As mentioned in Section 2, STD-NMR experiments on cells are most often performed in a STD-Difference (STDD) mode: two STD spectra are recorded, for example in the presence of mock-transfected cells (i.e. expressing only native cellular material) and in the presence of cells expressing the protein of interest. This is for two reasons. First, it would not be possible to use the choice of saturation frequency to select between the macromolecule of interest and the cellular background, as all signals of the macromolecule would inevitably be overlapped with signals from the cellular background. Second, saturation is anyway rapidly transmitted to the surrounding cellular species. Hence, compounds that only bind transiently with cellular membranes would show STD signals, but these should not be interpreted as an interaction with the membrane protein of interest. Moreover, a compound can also bind another endogenous receptor and consequently build-up STD signals. Finally, it is also important (and sometimes difficult) to be aware of the potential effects of variations in cellular metabolism and composition: the states of cells during NMR acquisition should still be as similar as possible to avoid any bias (e.g. binding to novel lipids, or phase-separated stalled mRNAs, etc...).

Finally, corrupted results may emerge from experiments using cells that progressively settle towards the base of the NMR tube. An elegant solution for this has been proposed by Jimenez-Barbero and his coworkers, that employs disposable rotor inserts designed for HRMAS [1475]: rotors can be filled with cell suspension and spun at a few kHz, which did not compromise cell viability according to the authors. Cells are thereby homogeneously sedimented on the rotor walls and STD experiments can be carried out. This approach was proposed almost 10 years ago, but we did not find any other later study in which it was used. The recent protocol by Konrat and colleagues may be easier to implement: cells are embedded in methylcellulose hydrogels to maintain a homogeneous dispersion, and settling in the NMR tube does not occur [1487].

### 3.4.3. On the use and present limits of on-cell NMR and the use of STD/TRNOE

As mentioned in Section 2, in the last 20 years ligand-observed NMR methods (STD-NMR, waterLogsy, T1rho/T2 relaxation) have been have been widely and successfully used for fragment-based drug design in vitro. The same has not been true for in-cell studies. Ligand-oriented NMR screening methods are valuable specifically because they detect weak affinity interactions [1386,1412], but these may often be uninterpretable in complex media: abundant metabolites also establish weak interactions [1413,1414], and reli-

ably identifying interesting hits would require highly reproducible metabolic states that current bioreactors can hardly generate.

Instead, in-cell NMR studies using ligand-based NMR detection have been focused on epitope mapping and structure-activity relationships: STD-NMR and TRNOE approaches have been favored due to their ability to deliver atomic-scale information on the interacting moieties of ligands and on their bound conformations. Although other ligand-observed NMR methods, such as waterLOGSY or T1rho relaxation enhancement, would presumably also work in complex media, they have not been extensively used for cellular NMR studies of drugs, probably because they do not provide structural information. However, waterLOGSY and T1rho relaxation enhancement have been shown to offer better sensitivity to binding events [1379,1413,1736,1740]. They may find increased application in the future.

Indeed, we feel that the capabilities of in-cell/on-cell STD/TRNOE/waterLOGSY/T1rho approaches may be exploited more broadly. On paper, their attractions are clear: it would be convenient and useful to observe direct contacts between a ligand and its membrane receptor in a native environment, without having to establish difficult purification and solubilization strategies. Defining standardized protocols for in-cell epitope mapping would promote trust and reproducibility. The use of isolated membrane fractions is likely to give access to simple protocols. However, we feel that avoiding membrane separation would be more convincing for biologists. Generating stable transfected cell lines for every protein target would thus be a prerequisite: it would yield reproducible expression of target proteins in the long term. Moreover, to permit the use of intact cells, experimental designs that avoid cell sedimentation are probably key success factors, as has recently been proposed [1487]. Applying automated procedures developed for background noise removal in metabolomics spectra may also be helpful to simplify the analysis of NMR cellular drug screening [1741,1742].

One reason why these epitope mapping approaches are not that popular for in/on-cell NMR studies may be the nature of the targets chosen to date: almost all publications of cellular experiments based on STD and TRNOE measurements have concerned surface proteins, mostly GPCRs [1469-1474,1489,1493] and integrins [1476,1479-1487] (see Section 2). Ligands of GPCRs commonly have long residence times with off-rates of 1 to 0.01 min⁻¹, as characterized by radioligand assays [1743]. Similar orders of magnitude have been reported for FDA-approved kinase inhibitors in vitro [1744]. Unfortunately, STD and TRNOE have the same limits with cells as they do with purified proteins: they can report on

<figure><img src="image_41.png" /></figure>

<figure><img src="image_42.png" /><figcaption>**Fig. 34.** Distribution of protein-size as a function of protein-copy-number per cell within two tissue types (adapted from Wiesniewski 2014 [1175]). Every dot represents one experimental point from mass-spectrometry quantification. The red line indicates the average protein size for a given copy number per cell. Important differences can be observed between cultured cell lines and the tissues from which they are derived [1176].</figcaption></figure>
interactions showing relatively weak affinities and fast on/off rates (Kd~10⁻⁷·10⁻³ mol/L, koff~10²·10⁶ s⁻¹ [1379,1412]). STD-NMR fragment screening on a solubilized GPCR has been recently proved to be feasible in vitro [1745], and, more generally, NMR has made recognized inputs in membrane protein drug research [1746]. Novel NMR approaches will have to be found for sensitive, ligand-observed studies of interactions with slow on- and off-rates, as has been discussed elsewhere [1379]. Such methods would find immediate applications for in- and on-cell studies. Alternatively, detecting ligand:protein interactions by the observation of the protein is a possible solution (see Sections 2.5.1. and 3.5.4.).

## 3.5. Cellular structural biology

### 3.5.1. How relevant is the intracellular concentration of the studied species?

If we had to select the single most crucial question concerning in-cell structural biology, that would be the issue of concentration. Conventional NMR structural biology studies use highly purified nucleic acids or proteins at high concentrations, i.e. in the 0.1-1 mM range, orders of magnitude above cellular concentrations. Exceptions exist, such as hemoglobin in erythrocytes (~5 mM). We know the protein copy numbers in many mammalian cells, which are the best characterized cells in this regard. Such cells are about 20 µm across, contain about 200 g/L of proteins (~4% from ribosomal proteins, ~4% from histone proteins) and contain an average of ~5.10⁴ copies per cell of every single protein species [1175-1177], which corresponds to a concentration of 1 µM. However, the distribution is broad: copy numbers span a wide range of values from 10² to 10⁷ per cell. In addition, all these numbers vary between cell lines and through the cell cycle [1747]. An interesting anti-correlation between protein size and copy number has been reported (Fig. 34). Moreover, many proteins of particular interest show high expression levels, e.g. oncogenes in cancer.

These micromolar concentrations are not generally suitable for acquisition of 3D NMR spectra, but they are compatible with 1D and 2D spectra. However, even when they are packed in wet pellets, cells cannot in practice comprise more than ~20-40% of the volume of an NMR sample, according to our tests (the actual volume of a cell is very difficult to quantify accurately [1175]). This unavoidably reduces the maximum number of molecules from which an NMR signal can be observed, which forces investigators to use unnaturally high intracellular concentrations of the macromolecule(s) of interest, i.e. above 10 µM and often above 100 µM (up to millimolar concentrations for structure determination in cells).

