</td><td>Calculated based on [1586]</td><td>Hela 12 to 20 mm diameter</td><td>-6.45 to -6.5</td><td>[1587]</td></tr><tr><td>culture medium (mammalian cells, classical medium 199 [1588])</td><td>-7.5</td><td>[1587]</td><td>lymphocyte</td><td>-9.1</td><td>[1587]</td></tr><tr><td>"Pure protein"(d=1.38 g/mL)</td><td>-10.5</td><td>[1569,1570]</td><td>Frog oocyte</td><td>-8.7</td><td>[1589]</td></tr><tr><td>100 g/L BSA in water</td><td>-9.1</td><td>Calculated based on [1570]</td><td>Pellet/yolk (proteins and lipoproteins)</td><td>-8.0</td><td>[1589]</td></tr><tr><td>DNA (d=1.7 g/mL)χ(diamagnetic)</td><td>-12.8</td><td>[1575,1590]</td><td>Cytosolic</td><td>-8.8</td><td>[1589]</td></tr><tr><td>Fatty acids (d=0.85 g/mL)</td><td>-8.4</td><td>[1552,1591,1592]</td><td>Lipid fraction</td><td>-8.1</td><td>[1589]</td></tr><tr><td>Cholesterol</td><td>-9.7</td><td>[1552]</td><td>Phospholipids (DPPC) (d=0.94 g/mL)</td><td>-9.4</td><td>[1571]</td></tr><tr><td>Glycerol</td><td>-9.8</td><td>[1552]</td><td>Glucose</td><td>-10.9</td><td>[1552]</td></tr><tr><td>Oxygenated hemoglobin (diamagnetic)</td><td>-9.2</td><td>[1584,1587]</td><td>Starch granules</td><td>-10.9</td><td>[1593]</td></tr><tr><td>Deoxygenated hemoglobin (paramagnetic)</td><td>-5.8 to -6.7</td><td>[1537,1584]</td><td>CNE-2Z carcinoma cell line human (for d=1 g/mL)</td><td>Nucleus: -6.8 Cytosol: 9.9</td><td>[1581] Should be interpreted cautiously, in regards to all other reported values</td></tr></tbody></table>
<figure><img src="image_30.png" /><figcaption>**Fig. 26.** One-dimensional ¹H NMR spectra of an erythrocyte suspension in a rotor spinning at varying angles θ with respect to the spectrometer field: close up views of the water peaks: **A)** Magic-Angle Spinning, which cancels the effects of varying magnetic susceptibilities in the sample, **B)** 0° spinning and **C)** 90° spinning. The labels "Extra." and "Intra." indicate the peaks from water molecules in the supernatant and inside the cells, respectively. The spectrum at the magic-angle reveals the isotropic chemical shifts due to the different chemical environments inside and outside the cells. The difference of magnetic susceptibility between inside and outside the cells is given by $Δχ=2(Δν₀ - Δν₀)/Δν₀$, with $Δν₀$ and $Δν₀$ the intervals in Hz between the "Intra." and "Extra." peaks when spinning at 0° and 90° with respect to the spectrometer field. Here, $Δχ$ was measured to be $-2.18×10⁻⁷$ (SI units, dimensionless). The rotor was spun at 250 Hz, spectra were recorded at 9.4 T (400 MHz). Adapted from Kuchel et al. Prog NMR Spec 2018 [1566].</figcaption></figure>

<figure><img src="image_31.png" /><figcaption>**Fig. 27.** Schematic representations of **A)** the relative magnetic field variation induced by $Δχ$, the difference of magnetic susceptibility inside and outside an idealized mammalian erythrocyte (4 µm of diameter); if $Δχ = 1$ ppm, molecules in contact with the erythrocyte membrane can resonate 0.5 ppm away from their remote counterparts (adapted from Gillis et al. 1995 [1596]); **B)** a sample of cells in suspensions (white) with a volume fraction of 0.46, whose intracellular magnetic susceptibility (white) is different from the extracellular medium (black), and **C)** the corresponding magnetic fields (or Larmor frequencies) induced by a field applied vertically, represented using shades of grey. Every intracellular magnetic field is influenced by the presence of neighboring cells and magnetic susceptibility inhomogeneities. Adapted from Novikov and Kiselev 2008 J Magn Reson [1600].</figcaption></figure>

about 0.1-0.5 micrometer per millisecond, which is enough to explore intracellular spaces comprising multiple subcellular compartments. Lipids are on average much less mobile and are thus not considered here. Let us first focus on chemical shifts. Recently, Kiselev and colleagues calculated the frequency shifts of molecules freely diffusing in a medium containing spheroid inclusions: whatever the diffusion rates and cell shapes, chemical shift variations are predicted to be limited to a few hundredths of the difference between magnetic susceptibilities of the solvent and of the inclusions [1601,1602]. We can model cells in their surrounding medium, as well as organelles in the cytosol, in the same way as such diamagnetic inclusions. Hence, as compared to chemical shifts of purified molecules measured in aqueous buffers, we expect chemical shift changes of about 0.1 ppm for both intra- and extracellular molecules in samples containing cells. Again, this discussion applies for a model of extracellular molecules that do not interact with cells, or of intracellular molecules that do not interact with organelle membranes.

As a final point concerning the effects of magnetic susceptibility on NMR line-broadening in cell samples, we must discuss an additional effect that is associated with diffusion in an inhomogeneous field. In the presence of magnetic field gradients, diffusion generates a specific increase in the rate of loss of transverse magnetiza-

tion [68]. This effect has been fruitfully used for about 50 years to measure diffusion rates by solution NMR spectroscopy. In the context of cellular samples, we face not only the difficulty that magnitudes of the local gradient fields are unknown, but so are the diffusion coefficients and diffusion regimes of the observed molecules. In the case of free diffusion in a steady and linear magnetic field gradient G, the signals attenuate in time (t) proportionally to $exp(-γ²D²G²t³/12)$, where γ is the gyromagnetic ratio and D is the diffusion coefficient [71]. We have discussed in 3.2.1 the fact that molecular diffusion can be considered to be restricted in the millisecond timescale. This leads to signal attenuation regimes that can be very different from those observed for free diffusion in steady magnetic fields. There is a wealth of literature on NMR measurement of diffusion in heterogeneous media, and the number and sophistication of the physical models in use makes it impossible to present them here in a few lines [69,71,1535,1597,1603-1609]. Rather, we can just keep in mind that diffusion-related transverse signal loss is very dependent on diffusion coefficients and the length and shape of the relevant compartment. Indeed, we did not succeed in finding simple analytical equations in the literature that could be used as simple guides for transverse signal loss in cell samples. The proposed attenuation regimes are quite complex, depending on the starting assumptions, and we would
only be able to provide a long list of possible situations together with impractical equations containing large exponents for every term, given the heterogeneity of magnetic susceptibilities and shapes of cells and subcellular compartments, as well as the variability of cell packing and molecular diffusion coefficients. A phenomenon that is close to this topic is water relaxation in blood, which has been much studied using CMPG sequences and diffusion NMR experiments. The measured diffusion-related ¹H relaxation due to susceptibility heterogeneity is dominant from 1.5 T upwards, proportional to $\Delta\chi^2$, $\gamma^2$ and $B_0^2$, and shows dispersion when varying the inter-echo spacing in CPMG experiments because of various exchange mechanisms [933,1536,1584,1596,1597,1599,1603,1610-1612]. Interestingly, if we focus on signal broadening, the diffusion of water molecules in the cellular field gradients has a stronger impact on attenuation of the FID than on CPMG relaxation [1604,1606,1613,1614]. As an extreme example, at a spectrometer field of 300 MHz, the specific FID R2 (1/T2) of intracellular water due to diffusion ranges from ~60 Hz to ~700 Hz for paramagnetic erythrocytes at ~100% and 40% hematocrit (volume percentage of the pelleted cells), respectively [1599]. Fully oxygenated, diamagnetic erythrocytes show diffusion-related water line-broadening below 0.1 ppm. These numbers will hold true for small molecules. As far as macromolecules are concerned, we can mention the fact that, up to 1.2 GHz, $\alpha$-synuclein linewidth did not increase with the square of the magnetic field in cultured mammalian cells [1615]. This may show that diffusion in cellular samples makes only weak contributions to protein relaxation. Finally, the effects of compartmental exchange between intra- and extra-cellular media are about 5-fold lower for water relaxation in erythrocytes, where water has a lifetime of about 10 ms [1536]. We can expect the effects to be smaller for most metabolites because of their slower exchange kinetics through the plasma membrane.

