# In-cell NMR: Why and how?

François-Xavier Theillet, Enrico Luchinat

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Francois-Xavier Theillet, Enrico Luchinat. In-cell NMR: Why and how?. Progress in Nuclear Magnetic Resonance Spectroscopy, 2022, 132-133, pp.1 - 112. 10.1016/j.pnmrs.2022.04.002. hal-03787648

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# In-cell NMR: Why and how?

Francois-Xavier Theillet<sup>a,*</sup>, Enrico Luchinat<sup>b,c</sup>

Edited by David Neuhaus and David Gadian

<figure><img src="image_4.png" /></figure>

## ARTICLE INFO

**Article history:**

Received 15 September 2021

Accepted 27 April 2022

Available online 04 May 2022

**Keywords:**

Structural biology

Biophysics

Metabolomics

Drug research

Cellular environment

## ABSTRACT

NMR spectroscopy has been applied to cells and tissues analysis since its beginnings, as early as 1950. We have attempted to gather here in a didactic fashion the broad diversity of data and ideas that emerged from NMR investigations on living cells. Covering a large proportion of the periodic table, NMR spectroscopy permits scrutiny of a great variety of atomic nuclei in all living organisms non-invasively. It has thus provided quantitative information on cellular atoms and their chemical environment, dynamics, or interactions. We will show that NMR studies have generated valuable knowledge on a vast array of cellular molecules and events, from water, salts, metabolites, cell walls, proteins, nucleic acids, drugs and drug targets, to pH, redox equilibria and chemical reactions. The characterization of such a multitude of objects at the atomic scale has thus shaped our mental representation of cellular life at multiple levels, together with major techniques like mass-spectrometry or microscopies.

NMR studies on cells has accompanied the developments of MRI and metabolomics, and various subfields have flourished, coined with appealing names: fluxomics, foodomics, MRI and MRS (i.e. imaging and localized spectroscopy of living tissues, respectively), whole-cell NMR, on-cell ligand-based NMR, systems NMR, cellular structural biology, in-cell NMR. All these have not grown separately, but rather by reinforcing each other like a braided trunk. Hence, we try here to provide an analytical account of a large ensemble of intricately linked approaches, whose integration has been and will be key to their success.

We present extensive overviews, firstly on the various types of information provided by NMR in a cellular environment (the “why”, oriented towards a broad readership), and secondly on the employed NMR techniques and setups (the “how”, where we discuss the past, current and future methods). Each subsection is constructed as a historical anthology, showing how the intrinsic properties of NMR spectroscopy and its developments structured the accessible knowledge on cellular phenomena. Using this systematic approach, we sought i) to make this review accessible to the broadest audience and ii) to highlight some early techniques that may find renewed interest. Finally, we present a brief discussion on what may be potential and desirable developments in the context of integrative studies in biology.

© 2022 The Authors. Published by Elsevier B.V. This is an open access article under the CC BY license (http://creativecommons.org/licenses/by/4.0/).

## Contents

<table><tr><td>1. Introduction</td><td>3</td></tr><tr><td>1.1. Definitions and scope</td><td>3</td></tr><tr><td>1.2. The basic principles of in-cell NMR</td><td>3</td></tr><tr><td>1.2.1. Observing a broad range of phenomena</td><td>3</td></tr><tr><td>1.2.2. Achieving selective detection from the cellular background</td><td>3</td></tr><tr><td>1.3. Which nuclei?</td><td>4</td></tr><tr><td>2. Applications of NMR spectroscopy on living cells.</td><td>5</td></tr></table>

<footnote><sup>a</sup>Université Paris-Saclay, CEA, CNRS, Institute for Integrative Biology of the Cell (I2BC), 91198 Gif-sur-Yvette, France</footnote>

<footnote><sup>b</sup>Dipartimento di Scienze e Tecnologie Agro-Alimentari, Alma Mater Studiorum – Università di Bologna, Piazza Goidanich 60, 47521 Cesena, Italy</footnote>

<footnote><sup>c</sup>CERM – Magnetic Resonance Center, and Neurofarba Department, Università degli Studi di Firenze, 50019 Sesto Fiorentino, Italy</footnote>

<footnote>* Corresponding author at: CEA-Saclay, bât. 144, 91191 Gif-sur-Yvette, France.</footnote>