Daunting questions remain moreover concerning protein localization and NMR-detectability. In-cell protein NMR yields averaged signals from all the species that i) tumble sufficiently fast (solution NMR), ii) do not interact with too many partners, each one of these interactions generating a new fractional population in a distinct chemical environment, and thus a supplementary dispersion of the resonances (solution and solid-state NMR) and iii) do not bind their partners with on/off rates in the intermediate exchange time-scale (solution NMR). Among other issues, high concentrations of a macromolecule of interest can have saturating effects on the pool of native binding partners: if the total population of binding partners is engaged in interactions with the studied species, supplemented or overexpressed in excess, it can leave a population of molecules free and well-observable by NMR. Hence, situations can be generated where the NMR-detected population is no longer biologically relevant. These aspects should not be forgotten when reaching conclusions for in-cell NMR studies.

Hence, in-cell NMR-assisted structural studies are most often not likely to deliver results directly related to functional biology. Rather, such investigations explore the consequences of the cellu-

lar milieux on the structural behavior of macromolecules, which is nonetheless of fundamental importance. It would thus also be unfair to describe them as irrelevant simply because of concentration issues. These concentrations delimit the degree of generalization that can be claimed.

In any event, it is clear that an important goal for improving isotopic labeling schemes and NMR methods should be to help facilitate in-cell NMR studies at lower concentrations.

### 3.5.2. Sample preparation

Sample preparation for in-cell protein NMR has been reviewed multiple times. We discuss studies focusing on nucleic acids separately (see Section 3.5.6.). Detailed protocols exist for all sorts of samples in prokaryotic or eukaryotic systems, generated from in situ recombinant expression or from protein delivery in cells [4,9,25,1186,1236,1748-1753]. Here, we will simply recommend to check cell viability, protein integrity (by western-blot for example) and protein leakage. Quantifying the intracellular quantity of a protein of interest allows comparison of expected and experimental NMR signal intensities. This requires a good estimation of the signal losses due to ¹H/¹³C/¹⁵N T2 relaxation during the pulse sequence: these are rather different in a dilute reference sample than they are in the cellular environment. Such quantification is highly relevant for evaluating the percentage of detectable protein. None of these quality controls is individually perfect, but all of them should reinforce each other.

### 3.5.3. Protein structure determination

In the following, we discuss studies and methods using solution NMR. Although many solid-state NMR studies have reported structural information in cellular environments, these were not used for solving structures.

A normal protein structure determination procedure by NMR starts with the assignment of resonances, which classically requires at least a set of 3-dimensional spectra. Assignment procedures of NOE peaks are most often intertwined with structure calculation within recursive algorithms. This subject is well established from the field of protein NMR and is out of scope for this review. We will only mention that, as far as solution NMR is concerned, NMR assignment will always be hampered in cells, because of faster T2 relaxation (see Section 3.2.) and because 3D spectra necessitate long acquisition times that are not compatible with the lifetime of cellular samples. In this regard, using bioreactors helps to maintain cells viable across extended periods [1188]. Improved strategies have been proposed, notably by Ito and colleagues, to adapt the usual in vitro approaches to the constraints generated by cellular samples. These aim at shortening the acquisition time of 3D spectra by using sampling schemes that reduce the number of increments required [1184-1188,1754]. They also use amino acid specific labeling to decongest the spectra. It is recommended to use 3D pulse sequences that are intrinsically less vulnerable to T2 relaxation losses, such as HNCA, HN(CO)CA, CBCA(CO)NH or NOESY-HSQC.

Another approach relies on the use of lanthanide tags: linking these at multiple selected positions (in separate samples) permits measurement of pseudo-contact shifts (PCS), which provide a very powerful NMR restraint for structure calculation [1755-1757]. Structural models were obtained from tagged proteins delivered in frog oocytes or mammalian cells [1193-1195]. The PCS can be analyzed by the GPS Rosetta algorithm [1758]. This strategy proved to be efficient to solve the structure of small model proteins in cells, using ~50 µM intracellular concentrations. It has been used mostly to obtain backbone restraints, and is probably not likely to provide high resolution structures. Moreover, lanthanide tags can cause protein alignment in the magnetic field, which may be sufficient to measure residual dipolar couplings (RDCs). Although
RDCs have not been as useful as PCSs in the studies mentioned above, some amide ¹H-¹⁵N RDCs were measured to be as strong as 25 Hz at 600 MHz. They could possibly provide valuable information in future structural/dynamics studies at higher fields. PCSs can provide information up to ~4-5 nm away from the lanthanide tag, while RDCs do not depend on the distance to the tag. As in classical studies on purified proteins, both PCSs and RDCs are of course complementary to NOE-derived structural restraints.

### 3.5.4. Protein:protein interactions and protein:ligand binding

Questions related to intracellular protein diffusion and multiple, transient interactions with cellular constituents have been discussed in Sections 3.2.1. and 3.2.3.

Strategies to characterize protein:protein interactions by NMR in cells have been developed, notably by Shekhtman and colleagues: E. coli cells are i) transformed to carry two plasmids coding for the inducible expression of the two protein partners of interest, ii) grown in Luria Bertani (LB) medium containing a natural abundance isotopic distribution (i.e. 1.1% of ¹³C and 0.4% of ¹⁵N), iii) transferred into a minimal medium containing ¹⁵NH₄⁺ as sole source of nitrogen, in which recombinant expression of the protein target is triggered for a few hours, iv) re-transferred into an LB medium where the recombinant expression of the second protein is triggered [1232,1233,1274]. Hence, the first recombinant protein (but not the second) is ¹⁵N-labeled in a mostly ¹⁴N cellular background. Long-lived molecules produced during the incubation in the minimal medium are also ¹⁵N-labeled, but represent generally a heterogeneous pool of species, whose spectroscopic signature is sufficiently diffuse to not perturb selective NMR observation of the protein of interest. Protein:protein interactions are thus revealed by the ¹⁵N-protein signals that shift or disappear upon expression of the second recombinant (¹⁴N-labeled) protein. This approach has been used to screen inhibitors or stabilizers in cells, in an attempt to find a complementarity using high-throughput screening approaches [1390,1391].

Luchinat, Banci and their coworkers reported an approach to express and isotope-label target proteins sequentially in mammalian cells [1759]: i) a stable cell line overexpressing a first protein is established and grown on a natural abundance medium; ii) cells are transfected concomitantly with a plasmid encoding for a second protein and silencing siRNA for the first protein; iii) cells are later transferred into a medium supplemented with ¹⁵N-labeled amino acids. The precise timing between steps ii and iii is critical to restrict ¹⁵N-labeling of the first protein and of the cellular background, while maximizing that of the second protein. A more classical co-transfection and overexpression of two protein partners requires finding the best proportions of the corresponding plasmids and of the transfection reagent. This strategy has been used by Luchinat, Banci and their coworkers, notably to co-transfect and co-overexpress a protein and its chaperones in appropriate proportions [9,1280,1283]. However, this strategy results in an identical isotope-labeling of both proteins.

Protein-observed ligand binding assays have been carried out in cells based on single overexpressed proteins, either in prokaryotic or eukaryotic systems [867,1392,1393,1760,1761]. After overexpression in an isotope-labeling medium, cells can be exposed to the ligand, which provides simultaneously several pieces of information: i) does the ligand bind the protein target in cells and if so with what apparent Kd?; ii) does the ligand penetrate the cells and if so at what rate?; iii) does ligand binding have an impact on the protein target conformation or post-translational-modifications? Naturally, as discussed in Section 3.2.1., the usefulness of the experiment depends on the protein concentration, its cellular localization and its NMR-detectability in cells (is the whole population detected or only a limited proportion?). This approach is nevertheless very promising. It could be further improved by

designing experimental procedures i) to obtain valuable NMR spectra using lower and better controlled protein concentrations; ii) to establish reproducible conditions using more standardized bioreactors but also more biologically relevant cell lines.