As a last, speculative point: we should also consider membrane-less organelles produced by liquid-liquid phase separation, which can contain high concentrations of proteins and/or nucleic acids [1616,1617]. These may also be treated as examples of diamagnetic inclusions, although this possibility has not been thoroughly discussed in the literature to the best of our knowledge [1618].

To summarize, according to theoretical calculations on local magnetic field susceptibilities, cellular samples can generate chemical shift distributions, whose width is in the order of 0.1-0.2 ppm for a single molecular species localized either inside or outside cells. These distributions can even be asymmetric if cells are not spherical and molecular diffusion is sufficiently slow (Fig. 28). Larger cells such as oocytes and pluricellular organisms that include large non-spherical portions can provoke larger spreads in the 0.3 ppm range.

### 3.2.3. Multiple interactions. A warning concerning quantification and referencing.

Interactions with cellular entities are very unpredictable phenomena for any molecule in a cell, yet they are at the core of the in-cell NMR approach, for which the aim is to find information on what happens in the presence of native partners in a native context. They are also possibly the most damaging for NMR investigations, because they can cause signal disappearance. NMR spectroscopists are used to dealing with this possibility in vitro, where causes and consequences are well-known: in the case of solution NMR they comprise interactions in the intermediate regime (µs-ms) or with large macromolecular assemblies; in the case of solid-state NMR, multiple interactions and multiple chemical environments or inappropriate conformational dynamics at room temperature. We highlight here some considerations of broad interest.

A

<figure><img src="image_32.png" /></figure>

B

Chemical shift distribution

<figure><img src="image_33.png" /></figure>

C

<figure><img src="image_34.png" /></figure>

D

<figure><img src="image_35.png" /></figure>

**Fig. 28.** A) Representation of the magnetic flux induced by ellipsoid cells of homogeneous magnetic susceptibility $\chi_i$ in a medium of susceptibility $\chi_e > \chi_i$ and a horizontal magnetic field $B_0$; B) Because randomly oriented cells more often adopt an orientation perpendicular to $B_0$, the chemical shift distribution of intracellular molecules corresponds to that seen in an NMR powder pattern; C) Illustration of the path of a water molecule during a 60 ms long Monte Carlo simulation in a medium where ellipsoid cells (shown semi-transparent red) are randomly oriented and occupy 38% of the volume; through the course of the simulation parts of the pathway shown in blue are extracellular, while the three green sections correspond to cell penetrations; D) Calculated frequency shift at every step of the Monte Carlo simulation presented above, using realistic parameters corresponding to diamagnetic erythrocytes at 3T mixed with an external medium containing 8 mM of Gd³⁺-contrast agent. Adapted from Wilson et al. 2017 [1619].
An important aspect is the lower availability of water for hydrogen-bonding in cells. Cells contain high concentrations of molecules that establish hydrogen bonds with water, e.g. metabolites, proteins or nucleic acids. Hence, water is on average less available for each of these hydrogen bonding events than it would be in dilute systems containing a single solute. Once the effects of magnetic susceptibility on chemical shifts have been removed (using HR-MAS for example), chemical shift differences between intra- and extra-cellular species are reminiscent of those observed between protic and aprotic solvents. This solvent effect makes an important contribution to the NMR split peak effect observed between intra- and extra-cellular populations of small molecules in cell suspensions. It has been extremely well presented by Kuchel and colleagues in a recent review [1568]. All hydrogen bonding metabolites or drugs are affected, notably ³¹P- or ¹⁹F-bearing molecules.

Turning to consideration of the water, protein solvation has consequences on water chemical shift and relaxation. The contribution of water-protein exchange has been measured to be about 0.2 parts-per-billion (ppb) per g/L of protein [1567,1570,1620,1621]. To calculate the effective water frequency in protein solution, this exchange contribution has to be combined with the effects of protein magnetic susceptibility, which is of the opposite sign and about twice as large. These two contributions are very variable among small molecules, and, unfortunately, standard NMR referencing molecules like DSS, TSP, or MDPA interact with proteins at high concentration, which can affect internal chemical shift and quantification referencing [1570,1620,1622]. Although they do not interact with proteins, other referencing molecules, like dioxane, can be affected by the solvent-induced shift presented above [1620]. Considering metabolite quantification, amino acids and important metabolites, e.g. pyruvate or creatine, also interact with proteins [1413,1414,1623]. These interactions depend naturally on the protein, and the binding kinetics vary from slow to fast exchange in the NMR timescale, giving rise to variable changes in signal intensity and chemical shift.

Proteins also have an impact on water relaxation, which is dominated at high field by chemical exchange between protein exchangeable protons, the water molecules from the hydration shell of proteins, and the bulk water molecules [1536,1612,1624-1626]. This exchange shows a strong dispersion at short CPMG inter-echo spacing, and can be modeled in different ways, but we can keep in mind that albumin and diamagnetic hemoglobin have relaxivities of about 10-15 Hz per millimolar of protein (70 Hz/mM for paramagnetic deoxy-hemoglobin) at 500-700 MHz and 310K. These values are naturally very dependent on molecular weight and the folding status of proteins. Nonetheless, one can still extrapolate rough relaxivity values of about 25 Hz at these fields, yielding a water line-broadening value of about 0.1 ppm. This increases with the square of the magnetic field, and is comparable to line-broadenings due to magnetic susceptibility heterogeneity discussed above. The impact of other macromolecular constituents like lipids [1627] or nucleic acids have been less thoroughly characterized, to the best of our knowledge. However, relaxivity effects of glucose and maltose on water can reach 0.1 Hz/mM at 500 MHz, a number that is probably close to those for other carbohydrates [1628-1631]. Glutamate and glutamine also show 0.1 Hz/mM relaxivities (at 400 MHz) [1632]. Overall, these cellular components probably yield water signal line-broadenings of about 0.2-0.4 ppm at high fields. Consequences in terms of water-suppression, cross-relaxation mechanisms and signal intensities of water-exchangeable protons may be non-negligible.