<footnote>E-mail address: francois-xavier.theillet@cnrs.fr (F.-X. Theillet).</footnote>
<table><tbody><tr><td>2.1. Cellular solvent</td><td>5</td></tr><tr><td>2.1.1. Water: intracellular and transmembrane diffusion, interactions and correlation times</td><td>5</td></tr><tr><td>2.1.2. Quantification and interactions of metal ions:</td><td>7</td></tr><tr><td>2.1.3. Li<sup>+</sup></td><td>7</td></tr><tr><td>2.1.4. Na<sup>+</sup></td><td>9</td></tr><tr><td>2.1.5. K<sup>+</sup>, and Rb<sup>+</sup> as a surrogate</td><td>10</td></tr><tr><td>2.1.6. Mg<sup>2+</sup></td><td>11</td></tr><tr><td>2.1.7. Ca<sup>2+</sup></td><td>12</td></tr><tr><td>2.1.8. Transition metal ions Mn<sup>2+</sup>, Fe<sup>2+</sup>/Fe<sup>3+</sup>, Co<sup>2+</sup>, Ni<sup>2+</sup>, Cu<sup>+</sup>/Cu<sup>2+</sup> and Zn<sup>2+</sup>, Pb<sup>2+</sup></td><td>13</td></tr><tr><td>2.1.9. More highly toxic species: Al<sup>3+</sup>, Cd<sup>2+</sup> and Cs<sup>+</sup></td><td>13</td></tr><tr><td>2.1.10. Common anions</td><td>14</td></tr><tr><td>2.1.11. Viscosity, pH, redox potential</td><td>14</td></tr><tr><td>2.2. Cell wall composition and dynamics</td><td>15</td></tr><tr><td>2.2.1. Plant cell walls</td><td>15</td></tr><tr><td>2.2.2. Peptidoglycans</td><td>16</td></tr><tr><td>2.2.3. Lipopolysaccharides and beta-glucans</td><td>18</td></tr><tr><td>2.2.4. Lipid dynamics in membranes</td><td>19</td></tr><tr><td>2.3. Metabolic activity</td><td>21</td></tr><tr><td>2.3.1. The early days: studying carbon fluxes using <sup>13</sup>C-NMR.</td><td>21</td></tr><tr><td>2.3.2. <sup>31</sup>P-NMR of phospho-species, energy fluxes and cellular localization.</td><td>22</td></tr><tr><td>2.3.3. Nitrogen flux studied by <sup>15</sup>N-NMR and <sup>14</sup>N-NMR</td><td>23</td></tr><tr><td>2.3.4. Overcoming spectral congestion in <sup>1</sup>H-NMR spectroscopy for higher sensitivity: monitoring cellular species below millimolar concentrations in the timescale of minutes.</td><td>24</td></tr><tr><td>2.3.5. HR-MAS <sup>1</sup>H-NMR spectroscopy: lipid filtering/editing and metabolome of intact cells, tissues and small living organisms.</td><td>25</td></tr><tr><td>2.3.6. 2D <sup>1</sup>H/<sup>13</sup>C-correlation NMR of live cells and small animals: metabolites at submillimolar concentrations on time-scales of minutes to hours</td><td>27</td></tr><tr><td>2.3.7. <sup>13</sup>C-hyperpolarization for carbon-flux quantification at low-millimolar concentrations and time scales of seconds</td><td>29</td></tr><tr><td>2.4. Cellular structural biology</td><td>31</td></tr><tr><td>2.4.1. Solving structures of folded, soluble proteins in cells</td><td>32</td></tr><tr><td>2.4.2. Structural studies of membrane and cytoplasmic proteins using solid-state NMR</td><td>33</td></tr><tr><td>2.4.3. Disordered proteins: intracellular conformations, dynamics and interactions.</td><td>34</td></tr><tr><td>2.4.4. Protein maturation: metal binding, post-translational modifications and redox regulation</td><td>36</td></tr><tr><td>2.4.5. Nucleic acids.</td><td>37</td></tr><tr><td>2.4.6. Protein folding, quinary structure and large assemblies</td><td>38</td></tr><tr><td>2.5. Cellular drug assays</td><td>40</td></tr><tr><td>2.5.1. In-cell drug screening through binding.</td><td>40</td></tr><tr><td>2.5.2. Drug modes of action and cell penetrance</td><td>42</td></tr><tr><td>2.6. Epitope mapping, structure-activity relationship</td><td>43</td></tr><tr><td>2.6.1. Virus receptors</td><td>43</td></tr><tr><td>2.6.2. Mammalian membrane proteins</td><td>43</td></tr><tr><td>3. In-cell NMR theoretical and practical considerations</td><td>44</td></tr><tr><td>3.1. A historical aside: the beginnings of NMR of biological material</td><td>44</td></tr><tr><td>3.1.1. Early controversies in NMR studies of water (1950-70's), and what can be learned from them</td><td>44</td></tr><tr><td>3.1.2. The case of quadrupolar nuclei</td><td>45</td></tr><tr><td>3.2. Line-broadening and observability.