### 3.5.5. In-cell solid-state NMR and the road to DNP-enhanced in-cell NMR

Solid-state NMR (ssNMR) using Magic-Angle Spinning (MAS) potentially has a great advantage over solution NMR in that it can acquire signals from proteins immobilized in membranes or in large macromolecular assemblies. As briefly explained in Section 3.3.4., MAS can suppress the effects of magnetic susceptibility inhomogeneities or unfavorable line-broadening due to slow molecular tumbling, which causes large dipole-dipole interactions, chemical shift anisotropy, or first order quadrupolar interactions.

ssNMR has consistently provided useful information on proteins and peptides in native membranes, starting with the work of Schaefer and colleagues on antibacterial peptidoglycan-binding drugs in the 1990's (see Section 2.2.2.). There is currently a growing number of ssNMR studies on transmembrane proteins in native membranes. To the best of our knowledge, only a few solid-state NMR investigations have reported de novo assignment in a native environment (bacterial membranes in most cases) [1173,1199,1752,1762-1764]. The many other ssNMR studies in the literature successfully transferred assignments made in vitro, in a similar fashion to what has been seen with in-cell solution NMR. ssNMR requires high MAS frequencies, which would affect cellular viability in physiological conditions (see Section 3.3.4). Frozen samples can preserve cell integrity during the ssNMR measurements, but they also suffer from peak broadening, notably also resulting from freezing heterogeneous ensembles of conformations and interactions with the cellular milieu. These studies often rely on carefully designed amino acid specific labeling to compensate their common poor resolution.

Most of the ssNMR structural characterizations in "native" conditions concern membrane proteins: the cellular membranes represent good 'native environments' and the removal of cytosol increases the proportion of useful sample in the rotor. A number of publications describe the isolation and packing of cellular membranes in ssNMR rotors following various protocols [1173,1197-1200,1203,1204,1207,1210-1211,1752]. Until recently, (DNP-) ssNMR studies on membrane proteins in native membranes relied on ¹³C-detected experiments. In principle, ¹H-detected spectroscopy could offer higher sensitivity. However, it requires either MAS above 60 kHz [1765] or extensive deuteration of the sample matrix to suppress dipolar ¹H-¹H relaxation, which makes it difficult to use this approach for complex biological samples. Medeiros-Silva et al. proposed a novel labeling scheme, namely inverse fractional deuteration, to obtain high resolution ¹H-detected ssNMR spectra of the K⁺ channel KcsA in liposomes and in native membranes [1209]. This approach improved the detection of water-inaccessible protons and enabled backbone assignment in vitro through a 'classical' strategy using triple resonance 3D spectra. These assignments were transferred to signals of the sample in situ via 2D ¹H-¹⁵N spectra, thereby permitting the characterization of as few as 3 nanomoles of TM domain in native membranes; in addition the authors carried out a detailed analysis of its motions in vitro through ¹⁵N relaxation experiments.

The first application of ssNMR to investigate a soluble protein in intact cells was published by the Dötsch group in 2012, in which Reckel et al. analyzed bacterial cells under freezing conditions using ¹³C-detected ssNMR [1212]. Bacterial thioredoxin and human FK506-binding protein (FKBP) were chosen because both proteins interact diffusely with the cellular environment: this causes extensive signal broadening at physiological temperatures, and prevents detection by solution NMR. In that work, freezing
temperatures were necessary to preserve cellular integrity. Also, ¹³C-depleted glucose had to be employed in the culture medium before protein expression, in order to suppress the unwanted signals arising from natural abundance ¹³C. In their concluding remarks, the authors noted that DNP would likely become an indispensable tool to increase the sensitivity of the method. Since then, DNP-ssNMR has been applied to study proteins in quasi-physiological samples such as cellular lysates and, more recently, directly in intact human cells. Frederick et al. investigated the folding and aggregation of the yeast prion protein NM (i.e. the prion domain of Sup35) in frozen yeast cell lysates using DNP-ssNMR [1172]. When pure NM was added at 1 µM to lysates already containing the prion form of Sup35, it adopted an amyloid form, which was more structured than the amyloid fibrils formed *in vitro*. Deuterated lysates were employed to render the cellular milieu NMR-invisible and to increase polarization efficiency [1170]. Viennot et al. used Bcl-xL as a model protein to assess the DNP enhancement selectivity in bacterial cell lysates, using a radical-labeled binding partner to selectively enhance the protein signals with respect to the background caused by other labeled cellular components [1766].

The first test of the application of DNP-ssNMR to intact human cells was published by the group of Barnes [1767]. Using biradicals and sterically shielded radicals as polarizing agents, DNP enhancements of ~50 were obtained at temperatures ranging from 6 to 90 K. A novel fluorophore-containing polarizing agent was introduced, which allowed simultaneous subcellular localization studies through fluorescence microscopy on the same cellular samples. The approach was recently combined with cell sorting through flow-cytometry to investigate the endogenous activation of the HIV virus by correlating GFP expression with increased ¹⁵N-amide resonance intensities in highly homogenous cell populations [1768]. The same research group further explored the effects of electron decoupling on cellular DNP-ssNMR spectra, reporting sensitivity improvements at cryogenic temperatures [1769].

A DNP approach to investigate intracellular proteins in frozen human cells was reported recently by Baldus and coworkers [1213]. They delivered isotopically-labeled ubiquitin to cells using electroporation. Then, they proved notably that their DNP polarizing agent diffused within cells using a fluorophore-tagged DNP-biradical: they observed a uniform fluorescence distribution 15 minutes after exposing the cells before filling the rotor with these cells. The preparation of cells for DNP measurement is still not a routine procedure: the mixing of the "DNP-juice", the timing, freezing method, etc., can all benefit from optimization. The Frederick group has recently addressed these aspects: they reported a detailed workflow for the correct cryopreservation of human cells prior to DNP analysis, which is crucial for preserving high cell integrity and viability [1770].

### 3.5.6. Nucleic acids

Successful NMR studies of nucleic acids in mammalian cells became frequent only in the last 2-3 years, so that systematic guidelines are difficult to define as yet. Previous studies have been mostly performed in microinjected frog oocytes, and have been reviewed multiple times [4,5,1323,1771]. Using current methods, the concentrations of the nucleic acid of interest can reach ~15 µM in mammalian cells, and ~100 µM in oocytes [1322,1329]. Just as for in-cell studies of proteins, checking viability, leakage and localization of the studied nucleic acids is mandatory, especially in cases where they have been delivered into the cells using membrane destabilizing methods.

Detailed protocols exist for sample preparation of phages and NMR analysis of their DNA content [1307,1311,1312,1772]. Such studies benefit from the rather standard and economical ¹³C/¹⁵N labeling procedures of bacteria. It is worth mentioning that supple-

menting the culture minimal medium with unlabeled amino acids together with ¹³C-glucose and ¹⁵N-ammonium helps to obtain NMR spectra showing almost only ribose signals; supplementing with unlabeled Phe, Tyr Trp, Tyr removes side chain resonances of these residues from ¹³C/¹⁵N-edited spectra, while retaining ¹³C/¹⁵N labeling of DNA bases [1310,1312]. ¹³C/¹⁵N labeling of nucleic acids is extremely helpful to characterize structural ensembles of nucleic acids, which commonly explore different conformations in dynamic equilibria occurring over a broad range of timescales [1291,1292,1294,1773]. Such labelling also helps to define any chemical modifications they may have [1331].