Concerning intracellular interactions and their consequences for proteins and in-cell NMR, we selected below a few striking observations from the literature. The first measurements of protein tumbling features in cells were reported 45 years ago using

¹³C-histidine labeling of mice (fed with [2-¹³C]-histidine) and their hemoglobin in erythrocytes [1178]. The intracellular correlation time was consistent with an intracellular viscosity only twice as large as that of an aqueous buffer. Twenty years later, using 5-fluorotryptophan incorporation and ¹⁹F-NMR T1/T2 relaxation measurements, Brindle and colleagues reported similar increases of correlation times for two enzymes overexpressed in yeast [1179]. They noticed however that two other enzymes were not observable in intact cells, whereas they recovered their ¹⁹F signals upon cell lysis [1179,1343]. We recall briefly here the strong size-dependency of apparent viscosity experienced by proteins (see Section 3.2.1.). Besides these effects, we meet here the multiple weak interactions that proteins can establish in cells, and their deleterious consequences for NMR signals. The use of ¹⁹F-amino acids was later fruitfully exploited by Pielak, Li and their coworkers: T1 and T2 relaxation were very differently affected by the effective local viscosity and transient interactions with other macromolecules [1344]. Hence, by measuring T1 and T2 rates in water-glycerol solutions, diluted *E. coli* extracts and intact *E. coli* cells, they could show that for ubiquitin and GB1 tumbling was similarly affected by a 2-3-fold change in intracellular viscosity, but that ubiquitin was involved in substantial transient interactions in cells while GB1 was almost not. Consequently, the ¹⁹F-line-widths of intracellular ubiquitin (8.6 kDa) and GB1 (6.2 kDa) corresponded to those of ~200 kDa and ~50 kDa proteins in buffer, respectively. Similar observations had been previously reported by Gierasch and her coworkers, using an ingenious approach: they measured linewidths of both TROSY and anti-TROSY peaks in ¹H-¹⁵N spectra, their difference being determined by local dynamics and not by chemical exchange or magnetic field inhomogeneity [1191]. They could show that about 30% of the intracellular GB1 ¹Hₙ line-width was due to transient interactions. As a consequence, GB1 yielded good quality NMR spectra in cells, while ubiquitin did not. A triple mutation of hydrophobic amino acids at the surface of ubiquitin (L8A-I44A-V70A) sufficiently reduced the unspecific interactions to result in NMR signal recovery in cell lysate, but not in cells. More generally, hydrophobic residues on protein surfaces, a net-positive charge and also a large dipole moment have been described to negatively affect in-cell NMR signal [1345,1349,1633]. Molecules present in high concentrations, such as ribosomes, may play a role in enhancing transverse relaxation of proteins in cells [1351,1354]. Burmann et al. have shown that for α-synuclein in mammalian cells, chaperone silencing reduced transverse relaxation of the hydrophobic patches involved in loose interactions verified in vitro. [1250]. This suggests a more global binding of exposed hydrophobic patches by chaperones. It is well-known by NMR spectroscopists that transient interactions engaging even low populations can cause considerable transverse relaxation. The differential impact of viscosity and transient interactions on ¹H-¹⁵N T1 and T2 relaxations holds true in cells and may carry information on the size distribution of interacting partners, as proposed recently by Danielsson and colleagues [1350]. Based on their characterization of relevant parameters (namely et charge, surface hydrophobicity and dipolar moment), they also used mutations to greatly improve in-cell NMR spectra of their proteins by removing unspecific transient interactions that spontaneously occur in the cellular environment [1349,1350]. This strategy may be fruitful in the future, even though its use should presumably be limited to specific questions that do not require maintenance of the whole set of native interactions; this, in turn, raises the question of how to know in advance whether the whole set of native interactions is to be maintained. A less-questionable strategy consists in avoiding positively charged peptide tags, which have undesirable effects on intracellular tumbling and on the NMR signal [1190,1347,1561,1562].
### 3.3. Observation of ions and metabolites

This section mostly focusses on in-cell studies by solution and HR-MAS NMR spectroscopy. We aim to provide information complementary to that presented in the applications Section 2. Because we will be discussing spin-echoes and T2/T1rho filters, we briefly mention the potential sample heating due to the applied RF.

#### 3.3.1. Metal ions

Many metal ions have been used for direct NMR detection in cells: ⁶Li and ⁷Li (s=3/2 and s=3/2, natural abundance of 7.6 and 92.4 %, respectively), ¹⁰B and ¹¹B (s=3 and s=5/2, n.a. 19.9 and 80.1%, respectively), ²³Na (s=5/2, n.a. 100%), ²⁷Al (s=5/2, n.a. 100%), ³⁹K (s=3/2, n.a. 93.3%), ⁸⁵Rb and ⁸⁷Rb (s=5/2 and s=3/2, n.a. 72.2 and 27.8%), ¹¹³Cd (s=1/2, n.a. 12.2%), ¹³³Cs (s=7/2, n.a. 100%). Unfortunately, for many important metals their only NMR-active isotope has low abundance, which precludes sensitive direct NMR detection: ²⁵Mg (s=5/2 n.a. 10.1%), ⁴³Ca (s=7/2, n.a. 0.14%), ⁵⁷Fe (n.a. 2.1 %), ⁶⁷Zn (s=5/2, n.a. 4.1%), ⁷⁷Se (s=1/2, n.a. 7.6%). Moreover, paramagnetic states prevent direct NMR detection of, e.g., Mn (II), Mn(III), Cu(II), Co(II): ⁵⁵Mn (s=5/2, n.a. 100%), ⁵⁹Co (s=7/2, n.a. 100%), ⁶³Cu and ⁶⁵Cu (s=3/2 and s=3/2, n.a. 69.2 and 30.8%, respectively).

Metal ions often show broad, overlapping signals from outside and inside cells [202]. Membrane-impermeable molecules, so-called “shift reagents”, have thus been developed from the early 1980’s to generate exploitable chemical shift changes for extracellular nuclei. The rationale for this was presented in Section 2.1.2. These “shift reagents” are commonly chelated lanthanides, notably dysprosium or thulium [131,198,199,201,202,225,242,1634-1636], whose paramagnetic properties yield significant frequency shifts and little signal broadening [133]. These reagents have their “signal-broadening only” counterparts in gadolinium-complexes MRI contrast agents, which can effectively shut off NMR signals in their vicinity [1637]. It was soon realized that some of the “shift reagents” could be unstable [216,238,1638], toxic [239,240,1639], could modify Li⁺/Na⁺ transport rates and membrane potential in red blood cells [1640,1641], and were dependent on pH or Ca²⁺ concentrations [1638,1641,1642]. Although not perfect, a more harmless Tm-DOTP complex “shift reagent” has been regularly used later for cellular studies [1643,1644]. Advanced versions of “shift reagents” are still being developed for MRI purposes [130].

Most of these NMR-observable isotopes are quadrupolar, making them more challenging for NMR measurements. Interestingly, increases in their relaxation rates can also reveal intracellular interactions in a sensitive fashion (see Sections 2.1.3., 2.1.4, 2.1.5. and 3.1.2.). Double- and Triple-Quantum Filtered experiments allow attenuation of the signals from extracellular quadrupolar nuclei that do not interact with macromolecular assemblies (see Section 2.1.4.).