</td><td>46</td></tr><tr><td>3.2.1. Intra- and extra-cellular mobilities of molecules.</td><td>46</td></tr><tr><td>3.2.2. Inhomogeneous magnetic susceptibility.</td><td>47</td></tr><tr><td>3.2.3. Multiple interactions. A warning concerning quantification and referencing.</td><td>50</td></tr><tr><td>3.3. Observation of ions and metabolites</td><td>52</td></tr><tr><td>3.3.1. Metal ions.</td><td>52</td></tr><tr><td>3.3.2. Spin-echo as an editing strategy</td><td>52</td></tr><tr><td>3.3.3. T2, T1-rho filters and other filtering/editing methods for signals from mobile species.</td><td>52</td></tr><tr><td>3.3.4. High-Resolution Magic-Angle Spinning (HR-MAS) to alleviate sample heterogeneity: advantages, drawbacks and late developments</td><td>54</td></tr><tr><td>3.3.5. Other solutions to sample heterogeneity, molecular diffusion heterogeneity and sample evolution.</td><td>55</td></tr><tr><td>3.3.6. Hyperpolarized metabolites for live-cell studies: 10,000 times more signal, but how many constraints?</td><td>56</td></tr><tr><td>3.4. Observation of ligands</td><td>58</td></tr><tr><td>3.4.1. Technical recommendations for ligand observed NMR in presence of cells.</td><td>58</td></tr><tr><td>3.4.2. Technical recommendations for STD experiments in presence of cells.</td><td>58</td></tr><tr><td>3.4.3. On the use and present limits of on-cell NMR and the use of STD/TRNOE.</td><td>59</td></tr><tr><td>3.5. Cellular structural biology</td><td>60</td></tr><tr><td>3.5.1. How relevant is the intracellular concentration of the studied species?</td><td>60</td></tr><tr><td>3.5.2. Sample preparation</td><td>60</td></tr><tr><td>3.5.3. Protein structure determination</td><td>60</td></tr><tr><td>3.5.4. Protein:protein interactions and protein:ligand binding</td><td>61</td></tr><tr><td>3.5.5. In-cell solid-state NMR and the road to DNP-enhanced in-cell NMR</td><td>61</td></tr><tr><td>3.5.6. Nucleic acids.</td><td>62</td></tr><tr><td>3.6. Whole cell analysis</td><td>62</td></tr><tr><td>3.6.1. Solid-state NMR analysis of cellular content, molecular proximities and chemical bonds</td><td>62</td></tr><tr><td>3.6.2. The case of hyperpolarized <sup>129</sup>Xe, a multitask MRI-oriented reporter</td><td>63</td></tr><tr><td>3.7. The quest for a bioreactor.</td><td>63</td></tr><tr><td>3.7.1. A re-emerging necessity.</td><td>63</td></tr></tbody></table>
<table><tbody><tr><td>3.7.2.</td><td>Cell immobilization strategies</td><td>63</td></tr><tr><td>3.7.3.</td><td>Particular problems for consideration.</td><td>64</td></tr><tr><td>3.7.4.</td><td>Past and recent applications</td><td>65</td></tr><tr><td>4.</td><td>Where next?</td><td>65</td></tr><tr><td>4.1.</td><td>Comparison and complementarity with other spectroscopies, spectrometries and microscopies</td><td>66</td></tr><tr><td>4.1.1.</td><td>In-cell structural biology</td><td>66</td></tr><tr><td>4.1.2.</td><td>Live-cell enzymatic activities and drug binding.</td><td>66</td></tr><tr><td>4.2.</td><td>Where to go - what to improve.</td><td>67</td></tr><tr><td>4.2.1.</td><td>Integrating advanced methods</td><td>67</td></tr><tr><td>4.2.2.</td><td>Specific strengths of NMR that complement other techniques</td><td>68</td></tr><tr><td colspan="2">Declaration of Competing Interest</td><td>68</td></tr><tr><td colspan="2">Acknowledgements</td><td>68</td></tr><tr><td colspan="2">References</td><td>68</td></tr></tbody></table>