Unfortunately, there is a high financial cost for the methods used to date for preparing ¹³C/¹⁵N labeled nucleic acids, before delivering them in mammalian cells using the published methods, electroporation and pore-forming toxins (electroporation performing better in terms of cell viability and leakage according to the literature [1320,1322,1323,1326,1329]). Hence, the studies have mostly been restricted to observation of ¹H-signals from the imino groups, which are largely free of interference from cellular background signals [1321,1324-1326,1329]. ¹⁹F-labeled nucleic acids have been also tested recently, which have the advantage that ¹⁹F NMR has no cellular background signal, but these molecules could also modify the conformational ensemble being investigated [1326-1328]. In both cases, the stability [1329] and the multiple conformations and/or interactions [1326] of nucleic acids in cells can be a severe obstacle. As discussed for proteins in Section 3.2.3., multiple transient interactions in cells have consequences for the NMR detectability of nucleic acids. Interestingly, Trantirek and colleagues showed recently how G-quadruplexes of similar size can adopt very different structural dynamics, how these fluctuate in presence of G-quadruplex ligands, and how the corresponding NMR spectral fingerprints allow or prevent in-cell NMR studies [1326]; this group has provided some very useful detailed accounts of work in this field. The current in-cell NMR techniques are thus limited in the quality of structural information they can offer and in the number of nucleic acids that can be studied. Even though multidimensional NMR of nucleic acids seems often to be impractical in cells [1329], ¹³C/¹⁵N labeling and NMR-editing of single bases in the nucleic acids of interest might be used in the future to help elucidate conformations or interactions with ligands. Alternatively, monitoring the NMR signals of a ligand can sometimes be sufficiently informative [1329].

## 3.6. Whole cell analysis

### 3.6.1. Solid-state NMR analysis of cellular content, molecular proximities and chemical bonds

As mentioned earlier, solid-state NMR coupled to magic-angle spinning permits the detection of the whole set of molecules carrying a specific isotope labeling when edited by a chosen pulse sequence (see Section 3.3.4.). This capability has been extensively exploited by Schaefer and colleagues to characterize a number of connections in bacterial cell walls and to determine the position of some antibiotics in the bacterial cell walls (see Sections 2.2.2. and 2.5.2). It has also been used to quantify the cellular content of certain molecular species upon exposure to chemicals. These techniques are appealing particularly because they do not require prior extraction steps before the NMR analysis.

In the early 1980's, authors used the so-called Double Cross-Polarization (DCP) [1774] from ¹H to ¹³C and then to ¹⁵N to enable selective observation of branched ¹⁵N-Lys and ¹³C-D-Ala [584-586]. However, these experiments suffered from weak signal intensities, and moreover were contaminated by signals from cellular components containing ¹⁵N at natural abundance (these were removed during processing by spectral subtraction). From the late 1980's, Schaefer and coworkers started i) to remove cellular back-
ground using ¹³C-¹³C connectivity filters (DARR), which they demonstrated for the selective observation of [1,2-¹³C₂]-Gly in Klebsiella pneumoniae [592] and ii) to improve the sensitivity of ¹³C-¹⁵N proximity filtering using the recently invented REDOR [593] and TEDOR [1775] techniques. The combination of DARR ¹³C-¹³C and REDOR ¹³C-¹⁵N or ¹³C-¹⁹F was indeed found to be effective. Importantly, the REDOR approach depends on dipolar couplings, which are distance dependent: distance measurements became possible between ¹³C and ¹⁵N nuclei [606], and later between ¹⁵N and ³¹P (for the analysis of DNA in phages [1306]), or ¹³C and ¹⁹F (for analyzing ¹⁹F-containing molecules inserted into cells). Hence, the studies described in Sections 2.2.2. and 2.5.2 rely substantially on REDOR or TEDOR. We thought it would not be appropriate to present them extensively here, instead citing relevant publications [570,587-591].

### 3.6.2. The case of hyperpolarized ¹²⁹Xe, a multitask MRI-oriented reporter

Hyperpolarized ¹²⁹Xe detection represents a peculiar case in the zoo of observable species that have been used for NMR cellular studies. We mentioned ¹²⁹Xe-based sensors in different sections for various applications. These have been developed for the purpose of designing new sensitive probes for MRI, e.g. to detect cell surface proteins [1776-1779], or to probe pH [1780,1781] and intracellular chemical entities like thiols or ribose [1782,1783], without any fundamental interest for cellular processes themselves. This topic will not be covered further here, but some excellent reviews on ¹²⁹Xe based sensors have been published [1784,1785].

## 3.7. The quest for a bioreactor

### 3.7.1. A re-emerging necessity

Keeping cells healthy in the spectrometer has been a long-term goal in the field. It is necessary for real-time monitoring of metabolic changes or molecular interactions, or to maintain cells in well-defined conditions when long acquisitions are required. We found about 200 reports using or presenting the design of an intra-spectrometer flow-probe bioreactor since the 1970's. The NMR-compatible bioreactor has been re-invented dozens of times, but the individual experimental designs apparently were not widely taken up beyond the labs where they had been developed. Although not fully exhaustive, our statistics show two main periods of activity (Fig. 35). Between the 1970's and the year 2000, a first period started with experiments on perfused animal organs, progressing later to home-made systems for metabolic studies. Since the 2000's, a second period saw in particular the emergence of low-diameter cryoprobes, whose higher intrinsic sensitivity is offset here by their small diameter (< 10 mm), close design and lower flexibility. Those familiar with standardized commercial systems are probably less tolerant of setting up and tuning home-built probes.

Initially, systems to perfuse organs were used in the 1970-80's, mostly to monitor ion equilibria or metabolism in rat or rabbit hearts [277,378,807,812,818,856-859,861,1786-1789], livers [725-727,809,814,1790-1794] and kidneys [808,819], using ³¹P-, ²³Na- ¹³C- or ¹H-detected NMR [1795-1797]. Perfusion systems for NMR spectroscopy of living plants were also proposed [884-886,900,1798]. We will not cover this field further.

In the vast majority of cases, bioreactors were designed in the context of studies on cell metabolism. As discussed in Section 2.3.1., in-cell NMR studies of metabolic activities are more suitable for high concentration metabolites. Hence, NMR spectroscopy of extracted and isolated material became dominant in the study of metabolic pathways in the 1990's, in combination with Gas- and Liquid-Chromatography coupled to Mass Spectrometry

[39,762,798-800]. Although they require tedious extraction and calibration procedures, these approaches allow the application of reproducible, systematic protocols. In the 2010's, bioreactors reappeared in the literature as appealing devices for i) in-cell structural biology (see Section 2.4.), ii) metabolism studies using ¹³C-hyperpolarized metabolites; iii) longitudinal metabolism and toxicology studies on cells (including for online monitoring of external, large-scale reactors), iv) and also on small animals like worms or water fleas [930,1057,1066].

Many NMR-bioreactors were proposed in the 1980-90's for studying cell metabolism. They all involve pumping systems of culture media equilibrated with the appropriate CO₂/O₂ gas ratios. A description of these home-made designs would be quite repetitive and not very informative. Equivalent commercial devices have been proposed in the last ten years, which are compatible with the actual (cryo)probes now present in most laboratories.