#### 3.3.2. Spin-echo as an editing strategy

Various versions of spin-echoes have been used in the past to edit certain resonances from cellular samples, and were important in early NMR studies of metabolites. These are now most often substituted by T2/T1rho filters to purge out signals from fast relaxing species, or by ¹³C/¹⁵N-editing pulse blocks such as INEPT to remove signals from non-labeled species. We earlier described a number of studies using spin-echoes in the 1970’s-1980’s (particularly in Section 2.3.4.), where we described the concept of spin-echo only in brief terms. Here we provide more detailed descriptions.

Spin-echo pulse sequences rely on the following principle: instead of recording ¹H-NMR signals immediately after the application of a 90° pulse, one can wait for a few tens of milliseconds, then refocus the spin magnetization (using a 180° pulse), wait

for the same amount of time and record the remaining “echoing” signals, those that relax slowly enough to have persisted through the delays (Fig. 15). This purges out the resonances of protons of slow tumbling species from the observable pool of signals, like those of proteins, nucleic acids polymers, and fatty acids from lipid droplets and membranes to some extent. Hence, this spin-echo method provokes a loss of information on slow-tumbling species, while it reveals the available information on fast-tumbling metabolites. Modern variants of this method are called T2- or T1rho-filters.

Scalar couplings can evolve during spin-echoes. This has been exploited notably in the case of ¹H-¹H and ¹H-¹³C scalar couplings. Indeed, peak inversion can help to assign peaks in ¹H-detected spectra, to monitor proton-deuterium exchange or ¹³C incorporation, as exemplified in Section 2.3.4. Interestingly, ¹³C metabolic insertion causes the appearance of characteristic splittings in ¹H-peaks due to scalar coupling, which coalesce back at the central resonance if ¹³C-decoupling is applied during acquisition. Hence, it is possible to choose to acquire FIDs after spin echoes that either do or do not allow scalar couplings to evolve, and to combine them by addition or subtraction to select either the ¹³C- or ¹²C-containing species in the final ¹H-detected spectra, while exploiting ¹H sensitivity.

The basic observation of ¹H-peak splitting due to ¹³C incorporation was first exploited in spectra from in living cells by Shulman and his coworkers: by measuring one-dimensional ¹H-NMR spectra with or without ¹³C-decoupling, and subtracting one from the other, they quantified ¹³C-incorporation from [2-¹³C]-acetate into glutamate or aspartate by S. cerevisiae [948] (Fig. 29). A more sophisticated ¹³C-modulated ¹H spin-echo was exploited by Brindle and colleagues: they combined ¹H-coupling and ¹³C-coupling evolution during spin echoes with ¹³C-decoupling during acquisition. Addition and subtraction of the FIDs obtained allowed monitoring of the ¹²C- and ¹³C-methyl content in the separated spectra. They could quantify ¹²C- and ¹³C-labeled alanine and pyruvate in real-time, and thus determine the activity of alanine aminotransferase in erythrocytes [949]. This approach produced spectra in which peak intensities were easier to analyze than those obtained by Shulman previously, by separating the protons bound to ¹³C from those bound to ¹²C nuclei into separate spectra.

There is a clear connection to the more standard non-in-cell NMR applications and the vast possibilities of spin-echo derived strategies. A great number of them have been proposed since the 1960’s, amongst the most famous being probably the INEPT transfer and BIRD filter for editing ¹³C/¹⁵N-bound protons [1043-1047, 1050,1051,1645,1646], and the TANGO filter for editing ¹²C-bound protons [1647,1648]. The field would be impossible to cover entirely and is well beyond our scope. The common INEPT has been the most widely used by the in-cell NMR community, but recent applications of BIRD or TANGO on live material show that these can also yield interesting information [1066,1649].

### 3.3.3. T2, T1-rho filters and other filtering/editing methods for signals from mobile species

As mentioned earlier, spin-echoes are nowadays substituted by T2 or T1-rho filters to relax away the magnetization from slow-tumbling molecules before acquisition, and thus simplify the NMR spectra from complex samples. These filters are notably supposed to suppress scalar coupling evolution of the small molecules of interest, though they do not always do so perfectly. It is recommended to use the improved CPMG scheme proposed by Morris, Bodenhausen and their colleagues, which includes J-refocusing [1650]. Using an adiabatic spinlock offers superior results for the long T2-filters (>400 ms) necessary to remove signals from cellular lipids (they reduce the power deposition and thus sample heating) [1039,1651]. T2-filters can also be combined with diffusion filters
<figure><img src="image_36.png" /><figcaption>**Fig. 29.** Spin-echoes and ¹³C-decoupling allow multiple editing/filtering strategies. A) One dimensional ¹H-NMR spectra or ¹²C- or ¹³C-labeled molecules can be distinguished by the large scalar coupling ¹J(¹H-¹³C), which generates typical doublets. However, ¹H-spectra are often sufficiently crowded to make analysis of both ¹²C- and ¹³C-bound proton signals difficult. B) A spin echo with τ=1/2*|J(¹H-¹H)| will invert signals both from ¹²C- and from ¹³C-bound protons, whereas C) a supplementary 180° pulse on ¹³C will result in positive signals of ¹³C-bound protons if τ=n/|J(¹H-¹³C)|. D) ¹³C-decoupling during ¹H-acquisition does not affect ¹²C-bound protons, but causes peak coalescence of ¹³C-bound proton signals. E) Applying ¹³C-decoupling during acquisition causes overlap and addition of signals from ¹²C- and ¹³C-linked protons, but F) the addition of a 180° pulse on ¹³C modulates the sign of ¹³C-linked protons in the sum. Hence, adding or subtracting spectra E and F permits monitoring of ¹²C- or ¹³C-labeled molecules, respectively.</figcaption></figure>

<figure><img src="image_37.png" /><figcaption>**Fig. 30.** a) The combined use of relaxation and diffusion filtering methods permits selective observation of different pools of molecules in complex samples (here a hemp seed extract): a short relaxation filter leaves high signal intensities from lipids (ii), which can be edited by a diffusion-filter (iii), the subtraction of (iii) from (ii) providing spectra mostly free of lipid signals (iv); b) Long spinlock mixing times are necessary to cancel lipid signals; c) the fast-tumbling species conserve high intensities upon the application of relaxation filters. Adapted from Hassan et al. 2019 [1039].</figcaption></figure>
to select signals from molecules with relatively fast correlation times but slow translational diffusion coefficients (e.g. lipids) (Fig. 30) [1006,1039,1056,1651].

Simpson and colleagues have recently proposed using ²H-detected NMR to detect the incorporation of ²H-labeled nutrients in fresh water shrimp and fleas: the quadrupolar nature of ²H ensures that only very mobile ²H-labeled species are observable, but it presupposes the joint use of low-speed HR-MAS [1038]. They also proposed to adapt typical protein NMR pulse schemes to monitor ¹³C-labeled levels of amino acids in water fleas [1065]: these sequences transfer magnetization through ¹³C-labeled amino-acid side chains and backbone, using ¹³C-selective pulses and long magnetization transfers that filter out non-peptidic and large molecules. They also applied a data-processing approach that acts like a relaxation filter: the window function is shifted to suppress early points of the FID (Fig. 31).