### 3.7.2. Cell immobilization strategies

With regard to the conditions in which cells are maintained in the measurement volume, several options have been explored. First, cells can be kept in suspension in the NMR tube: There has been a long and varied history of many stirring or bubbling devices designed through the years, which avoid settling and clumping and ensure (an)aerobic conditions (see [783,838,841,1802-1819]). These designs are often not compatible with recent, narrow probe designs, where keeping cells inside the active volume of the tube despite the flow of medium has been a long-term challenge. A number of strategies have been proposed.

Flow-probes for suspension cells were initially developed that used dialysis fibers for the circulation of the fresh medium [1820]. Soon afterwards in the mid-1980's, an advanced version of this principle was proposed to provide rapid flow of medium and high O₂ delivery for aerobic cells [1821,1822]. A similar approach involved circulating the fresh fluids through the lumen of hollow fibers, whose polymer porous walls permit nutrient diffusion to the interspace of the fibers, where suspension cells are maintained [771,797,1823]. Adherent cells can also attach on the hollow-fibers walls, either inside [1824] or outside [789,791,797,1823,1825-1830]. Good results have also been obtained with a recent simplified version using a 1 MDa dialysis membrane, in which fresh medium flows continuously [1761]. The most popular approach has been to immobilize cells in beads [1811,1831-1833] or threads made of agarose [167,179,765,777,780-782,786-788,795,796,852,866,867,1354,180 1,1834-1850], alginate beads [784,785,790,869,1103,1105,1369, 1804-1806,1840,1851-1858], carrageenan gels [1859,1860], or low-temperature melting PNIPAAm-PEG gels [475,868,1861]. Other designs took advantage of improved matrices composed of methylcellulose [1487], poly-L-Lysine containing alginate [1862,1863] or biological material composing extracellular matrices like collagen or Matrigel, which are better adapted to cell proliferation [167,387,785,789,794,1799,1849,1853,1864-1867]. The diameter and composition of gel threads or beads has of course an impact on diffusion of ions [167]. Adherent cells can be grown on micro-carrier polymer beads [167,778,779,792,793,1093, 1099,1100,1112,1800,1865,1868-1874], on fibers [1800] or even on polyethylene terephthalate (PET) scaffolds [1875]. Cell spheroids and plant cells or root nodules have convenient characteristics: unlike single bacterial or mammalian cells, they do not require any immobilization to be retained in the NMR tube, and simple perfusion systems can be used [882,964,1876,1877], possibly equipped with loose nets and completed by stirring devices [1878]. Remarkably, insect cells were also maintained viable in suspension at the bottom of the NMR tube while using a low flow rate of culture medium (2 mL/h) [1188].
<figure><img src="image_43.png" /><figcaption>**Fig. 35.** A) Primary publications of studies (those known to the authors) reporting the design or use of intra-spectrometer bioreactors and flow-probes; B) Scheme of a typical NMR bioreactor, using a well-defined gas mixture bubbled into the culture medium, as well as culture medium heating and pumping into the spectrometer using inlet and outlet tubing to deliver fresh nutrients in a NMR tube (typically 5 mm diameter and 0.5 mL for recent probes) (adapted from Hertig et al. 2021 [1799]); C) Electron microscopy of anchorage-dependent human breast cancer cells attached to agarose polyacrolein microcarrier beads, polyester or embedded in agarose gel threads (adapted from Neeman et al. 1988 [1800]); D) Example of an empty flow unit showing the NMR tube and the inlet tubing (adapted from Luchinat et al. 2020 [867]); E) Examples of 5 mm NMR tubes filled with culture medium (its pink color indicates a pH ~7, while it turns yellow when it becomes acidic because of cell metabolism in absence of medium replenishment) and gel threads or agarose beads (adapted from Carvalho et al. 2019 [1801], Kubo et al. 2013 [868], and Inomata et al. 2017 [1369]).</figcaption></figure>

Each of these immobilization methods has advantages and drawbacks. Gel encapsulation is very well suited to studies of suspension cells, while micro-carriers represent an appropriate technique for preserving adherent cell integrity and immediate accessibility of nutrients [167]. Cell density is higher with gel encapsulation methods (up to ~2.10⁸ mammalian cells per mL) than with adherent systems (up to ~1.10⁷ cells per mL) [768,1864]. Agarose is not electrically charged, while alginate gelification relies on ionic interactions that require Ca²⁺ (or Ba²⁺) supplementation and low EDTA or phosphate concentrations. Hence, the two gels interfere differently with the diffusion of charged metabolites and O₂ [1840]. The low melting point agarose that is commonly used has a low mechanical strength and large pores; in contrast, alginate beads have a higher mechanical resistance and a finer meshwork [1840]. Alginate has been well characterized as a viable tridimensional culturing system for multiple purposes and cell types [1879,1880]: among others, it produces beads of uniform, tunable size, showing possibly hypoxic cores, which can be very useful, e.g. for radiobiological studies of cancer cells [1879]. Cells may have difficulties multiplying in dense agarose or alginate. In this regard, gels made of components of extracellular matrices such as Matrigel are better mimics of native environments, and allow cells to attach, grow and function in a more satisfactory fashion. They were also reported to interfere less than agarose with ionic diffusion [1849]. Methylcellulose also has superior characteristics to those of agarose, by allowing cells to adhere to its three-dimensional meshwork and ensuring improved cell viability [969,1487]. "Normal" cell growth is also possible on microcarriers, but attached cells are subject to shear stress. Such stress

is absent for cells grown on hollow fibers, but these occupy the largest fraction of the effective volume. Moreover, hollow-fibers require the application of high fluid pressures to exchange nutrients through the fiber walls. Finally, adherent cells that are only able to grow as a monolayer may provide low signal, both with micro-carrier beads and hollow-fibers [1864]. Spheroids can suffer from necrosis at their center [768,964,1876,1878].

### 3.7.3. *Particular problems for consideration*

To be effective and notably to achieve sufficient SNR, in-cell NMR often requires high cell densities. This makes nutrient or oxygen diffusion and delivery difficult, although these are essential for maintaining a constant "normal" cell metabolism [1881-1883]. Studies have been reported that attempt to define the best balance between cell density, flow rates and O₂ concentration in the perfused medium [1823,1828,1884,1885]. Non-invasive measurement of dissolved O₂ over time in the bioreactor is accessible, using perfluorocarbons entrapped in the gel matrices, whose T1 relaxation depends on O₂ partial pressure [1799,1853,1873,1886,1887].

Naturally, cellular composition and metabolic state vary over time. Notably, the compositions of lipids and their glycerol precursors are not the same depending on cell density on carriers or in threads [1800,1839,1864]. It is clearly important to perform control experiments to verify that cells are kept healthy: ¹³C- and ³¹P-NMR monitoring of metabolite consumption and ATP levels are good indicators, complementary to cell viability assays [867-869,1105,1761,1888].
Finally, as far as quantification is concerned, it is important to consider the fact that effective T1 relaxation can be affected by the flow rate and the resulting renewal rate of extracellular species: molecules freshly inserted in the tube are not perturbed by previous radiofrequency pulses and therefore provide more signal per mole than molecules that remain within the tube and do not attain their equilibrium magnetization between pulses. For example, the use of a long-T1 species such as inorganic phosphate as a concentration reference was shown to be potentially misleading because its signal intensity was dependent on the flow rate [1889].

### 3.7.4. Past and recent applications

Bioreactors have been used to characterize cell metabolism in real-time with ¹³C- and ³¹P-NMR. A broad variety of studies using NMR-bioreactors have been concerned with the effects of drug-exposure and irradiation on mammalian cancer cell metabolism [768,777-794,1093,1094,1099,1100,1103,1858,1866,1890], the impact of ethanol and chemicals on hepatocytes [795,796], metabolism in hybridoma [771,797] and bacteria [1819,1883,1891-1894], comparisons of the metabolism of cells in suspension or embedded in gels, which has industrial applications for yeast metabolism [1804-1806,1811,1831,1832,1859], and antibody secretion of hybridoma [866]. They also served in the study of intra- extra-cellular ²³Na⁺ or ⁷Li⁺ fluxes in mammalian cells [167,179,1844,1849]. Real-time monitoring has its advantages, by permitting the longitudinal observation of a single sample without repeated quenching and extraction of solutes. It can provide very well defined kinetics with dense timeframes, thus allowing advanced modeling of metabolism, as shown recently for melanoma cells [1874]. However, it requires the establishment of peculiar arrangements, and more classical metabolomics methods that employ solute extraction and high-resolution NMR and mass-spectrometry are more popular currently.

Hyperpolarization methods hold the promise of providing large sensitivity enhancements (see Sections 2.3.7. and 3.3.6.), and dedicated bioreactors have been proposed since the late 2000's. Cells can be steadily supplied with nutrients and possibly with drugs of interest, and can be exposed at controlled times to hyperpolarized metabolites (mostly ¹³C-pyruvate). The evolution of their NMR signal and those of their cellular byproducts reveals the metabolic states of the cells. The short half-life of metabolite hyperpolarization limits the application of this approach to fast enzymatic reactions that can be monitored within a few tens of seconds. As was the case in the 1980-90's, ¹³C-NMR of hyperpolarized metabolites has been studied in bioreactors to establish good references for future MRI/MRS applications [1093,1100,1103,1112,1867,1890], or to examine the effects of various drugs on various cancer cells [1093,1099,1100]. Classical immobilization methods were adopted, using alginate beads [1094,1103,1105,1850,1890] or cell attachment to polymer beads [1093,1099,1100,1112]. Original designs were also proposed: cells were grown on 3D scaffolds through which culture medium and hyperpolarized metabolites were perfused [1895], or embedded in a 3D collagen matrix placed in a 3D-printed bioreactor designed for allowing both NMR spectroscopy and optical Fluorescence Lifetime Imaging Microscopy (FLIM) [1867].

Real-time monitoring of cellular metabolism has also been carried out to characterize the evolution of large, external bioreactors. Continuous "online" monitoring can be achieved by circulating the culture broth through the spectrometer using a bypass tube [739,962,963,1611,1859,1883,1891-1894,1896-1899]: this approach gives access to scale-up studies of potential industrial applications in a non-invasive fashion, and is well suited to use with low-field magnets [962,963,1898,1899]. Using similar designs, circulating the supernatant medium of an external bioreactor into the magnet gives access to indirect monitoring of cell

metabolism that can be applied to organoids or microbial biofilms [1813,1900,1901].

Bioreactors are now becoming more mainstream for in-cell structural biology studies. They are clearly necessary to maintain cells in relevant physiological states during the required hours-long acquisition times [867-869,1761,1888]. Evidence exists that the studied proteins can have different folding states or interactions depending on the cellular metabolic state [1355,1369]. Now that early proofs of principle have been realized, which show that observing proteins and nucleic acids in living cells is possible, the field will have to move towards wider adoption of bioreactors in the future.

Overall, it appears that NMR spectroscopists have been very inventive in developing home-made designs to maintain cells in a healthy state in the spectrometer. These designs have a broad variety of applications, from the monitoring of industrial microbial bioreactors to the simple preservation of steady conditions for in-cell structural biology studies. This profusion of systems is a blessing and a curse at the same time. Most of the proposed home-built designs were somehow lost and not transferred from lab to lab, and more standard systems would probably have helped in developing in-cell NMR applications. Commercial solutions proposed recently are derived from devices dedicated to the online-monitoring of external chemical reactors, and are not fully satisfactory when it comes to nutrient delivery. We speculate that solutions will be found to this problem. Online-monitoring of external reactors has a promising future, notably thanks to the recent availability of benchtop, permanent magnets (operating at ~1 T): these are more easily adapted to conditions in the field at reasonable costs, and they should make non-invasive, real-time NMR analysis possible more widely [47,962,963,1899,1901]. Finally, we can hope that the efforts of many groups to miniaturize their designs will succeed and have interesting applications. The integration of micro-fluidics and alternative micro-coil systems may push the capability of NMR to become a non-invasive approach for the characterization of nano- to micro-meter size tissues, egg cells, animals or organoids for therapeutic or toxicologic studies [964,1066,1109,1902-1904].

## 4. Where next?

NMR spectroscopy is extremely versatile, and the variety of types of information that it has provided from cellular samples is astonishing. The main applications of in-cell NMR have varied through time. The original investigations on cellular water, salts and basic metabolic reactions provided a good basis for clinical developments, notably for MRI, MRS and metabolomics. Unique contributions are regularly delivered by in-cell NMR studies in various fields as shown throughout this review, from the in-situ characterization of cell-walls, of protein folding and drug binding, to non-invasive toxicology studies in small living organisms. However, the latest technological developments have oriented modern biological applications towards high-throughput or single-molecule/single-cell analysis, as discussed below. NMR spectroscopy is not a method of choice in this regard and is thus not among the most fashionable methods for cellular analysis currently. The particular capabilities of NMR remain intact, however: the ability to observe multiple nuclei, to provide atomic-scale information in a non-invasive fashion. Hence, even though the designs used in the past for in-cell NMR studies can now appear outdated (using outdated cell strains and culture conditions), we believe a number of strategies may be readapted to more contemporary contexts. This requires identification of complementarity with the most popular current techniques for cellular analysis, the systems that would benefit from non-invasive NMR observa-
tion, the new technologies that would empower the NMR capacities, and those recent NMR developments that may find their use in the analysis of cellular samples.

## 4.1. Comparison and complementarity with other spectroscopies, spectrometries and microscopies

Many other techniques can provide molecular information from intact cells. Focusing on the fields where NMR can contribute to establishing knowledge on cellular processes, we can distinguish two broad fields: in-cell structural biology, and live-cell enzymatic activities. Although they are likely to evolve rapidly, the specific advantages and drawbacks of other spectroscopies must be considered in order to understand how NMR may be useful, complementary or even unique. We will not give any technical details here, but only mention the present capabilities and applications of the major "in-cell techniques".

### 4.1.1. In-cell structural biology

We start this section by considering a technique that is particularly closely related to in-cell NMR, namely in-cell EPR. This technique can provide two types of information in particular [1905-1908]: i) the local structural dynamics of a paramagnetic tag on a protein at room temperature, ii) the evaluation of distance distributions between two paramagnetic tags attached on the nucleic acid or protein(s) of interest at cryogenic temperature. In-cell EPR is well-suited to distance measurements in the ~2.5-6 nm range using fairly small amounts of material (down to submicromolar protein concentrations in ~5-10 million cells, as reported recently [1582]), which can be either nucleic acids or proteins, doubly tagged with nitroxides [1909-1914], trityl groups [1915-1917], or Gd³⁺-based labels [1582,1918-1924]. They yield information on frozen distance distributions measured at cryogenic temperatures, which are commonly reached by using significant quantities of liquid helium. In-cell EPR is an expanding field and may represent an interesting method in the toolbox in the coming years.