Another strategy has sometimes been to record one-dimensional ¹H-NMR spectra of similar cell samples, e.g. undifferentiated neuroblastoma/glioma cells and their differentiated counterparts. After normalization of the two spectra according to their global profile intensities, the second spectrum was subtracted from the first: NMR signals from metabolites showing varying levels in the two cell lines, like glutamine and glutamate, could be observed in the difference spectrum [1652]. We did not meet this method very often in the literature, even though a similar approach has

been applied recently to remove the cellular background in studies focusing on overexpressed proteins [9,1251,1280].

### 3.3.4. High-Resolution Magic-Angle Spinning (HR-MAS) to alleviate sample heterogeneity: advantages, drawbacks and late developments

Cells cause intrinsic heterogeneity in magnetic susceptibility (see Section 3.2.2.) and limit the mobility of a number of molecules. Even though cellular samples do not behave like solid matter, they partially lose some of the solution character that facilitates solution NMR. This leads to NMR line-broadening because of dipolar fields due to magnetic susceptibility heterogeneities, and restricted motions increase the negative impacts of dipole-dipole interactions, of chemical shift anisotropy, and of first order quadrupole interactions. All these terms show a dependence on the second-order Legendre polynomial, proportional to $3\cos^2(\theta) -1$, with $\theta$ the angle between the field $B_0$ and the interaction vectors or anisotropy tensors. Magic-Angle Spinning has the capacity to average out these deleterious effects if the rotation rate is faster than the strength of the above mentioned phenomena (Fig. 32) [631,641,1653-1657]. MAS in liquid-like and heterogeneous systems was proposed in the 1970's [627-629,1658-1660], and became progressively more popular in the 1990's under the name HR-MAS to study the mobile fraction in liquid-phase of heterogeneous samples including tissues or living cells

<figure><img src="image_38.png" /><figcaption>**Fig. 31.** A shifted sine-squared bell window function can act as a potent relaxation filter. These spectra were obtained from a 2D (H)CbCa(COCa)Ha pulse sequence applied to a ¹³C-enriched *D. magna* extract. Adapted from Lane et al. 2019 [1065].</figcaption></figure>

<figure><img src="image_39.png" /><figcaption>**Fig. 32.** A) Schematic representation of a MAS probe, where the rotor containing the sample is inserted in a MAS stator; B) HR-MAS one-dimensional ¹H NMR spectrum of 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) vesicles in D₂O acquired with MAS and C) without MAS. Adapted from Vermathen et al. 2017 [970].</figcaption></figure>
[630,636,641,974,1024,1068,1661]. Very good guidelines for sample preparation have been published elsewhere [972,1068,1662].

Carbohydrates and lipids represent a class of molecules that can clearly benefit from the line-narrowing effects of HR-MAS. Carbohydrates are notoriously difficult to study because of the limited dispersion of their NMR signals [1663-1666]. Moreover, in living cells, polysaccharides are often found in the neighborhood of cell membranes where magnetic susceptibility effects due to heterogeneity are particularly strong, and molecular mobility can be severely hampered. Typical examples include bacterial lipopolysaccharides or capsular polysaccharides, or N-glycosylation of eukaryotic membrane proteins. Hence, HR-MAS NMR spectroscopy has been instrumental for their NMR characterization in living cells (see Sections 2.2.3. and 2.3.5. for applications). The available carbohydrate databases will certainly facilitate spectral assignment and glycan studies in cells in the near future [634,1667]. Small molecules such as metabolites can also greatly benefit from HR-MAS, and the applications explored in the metabolomics field have been numerous (see Sections 2.4.5. and 2.4.6.).

There are nonetheless some shortcomings of HR-MAS on living samples. First, cells suffer if subjected to hours-long MAS at rates above ~500 Hz, which packs them tightly on the 4 mm diameter rotor walls and possibly damages their viability. The centrifugal forces generated by MAS are clearly problematic: using disposable 4 mm diameter rotors, 1-4 kHz spinning rates translate into forces of 8,000-100,000 g, which rapidly affects the integrity of mammalian cells [67]. These spinning rates are necessary to avoid signal artefacts known as "spinning sidebands". Samples can thus not be recovered after NMR experiments for a continuous survey. These samples are commonly snap-frozen and later thawed just before the NMR acquisition, causing cell damage and lysis that are often not reproducible. Then, ongoing enzymatic activity and metabolome evolution continues to be observed upon cell lysis and during NMR acquisition, so that additional heat-inactivation has recently been proposed for accurate measurements [1668], similar to earlier strategies using microwave irradiation of biopsy samples before analyzing them with HR-MAS [1669]. The delay between harvesting cells and freezing or analysis can also affect reproducibility. All these issues are also met with HR-MAS metabolic analysis of tissues [972,1622,1670], and have probably hampered its adoption among conventional diagnostic approaches [971,974,1068].

The appealing aspects of HR-MAS have nevertheless motivated improvements in different directions. First, efforts have been made to find pulse sequences and sample preparation protocols that decrease MAS rates without resulting in spinning sidebands. Pulse sequences suitable for slow-spinning rates have been adapted to HR-MAS conditions, and can provide exploitable ¹H spectra using MAS rates as low as ~40 Hz (PASS) [1031,1032] and even 1 Hz (PHORMAT) as shown for rat liver tissue [1033]. However, these techniques achieve these outcomes at the expense of NMR ease-of-use and can even cause ~10x sensitivity losses in the case of PHORMAT [636,1671], which probably explains why their use has rarely been reported in the last 20 years, to the best of our knowledge. Techniques have been proposed that recombine spectra obtained at various MAS rates of about 300-700 Hz, which permits localization and removal of spinning sidebands [1672,1673].

In the last 10 years, more attention has been paid to sample preparation: reducing the sample size to a few microliters, shaping it as a sphere, or carefully removing air bubbles from the rotor limits the appearance of spinning sideband artefacts at MAS frequencies of ~400 Hz [1030,1662,1674]. This rules out any renewal of oxygen in the sample. These conditions generate forces of only 1,000 g at which lumpfish eggs, for example, remained intact. Also because of its favorable, spherical shape minimizing the adverse

consequences of field heterogeneity, one single egg (~1 mm diameter) provided excellent NMR resolution using HR-MAS at 400 Hz [1030].

At such spinning rates, attention must be paid to J-coupling modulation during spin echoes, which can greatly alter signal quality [1030]. The non-J-modulated spin echo PROJECT [1650] has been usefully adapted in HR-MAS experiments by Caldarelli and Vermathen and their coworkers [1022,1028,1030]. Simpson and colleagues have also proposed the use of a low power adiabatic spin lock to filter out fast-relaxing species [1039], an approach that is further improved by diffusion-filtering [1006,1039]. A proper water suppression scheme is very helpful in clearly removing spinning side-bands, as shown again by Simpson and colleagues [929,1039].

Sakelariou, Wong and colleagues have designed inserts that integrate a rotating coil, enabling analysis of 100-500 nL samples by HR-MAS [1662,1675-1678]. Combined with NMR pulse sequences suitable for low spinning rates, these micro-HR-MAS systems provided a sufficient signal-to-noise ratio for metabolic analysis on a single *C. elegans* worm at a spectrometer field of 1 GHz with 300 Hz MAS [1041,1679,1680].