Förster Resonance Energy Transfer (FRET)-based methods deliver similar tag-to-tag distance measurements in the ~3-8 nm range, but in this case between two fluorescent labels [1925]. This approach gives access to the subcellular localization of the characterized proteins in single cells, and has been used to study protein thermal stability [1377,1926-1930] and protein:protein interactions [1931,1932]. Single-molecule (or single-particle) FRET (smFRET) approaches allow measurements in live-cells not only of intra- and inter-molecular distance distributions, but also of conformational or binding exchange dynamics [1933-1938], including for membrane proteins [1939]. In combination with Fluorescence Correlation Spectroscopy (FCS), structural dynamics on timescales between ~10 ns and ~1 ms can be observed [1940,1941]. smFRET might soon be effective for in-cell structural modeling [1942] and probably will be an important source of in-cell structural information in the coming years. Similarly to the labels used in EPR, the labels used in such fluorescence studies have sizes in the approximate range 1-2 nm but their flexibility, their potential interference with the studied structural properties, their (photo)stability and the necessity to deliver the labeled proteins in cells (for the non-GFP fluorophores) are current limiting factors [1939,1940,1943].

It is well known that cryogenic electron microscopy (cryo-EM) has become a dominant technique in the field of structural biology during the past several years. In parallel, cryo-electron tomography (cryo-ET) has permitted very considerable advances in the observation of macromolecules *in situ*, i.e. in plunge-frozen cells or even small organisms [1944-1946]. The latest technical developments make it feasible to obtain tomograms of ~100-200-nm-thin cell

slices at ~3-4 nm resolution [1945,1947,1948]. At the molecular level, this has already permitted characterization of amyloid aggregates [1949-1951], tubulin- or actin-associated proteins [1952-1954], and membrane protein complexes [1955-1960] at nanometer and subnanometer resolutions using sub-tomogram accumulation and averaging. Recent improvements in data acquisition and computational treatment make cryo-ET capable of reaching resolutions of 3-4 Å [1961-1965], including in cells for ribosomal particles [1965]. Information on "invisible" parts of the complexes can be obtained by complementary analysis from in-cell cross-linking mass-spectrometry [1966]. In-cell cryo-ET structure determination is clearly better suited to large, abundant macromolecular complexes above ~500 kDa like ribosomes and proteasomes, this limit being reportedly set by the cell slices thickness [1945,1964,1967].

The number of reports using the techniques discussed above has grown exponentially in the last 5 years. They have already provided descriptions of fundamental cellular processes and macromolecular assemblies, and provided important data on macromolecular conformational equilibria and dynamics. NMR spectroscopy is certainly not likely to solve intracellular structures of high-molecular weight assemblies, or determine nanometer-scale distances (such measurements are accessible using paramagnetic relaxation enhancement and pseudo-contact shifts, but these are not commonly useable in cells as yet). Nonetheless, NMR has complementary capabilities. Among others, these include: i) solution- and solid-state NMR can deliver atomic-scale information on small and large molecules at the same time; ii) solution NMR can also deliver this atomic-scale information on very flexible regions; iii) solid-state NMR can also deliver this atomic-scale information on the complete ensemble of cellular molecules of interest (targeted and observed via isotopic labeling); iv) solution- and solid-state NMR do not require fluorescent dyes or paramagnetic labels to provide a structural information; v) solution NMR can monitor structural evolution in living cells and in real-time. Ideally, one should keep in mind the actual weak points of NMR (the necessity for high concentrations, the absence of information on subcellular localization and the possibility that only a subpopulation of the molecule of interest gives rise to the signal) and countervail them with one of the potentially complementary techniques cited above.

### 4.1.2. Live-cell enzymatic activities and drug binding

A tremendous number of biosensors have been designed to report on enzyme activities or molecular interactions/proximities using various bioluminescence and fluorescence phenomena, notably FRET and Bioluminescence Resonance Energy Transfer (BRET) [1968-1971]. It need hardly be said that such light-based approaches are more sensitive and better suited to high-throughput drug screening than is NMR. Moreover, they can often give real-time, spatially resolved information, sometimes even in living animals. Many commercial kits exist requiring minimal equipment for quantifying the cell signaling activities being studied. However, all these methods use either i) competition assays detecting the binding of generic fluorescent probes to the protein of interest [1972,1973], or ii) downstream/secondary reactions triggered by the activity or the proximity of the protein of interest, which can bias the interpretation of the results [1968-1971,1974-1976]. In the context of drug screening, this is of considerable concern because the targeted signaling proteins are often pleiotropic. Consequently, they are prone to generate unreliable downstream outcomes upon activation or inhibition: i) the drug target can have multiple activities and a binding compound may impact only one of these, ii) feedback loops are often activated. Deeper discussions about so-called "biased signaling" have been published elsewhere [1977,1978]. The diversity of reporters is already large, and one can
assume that it will become broader and that multiplexing capabilities will be improved [1979].

High-throughput screening methods using mass-spectrometry (MS) analysis can draw the interaction maps of large compounds libraries at the proteome scale in cells or cell extracts [1980-1982], or identify drug targets even in tissues [1373,1983,1984]. These MS studies do not report direct target binding, but rely on either i) compound functionalization and covalent attachment to their target [1985-1987], or ii) the consequences of drug binding on proteins thermal stabilities or sensitivity to protease treatments [1371]. The thermal stability shifts measured in the presence of ligands are not always sensitive enough to detect weak binders, and such assays are not always able to distinguish between binding to active and unactive forms of the protein target. MS analysis moreover is a destructive method, which does not allow the longitudinal monitoring of a single sample.

In comparison, in-cell NMR drug studies have only low sensitivity and low throughput capabilities, but they deliver direct information on drug:target interactions at the atomic scale: i) The protein target or the screened compounds can be observed directly by NMR, and ii) enzymatic activities can be monitored using the NMR signals of substrates and of their products. In this last case, tailored reporters can be designed to provide NMR exploitable signals, which allows NMR drug:protein studies a great deal of flexibility and adaptability.

## 4.2. Where to go – what to improve

Prediction is a fool's game, and we do not claim in any way to be able to mentor the community. We thought nevertheless that it would be worthwhile to mention a few directions that we consider to be important and promising for in-cell NMR. We cannot be comprehensive, of course: NMR spectroscopy is far too versatile to guess all its future applications.

### 4.2.1. Integrating advanced methods

We feel that combining some recent biotechnologies would make in-cell NMR investigations more attractive. We limit our list to a handful of techniques. First, in-cell NMR approaches could be applied to the so-called organoids and organs-on-a-chip, which the literature suggests require supplementary readouts [1988-1992]. A significant proportion of the studies described in the present review have been carried out on living tissues or cultured cells to prepare *in vivo* imaging and metabolic analysis (MRI/MRS) applications. They were often not intended to provide information of great biological relevance, but rather to show the feasibility of such NMR studies on live material. However, many of the approaches described could now find direct applications in longitudinal studies of organoids: NMR spectroscopy is capable of reporting in real time and non-invasively on the organoid's metabolism, drug penetration, its cellular processing or its binding to cellular targets, or its toxic effects, for example. Second, a number of NMR studies on eukaryotic cells would be facilitated by expert handling of the inducible expression or degradation of proteins in eukaryotic cells. That would permit carrying out experiments in cleaner conditions, with and without the nucleic acid or the protein of interest, in closely similar cellular backgrounds. This would be helpful for several types of investigations, from metabolism monitoring, to drug pharmacology, drug binding assays, or protein:protein interactions studies. Among other applications, it could also help in structural biological studies to achieve specific isotope labeling *in situ*: by triggering the expression of the protein of interest concomitantly with the supplementation of the chosen isotopically labeled amino acids, one could limit the NMR signal from the cellular background. Third, integrating genetic code expansion techniques [1993-1995] could permit one to incorporate paramagnetic tags or chemical

functions that would provide high NMR sensitivity, and which could help to report on ligand binding or protein conformational changes, as recently exemplified in *in vitro* studies [1996]. CRISPR-Cas9 genome editing techniques may also be useful to control the expression of target proteins, which would be of particular interest for in/on-cell drug binding assays and structural biology. However, for most NMR labs, it would probably be as efficient to rely on the many commercial stable cell lines or the help of dedicated biology labs.