The development of slow (300 Hz) micro-HR-MAS systems by Wong and colleagues has even permitted analysis of samples containing only ~19 million yeast cells (~250 nL) and enabled their metabolome to be distinguished at various stages of growth or in osmotic stress [1042].

It is also important to emphasize the fact that HR-MAS rotors are hermetically closed before being inserted in the spectrometer, which prevents air exchange and brings about anaerobic conditions. Simpson and colleagues have shown that a "simple" hole drilled in the rotor cap allowed sufficient renewal of oxygen to keep fresh-water shrimps alive in the rotor [1006,1007]. According to the authors, the rotor spinning creates a vortex, which favors air exchange without buffer spilling or rotor instabilities. This rotor cap customization was also used to ensure aerobic conditions in a study on the fungus *N. crassa* [474].

Overall, problems arise from the centrifugal forces exerted on cells in HR-MAS NMR spectroscopy. These are not so problematic for microbial organisms, and novel techniques are being developed to achieve HR-MAS experiments at lower spinning rates. The resulting improved experimental conditions make it possible to study small animals, and, perhaps, organoid metabolism may also be monitored in the future using HR-MAS.

### 3.3.5. *Other solutions to sample heterogeneity, molecular diffusion heterogeneity and sample evolution*

We have seen in previous sections that sample heterogeneity causes peak broadening, which greatly hampers NMR analysis. We limit our discussion here to bulk magnetic susceptibility inhomogeneities, because in-cell experiments are usually performed in a constant, locked and shimmed magnetic field. Deleterious effects of such inhomogeneities are particularly problematic for ¹H-detected NMR, supposedly the most sensitive and versatile spectroscopy, because of the high gyromagnetic ratio of ¹H and its ubiquitous presence in biological samples. As described above, HR-MAS can eliminate some of the negative effects of sample heterogeneity, but it requires high spinning rates that can damage live cells and tissues. Other approaches have been applied to biological samples.

First, when the problem arises from the existence of different intra- and extra-cellular magnetic susceptibilities, it is possible to compensate using paramagnetic, or shift agents. This approach has been used notably in the 1980's to study erythrocytes: these contain high concentrations of hemoglobin, which can be paramagnetic in its deoxygenated form. Extracellular paramagnetic species can be used to match the intracellular magnetic susceptibility [1610,1681,1682]. This ensures lower intracellular
field gradients, and thus slower transverse relaxation times (see 3.2.2.). Whether this approach can be useful for cellular systems other than erythrocytes is questionable, but recent studies reported experimental values of mammalian cells magnetic susceptibility that suggest it may be [1581,1582,1583].

Pulse sequences exist that provide high-resolution spectra in inhomogeneous fields. Interesting research has been achieved using intermolecular zero-, single- or multiple-quantum coherences. In its later forms, this approach relies on establishing long-range dipolar couplings between protons of water and those of solute species [1683,1684]. In a rough simplification, this permits the correlation between local water chemical shifts in the indirect dimension (hence encoding the magnetic susceptibility variations) and local solutes chemical shifts in the direct dimension of 2D spectra. Post-processing treatment of 2D spectra can shear them back to corrected "inhomogeneity-free" spectra (Fig. 33) and yield high-resolution 1D projections. This has been used successfully on very inhomogeneous samples such as fish eggs [1685], grapes [1069], earthworms or water fleas [1686]. It has been combined with a supplementary J-resolved dimension on intact fish or on pig brain tissues [1687-1689]. At its current best, this approach suffers a 90% loss in S/N ratio per unit or time in 1D spectra [1686]. Arguably, a new type of quality criterion may be introduced, which would be the S/N per line-width ratio per unit of time.

Spatially encoded NMR spectroscopy, notably pulse sequences related to the Zanger-Sterk method for obtaining pure shifts, is also capable of removing field inhomogeneities. These rely on slice-by-slice resonance-selective excitation, decoupling and detection along the z-axis, and specific acquisition of chunks of the FID where the J-coupling evolution is negligible. This approach has been very well described recently in a didactic review by Dumez [1690]. To the best of our knowledge, it has been applied recently and only once to a heterogeneous biological sample, namely grape [1691]. This approach also suffers from low S/N and does not address the inhomogeneity in the x-y plane, although spinning the NMR tube can help with this. Above all, such spatially selective strategies cannot cancel the effect of sub-millimeter field inhomogeneities.

Temporal evolution of a living sample is the source of another type of inhomogeneity. This is commonly met when chemical reactions take place (e.g. metabolism, protein post-translational modifications) or when environmental conditions vary (e.g. exposure to chemicals). A classical approach to solve this problem relies on the successive recording of time series of NMR spectra. This implies a prior knowledge of the evolution timeframe to choose an appropriate acquisition time. ULTRAFAST approaches [1690,1692,1693] are rarely expected to be helpful for examining living samples, because the molecules to be detected are often at low concentrations. Time-resolved non-uniform sampling (TR-NUS) has been developed in the last years as a very specific approach to solve the issues related to temporal evolution. This method relies on the following principle: the NMR signal is acquired using a single uninterrupted experiment, during which the increments in the indirect dimension are allowed to vary in a randomized fashion. Sets of data points can be later extracted, which correspond to a limited experimental time window: even though they only partially sample the indirect dimensions, they permit reconstruction of the 2D or 3D spectra using dedicated NUS algorithms [1694]. This approach has been developed notably by Orekhov, Kazimierczuk and colleagues, and applied to characterize protein phosphorylation reactions in vitro [1695], bacterial metabolism [1061] and the metabolic response to growing toxin concentrations of water fleas [1066]. TR-NUS offers the appealing possibility to choose the experimental time windows in a post-acquisition fashion, and to use overlapping subsets of data or moving time-windows. The most suitable balance

between S/N, resolution and time-frame can thus be found. The available software to process TR-NUS data should help in making this approach popular [1696].

### 3.3.6. Hyperpolarized metabolites for live-cell studies: 10,000 times more signal, but how many constraints?

Sensitivity is a perennial concern for NMR applications. It is set primarily by poor overall nuclear magnetization under usual experimental conditions, i.e. room temperature and a 1-10 T magnetic field: at equilibrium, it does not reach more than ~0.01% for ¹H, the highest γ nucleus. This starting magnetization limits the finally detected signal level. Enhancing nuclear magnetic polarization has been one of the most sought-after goals for NMR spectroscopists. For the purposes of achieving this goal, only "dissolution-Dynamic Polarization Enhancement" (d-DNP) has been consistently used for cellular metabolism studies, although para-Hydrogen induced hyperpolarization (PHIP) shows interesting capabilities too.

We have described applications of d-DNP and PHIP for real-time metabolic studies in live cells in Section 2. Although substantial, these achievements have not yet resulted in d-DNP becoming widely adopted as a regular approach for characterizing metabolism. In the following paragraphs, we highlight some key pieces of information on d-DNP, in an attempt to popularize its capabilities (and shortcomings) for non-specialists. Dedicated experimental designs and NMR pulse sequences have been developed to exploit these dissolved hyperpolarized species, and have been reviewed recently [1697,1698].