Beyond this, one could think about advanced NMR methods that could profitably be implemented for in-cell studies. Here again, we can only include a few thoughts in a list that is necessarily far from comprehensive. In no particular order: applying pure-shift methods [1646,1649,1690,1691] or implementing post-acquisition data treatment [1741], notably deep-learning analysis [1997,1998], would certainly help to exploit NMR spectra from cellular samples; continuous readout using time-resolved non-uniform sampling could probably become a standard approach, as mentioned in Section 3.3.5. [1061,1066,1694-1696]; recent DNP-agents showing improved persistence in water and signal-enhancement at high magnetic fields will certainly help in/on-cell solid-state NMR approaches [1999-2002]. Methods developed for other spectroscopies might usefully be adapted also. Flash-freezing methods have been developed to improve the vitrification quality of cryo-EM/ET samples and to avoid the dissociation or denaturation of macromolecular complexes [2003,2004]: the slow freezing conditions used in solid-state NMR studies may bias observations and adapting fast freezing techniques may help to capture and analyze the actual cellular state at a precise timepoint. In-cell NMR protein observation will benefit from supplementary schemes of amino acid isotope labeling and combinations thereof, notably to generate NMR spectra with isolated signals and less noise from cellular background [1173,1211,1280,1392,1861] (see also Sections 2.2.2 or 2.5.2 for cell-wall analysis): protocols might be adapted from quantitative proteomics using stable-isotope labeling with amino acids in cell culture (SILAC) [2005-2007]. We can also suggest the future promise of nitrogen-vacancy (NV) centers in diamond and optically detected magnetic resonance: large species with highly restricted motion in cells could favor nano-NMR, but the expertise and the actual experimental designs are not yet sufficiently accessible to allow their application to in-cell studies [2008-2011].

Finally, a few technical aspects should probably be tackled in the future to make in-cell NMR more appealing. First, we already enjoy using commercial cryoprobes with up to five channels (¹H, ²H, ¹³C, ¹⁵N, ¹⁹F/³¹P) for studies using samples in conventional 5 mm NMR tubes. As shown in Section 2.1., a broad diversity of nuclei is observable by NMR, and their parallel monitoring would be highly valuable in a number of other situations. Let us imagine a study testing a drug candidate against high blood-pressure on a mini-brain or a mini-heart: biologists would doubtless appreciate a continuous readout of the levels of metabolites (¹H and ¹³C/¹⁵N), of ³¹P-phosphate-containing species and of ²³Na or ³⁹K, for example. Hence, we feel that a greater number of RF channels for different nuclei (possibly including tunable broad-band channels) would be highly desirable. Second, solution- and solid-state NMR approaches should be more often combined, which would bring about a more complete representation of the studied systems. Solution NMR can provide information only on fast-tumbling species, but it does so non-invasively in their native state and in real-time; solid-state NMR can report on the complete ensemble of labeled molecules, but only on frozen cells where dynamics cannot be characterized and where conformations and interactions can be influenced by the freezing techniques. Studying in-cell samples might well promote productive collaborations between labs with expertise in solution- and solid-state NMR.
Third, the field should work to establish common standards to validate cell viability and ensure better reproducibility of acquisition conditions. Unfortunately, these requirements will probably necessitate using bioreactors and probes permitting $ ^{31}P $-acquisition to monitor cellular phosphate-containing species, both of which would further stretch budgets. Finally, as explained in Section 3.4.3., NMR spectroscopy has remarkable capabilities in defining weak interactions ($ K_d \sim 10^{-7} \sim 10^{-3} $ mol/L, $ k_{off} \sim 10^2 \sim 10^6 $ s$^{-1}$ using STD-NMR or WaterLOGSY for example [1379,1412]), but approved drugs on the market generally show much slower off-rates (1 to 0.01 min$^{-1}$, see for example [1743,1744]). To be able to participate in the drug research pipeline, it would be desirable to establish in-cell NMR methods that apply to these slower types of binding kinetics.

### 4.2.2. Specific strengths of NMR that complement other techniques

Once more, the versatility of NMR spectroscopy is too broad to include all the possible subjects that could be tackled on cellular samples. In more concrete terms than in Section 1.2., we want here to put together some of the specific capabilities (and shortcomings) of NMR relevant for in-cell studies.

NMR spectroscopy permits the direct, atomic-scale observation of the studied object itself, rather than of a bound fluorophore or of a reaction product. NMR spectroscopy can report simultaneously on a broad variety of nuclei and of molecules, from ions and metabolites to proteins and membranes, in an isotope-selective fashion. NMR spectroscopy can trace the conformational and chemical environment of nuclei non-invasively and in real-time, and it allows the application of light and of chemical additives.

As far as in-cell studies are concerned, the main drawbacks of NMR are i) its poor sensitivity, ii) its inability to localize the detected species at submicrometer scale, iii) its slow measurement speed and consequent low-throughput. Also, practitioners of NMR spectroscopy require long years of training, and spectroscopists are most often specialized either in small or in large molecules, either in the solution- or in the solid-state.

Altogether, there are particular fields where in-cell NMR can be highly valuable: e.g. the monitoring of redox states, of metal chelation, of light-induced reactions, the characterization of flexible and/or middle-size (1-50 kDa) objects for which cryo-ET is so far "blind", studies where the simultaneous observation of small and large molecular species is required (such as an enzyme and its substrates/products/cofactors, or a ligand and the conformational state of its protein target), the longitudinal monitoring of large panels of metabolites or of protein targets in cells/organs/small animals upon drug exposure, etc. In-cell NMR will inevitably benefit from complementary techniques, especially from microscopies providing information on subcellular localization. By providing experimental evidence on virtually any chosen molecule in live material, in-cell NMR can complement high-throughput methods in the drug research pipeline, in a world where large-scale cell-based or virtual screenings have gained interest and trust (see for example [2012-2015]). Indeed, in-cell NMR delivers information at the atomic scale, which will always be invaluable.

## Declaration of Competing Interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

## Acknowledgements

This work was supported by the CNRS and the CEA-Saclay, and by iNEXT-Discovery, Grant No. 871037, funded by the Horizon 2020 research and innovation programme of the European Com-

mission the French Infrastructure for Integrated Structural Biology (https://frisbi.eu/, grant number ANR-10-INSB-05-01, Acronym FRISBI) and by the French National Research Agency (ANR; research grants ANR-14-ACHN-0015 and ANR-20-CE92-0013), and by the Fondazione Cassa di Risparmio di Firenze (grant CRF2020.1395). We also thank Pierre Barraud and Muriel Dele-pierre for their helpful reading and comments.

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