To increase the starting equilibrium nuclear magnetization, temperature is a possible parameter to tune that could bring great benefits theoretically: adopting a Boltzmann equilibrium, the useful magnetization of dipolar nuclei like ¹H, ¹³C or ¹⁵N is proportional to tanh(-γhB₀/2k_B T), where γ is the gyromagnetic ratio of the considered nucleus, h and k_B are the Planck and Boltzmann constants respectively, B₀ is the polarizing magnetic field and T is the absolute temperature. Reaching sub-Kelvin temperatures would theoretically allow nuclear polarization to reach over 1% [1699]. Moreover, polarizing agents carrying free electrons can transfer the high electron polarization (γe.~660γ₁H, ~2600γ₁₃C) to the nuclear spins upon microwave irradiation at appropriate frequencies, a strategy commonly called Dynamic Nuclear Polarization (DNP). This theoretically permits one to reach 10-50% polarization at temperatures below 50 K. DNP-enhanced NMR is a very active field, which has recently gained strong interest and support, but its fundamentals and developments are beyond the scope of this review (see [1073,1074]). For fruitful use in metabolism analysis, it is necessary to implement i) fast dilution as a means to quickly thaw samples hyperpolarized at cryogenic temperatures, and ii) fast transfer to the analyzing magnet. Since the pioneering studies by Ardenkjaer-Larsen and colleagues in the early 2000's [1070], the promising 10,000- to 50,000 fold signal enhancements have been regularly achieved and d-DNP investigations have flourished [1071,1072].

Para-hydrogen induced hyperpolarization (PHIP) has recently been explored as a source of hyperpolarization for metabolism studies on living cells [1138-1140]. It relies on dihydrogen hyperpolarization at ~30-70K, and gives access to high magnetizations at lower cost than d-DNP.

PHIP is however much less versatile than d-DNP if we consider the number of metabolites that can be polarized, even though great advances have been achieved showing the possibility to hyperpolarize pyruvate, fumarate, acetate, carbohydrates or amino acids [1700-1707]. Several methods exist to use para-hydrogen hyperpolarization [1708], which we will not discuss here given that there have been only a handful of applications with cells. We expect however that these will multiply in the future, notably
<figure><img src="image_40.png" /><figcaption>**Fig. 33.** **A)** Operating principle of intermolecular single-quantum correlation spectroscopy (iSQC), which correlates solute and water local resonance frequencies resonances and therefore allows a post-acquisition correction yielding an "inhomogeneity-free" final spectrum; ¹H-NMR spectra were recorded with a de-shimmed sample containing Tyrosine and Arginine; **B)** ¹H-NMR spectra of a live earthworm *E. fetida* recorded with a standard presaturation W5-WATERGATE pulse sequence (upper panel), and the IP-iSQC pulse sequence (lower panel) [1686]. Adapted from [1686].</figcaption></figure>

thanks to the efforts being spent in improving the biocompatibility and purification of catalysts and products [1707-1710].

D-DNP and PHIP have however been used for real-time monitoring of metabolites that occur only at relatively high concentrations, i.e. in the millimolar range. This is because the technique currently has a number of limitations. First, the initial magnetization cannot be replenished once the experiment starts, whereas monitoring chemical reactions in real-time requires recording of a time-series of NMR spectra. This means a series of spectra can only be obtained by using small flip-angle pulses: a pulse with a flip angle of only ~1-15° leaves more than 95% of the magnetization available for subsequent scans [1072]. However, the cost of using this approach is that the sensitivity of each spectrum is decreased substantially relative to the full intensity for a single spectrum obtained after a 90° pulse.

A still worse problem is that in solution, the T1 relaxation of the studied ¹³C nuclei ranges from 1 to 50 s for most metabolites at commonly used magnetic field strengths. This means that the hyperpolarization has to be used within minutes of its creation at best, before the global ¹³C-magnetization has relaxed back to its thermal equilibrium value in the order of ~0.001%. Fortunately, a number of important metabolic reactions are fast enough to meet this constraint. [1-¹³C]pyruvate is often used because of its "long" hyperpolarization half-life, c.a. ~30 s, compared to about 10 s for [U-²H,U-¹³C]glucose and just one second for protonated ¹³C-glucose [1079,1112]. Intracellular ¹³C-T1 relaxation has even been shown to be about 3 times faster than extracellular for hyperpolarized [1-¹³C]pyruvate, [1-¹³C]ketoisocaproate, [1-¹³C]acetate and [1-¹³C]butyrate [1711]; moreover, ¹³C-T1s of such molecules are pH-dependent [1712]. This must be taken into account for the analysis of the reaction kinetics, together with an evaluation of the intra- and extra-cellular pools of observed molecules [1095-11011105,1106].

In complete contrast, DNP hyperpolarization build-up itself takes about an hour (even without including the manipulation time for preparing and properly freezing the DNP mixture). Moreover, it requires expensive polarizers and consumes significant quantities of liquid helium [1071]. Finally, DNP requires mixing the metabolites of interest with polarizing agents carrying unpaired electrons, which themselves greatly enhance T1 relaxation in the frozen state once microwave irradiation stops. Hence, the hyperpolarization vanishes within a few tens of minutes when

using classical DNP conditions (glassy frozen solution in presence of "DNP-juice") even in the frozen state [1071,1698,1713,1714].

A number of possible solutions to improve the approach are currently being explored. Many strategies are being developed aiming at storing hyperpolarized metabolites in their frozen state, which would be useful for clinical applications. First, the so-called "brute force hyperpolarization" method has been tested, which relies on i) sustained ¹H polarization for ~24 hours at ~1 K and B₀> 10 T, ii) magnetization transfer from ¹H to ¹³C nuclei at low field (B₀ < 0.1 T) to reach a ¹³C-polarization of about 0.1% and iii) storage at cryogenic temperature (a few hours at T~30 K and B₀~2 T) or dissolution-thawing before acquisition [1699,1715,1716]. In these conditions, carrying out deoxygenation and a slow-freezing annealing have a beneficial impact on hyperpolarization lifetime [1715,1716]. However, it is not clear that it will prove sufficiently competitive with respect to DNP. Second, hyperpolarization can be maintained for hours by improved preparation schemes that introduce enough distance between the polarizing agents (that would otherwise enhance T1 relaxation due to their free electrons) and the metabolite of interest [1713], and it can be further enhanced by storing in a high magnetic field at cryogenic temperature [1717]. Third, radical precursors (notably pyruvate derivatives) have been developed in which unpaired electrons are generated upon UV-irradiation at ~80 K, hence turning them into DNP polarizing agents, but which readopt their non-radical form at ~200 K. Such molecules permit efficient DNP at ~5 K in their radical form, before being quenched at ~200 K, leaving hyperpolarized molecules in a favorable environment for their storage where they have relatively slow T1 relaxation during storage [1714,1718-1721].

Once in the solution state, hyperpolarization should be maintained for as long as possible. Quaternary ¹³C nuclei are those likely to show the slowest T1 relaxation, because they have only weak dipolar interactions with other ¹³C nuclei. For the same reason, deuteration of other ¹³C nuclei drastically lengthens their T1 relaxation times. Importantly, the quadrupolar nature of ¹⁴N has a negative impact on T2 relaxation of nitrogen-linked ¹³C nuclei, which makes ¹⁵N-labeling useful as a T2/signal-enhancing strategy [1125,1722]. ¹⁵N nuclei themselves can be hyperpolarized and show longer T1 values because of the lower gyromagnetic ratio of ¹⁵N than ¹H, ¹³C or ¹⁴N, which may makes ¹⁵N an interesting nucleus for d-DNP studies [1135]. Hyperpolarization can also be
maintained using so-called "long-lived states" (LLS) for certain
molecules, recently reviewed in [1723-1725]; however this topic
is out of the scope of this review. Although this approach has
allowed spin order lifetimes of minutes to be reached, metabolites
of interest have not yet been found that could support LLS for cel-
lular studies at high magnetic fields. The common large chemical
shift differences between ¹³C nuclei prevented the use of LLS as a
means of detecting, for example, conversion of pyruvate to lactate
[1726]. LLS have been generated mostly on ¹H nuclei couples and
do not outcompete ¹³C T1 relaxation times under usual experimen-
tal conditions [1702,1725,1727,1728], notably because of the neg-
ative impact of dissolved oxygen [1702,1729]. Hence, LLS
applications for cellular studies have yet to appear.

Solutions to decrease the cost of d-DNP are being explored, such as the use of a cryogen-free polarizer that would save about 100 L of He per week [1071]. It has been also proposed to analyze simultaneously multiple samples in parallel, or in series, by mixing them with a single split preparation d-DNP hyperpolarized stock [1109,1730]. Altogether, hyperpolarization may hold great promise for studies monitoring real-time metabolism in live-cells, but d-DNP methods are still costly and PHIP approaches remain to be further established. We repeat the attractive ability of NMR to evaluate the metabolic status of cells or of organoids over long periods of time in a non-destructive manner. This would also be of interest as a complement to studies focusing on the intracellular behavior of proteins.

3.4. Observation of ligands

3.4.1. Technical recommendations for ligand observed NMR in presence of cells

The observation of small compounds by NMR can be facilitated
by the use of T2/T1rho filters, as described in Sections 3.3.2. and
3.3.3. The removal of background signals arising from macromolec-
ular cellular species is typically achieved using a short T1rho filter
[1476] combined with the STD/TRNOE pulse sequences, which has
also become normal practice for in vitro applications. The large sig-
nals from lipids can be removed using long adiabatic spinlock fil-
ters, as discussed in Section 3.3.3.

3.4.2. Technical recommendations for STD experiments in presence of cells

We want here to underline some technical aspects of STD
experiments on cellular samples. STD-NMR experiments rely on
the transfer of magnetization from the target protein to the
observed ligand, more precisely on the transfer of magnetization
saturation, i.e. null magnetization. Saturation reduces or eliminates
signal intensity by driving the spin-state populations towards
equality. It is produced by repeated, selective pulses (or alterna-
tively very low-power continuous irradiation) at a ¹H-frequency
where some protons of the studied protein resonate. Starting from
the initially targeted protons, the saturation spreads initially to
neighboring protons via the same processes of longitudinal relax-
ation as drive the NOE; then, via spin-diffusion (which is essen-
tially just a process of many successively repeated NOE transfer
steps), it spreads progressively throughout the whole macro-
molecule. as well as to ligand protons that contact the macro-
molecule during binding events. This saturation must be applied
selectively at a frequency where only protein nuclei resonate to
avoid any direct saturation of the ligand, an absolute requirement.
Hence, the on-resonance saturation pulses must be applied at a
resonance frequency distant enough from the ligand resonances,
and simultaneously excite a sufficiently broad resonance window
to maximize the saturation of the target protein [1731,1732]. A
careful, case-dependent evaluation should probably be carried
out systematically. Applying on-resonance pulses at very distant

frequencies is feasible on cells, because their very long correlation
time and heterogeneous environment cause very broad linewidths.
On-resonance saturation at -4 ppm from the closest ligand reso-
nance is very effective for virus like particles for example, as shown
by Peters and colleagues [1448]. Saturation spreads rapidly in cel-
lular macromolecules because of fast longitudinal relaxation asso-
ciated with long correlation times. Two to four seconds-long
saturation pulse-trains are usually applied to build a consistent
saturation transfer through a maximal accumulation of binding
events. However, a clean epitope mapping requires saturation
times shorter than the T1 relaxation times of the observed ligand
nuclei, otherwise saturation will spread into the rest of the ligand
beyond the epitope [1448,1733,1734].

The transfer of saturation from the protein occurs while the ligand is bound, and acts to partially decrease the signal intensities of ligand nuclei that contact the saturated protein. To observe these intensity losses in the saturation spectrum, it is subtracted from a reference spectrum. This reference spectrum is acquired with the same pulse sequence, but the saturation pulses are applied at an "off-resonance" frequency that is sufficiently far removed from the protein spectrum that no protein resonances will be affected at all, even notwithstanding the very broad protein signal linewidths. Peters and colleagues have shown for virus-like particles that even off-resonance ¹H irradiation at -80 or +80 ppm (at 500 MHz) still exerted observable saturation for these large objects [1448]. They recommended using an off-resonance saturation frequency of -300 or +300 ppm for STD studies on cellular systems. We will not list all the later publications that reported off-resonance irradiation at only +/-20 or 30 ppm and probably therefore obtained unreliable results, but we note that such issues are not rare.

Another issue is the slow T1-relaxation of cellular species, nota-
bly that of surface proteins. Again while studying virus-like parti-
cles (estimated correlation time of 75 µs), Peters and
Rademacher have shown that interscan delays of 25 seconds were
necessary to avoid significant unwanted effects due to residual
protein saturation during the off-resonance experiment (in
schemes using interleaved on/off experiments) [1735]. This results
in a clean epitope mapping, but reduced interscan delays may be
more efficient in terms of S/N per unit time, while still delivering
STD levels that are sufficiently contrasted to distinguish between
ligand protons contacting or not-contacting the studied target.

STD-NMR has often been shown to be more sensitive in D₂O
buffers [1731,1736]: these reduce saturation transfers between
the protein and bulk water, hence maximizing the transfer of pro-
tein saturation directly to the protonated ligand, even though it
slows down the longitudinal relaxation and increases the required
length of interscan delays. Working in deuterated buffers is accept-
able for viruses or isolated membrane fractions, but cells rapidly
deteriorate in these conditions and caution is recommended.

Concerning sample preparation, Caffrey and coworkers have
shown that serum proteins can establish weak interactions with
the observed compounds, which are typically those detected by
STD-NMR [1454]. STD pulse sequences in normal use cannot satu-
rate selectively cellular species as opposed to serum species.
Advanced saturation schemes using isotope-filtering exist
[1737,1738] that would allow a selective saturation of cells, but
this would require prior cellular ¹³C/¹⁵N isotopic labeling. One
can also consider using ¹³C/¹⁵N labeled antibodies to transfer the
magnetization saturation [1739]. Martinek and colleagues have
shown that this approach works in cell extracts, but there would
likely be a number of complications in the context of living cells.
Moreover, it is probable that these strategies would severely com-
plicate the quantitative interpretation of STD results. Of course, the
simplest option is the use of serum-free medium during the NMR
experiments, which is acceptable for limited measurement times
of a few hours.