# 1. Introduction

## 1.1. Definitions and scope

The initial intention for this review was to focus on the modern concept of in-cell NMR, i.e. NMR studies of macromolecules in cellular milieus, as coined by Dötsch and his coworkers in 2001 [1]. It has been 10 years since the last review on this topic in this journal [2,3]. A significant body of results has been published in the meantime, but also many reviews [4-24] (see also the excellent book recently edited by Ito, Dötsch and Shirakawa [25]). Events in 2020-21 (particularly the COVID-19 pandemic and the “Great Lockdown”) gave us time in which to extend background research to include literature in “neighboring” NMR fields, and also publications from the early days of NMR spectroscopy. Hence, we identified consistent themes and patterns running through the very many publications from the overall field of NMR studies on cells. Such a continuity has been stressed in an inspiring fashion by Lip pens [15] (and earlier by Szwergold [26] or Dötsch [1], or more recently by Selenko [22]), but has not been explored extensively in a review. We thought it would be a good time to attempt to bring together an account of this long-term body of knowledge: this would help us to propose a perspective after 70 years of works in the broad, integrated field of in-cell NMR spectroscopy, a field encompassing NMR studies using intact cells.

Our aim here is thus to cover NMR studies analyzing cells and, when closely related, “in situ” studies on cell extracts, membranes, virus and viral particles, organelles, dissected tissues or small model animals such as flies or worms. NMR spectroscopy permits the analysis of living cells or tissues and their content in a non-invasive fashion. It has applications in a very broad variety of fields, e.g. metabolomics, MRI, cellular biophysics, structural biology, drug development, etc. We will only mention briefly MRI and metabolomics, which represent an immense, independent literature [27] (see [28-32] for a few selected applications in clinics or cognitive science, see [33-35] for MRS metabolic imaging and some future prospects [36,37], see [38-46] for metabolomics and some future NMR metabolomics strategies [47,48]).

Our overall goal was to attempt to integrate and analyze the wide landscape of past and present NMR studies on cells. We have adopted a chronological narrative in most sections to present the progressive technical achievements and the observations they gave access to. We felt such a presentation could be helpful for non-NMR readers and novice NMR students, as well as perhaps for more experienced spectroscopists.

We have attempted to present each section such that it can be read independently. We have deliberately avoided equations, but have tried to provide experimental numerical data wherever possible. We hope that this approach to presentation will be stimulating both for a broad non-expert readership and for NMR spectroscopists.

## 1.2. The basic principles of in-cell NMR

### 1.2.1. Observing a broad range of phenomena

The appealing concept of in-cell NMR is to obtain information at the atomic scale on chosen nuclear isotopes in living environments. Indeed, NMR spectroscopy reports on the impact of the local chemical environment, conformation or interaction dynamics on the detected populations of nuclei.

These nuclei can be incorporated in molecules of any size, and can become sensors for more global physical properties like pH or viscosity. NMR spectroscopy can also provide information on local homogeneity and dynamics, from the picosecond to the second timescales by measuring magnetization relaxation times. However, these can be difficult to interpret, because they are dependent on rotational correlation times, conformational dynamics and interaction kinetics at the same time [49].

In addition, because NMR is a non-destructive technique, time series of successive spectra can be recorded over minutes to days on a single sample, provided that conditions to maintain its integrity are met. Chemical reactions or environmental changes can thus be monitored through widely ranging experimental timescales.

Hence, in-cell NMR can provide a broad range of information at multiple scales. Of course, several limitations exist that we will explore later. Still, we can mention the two we see as the most important: i) “NMR-visible” molecular species are limited to those that do not interact or interact very weakly (i.e. rapid exchange below the µs time scale and low populations in the order of magnitude of 1 %) with large macromolecular assemblies (in the case of solution NMR) or those that do not interact with too many cellular binding partners (in the cases of either solution or solid-state NMR); ii) standard NMR techniques cannot determine the spatial localization of the detected molecules at the micrometer scale and below, i.e. the subcellular scale. A subcellular position can thus be inferred only from secondary pieces of evidence, in those cases where organelles/compartments contain markedly distinct chemical environments (e.g. pH, redox potential, presence of binding partners, etc.) that result in well-defined spectral changes.

### 1.2.2. Achieving selective detection from the cellular background

NMR spectroscopy of cellular species permits the selective observation of specific nuclei (more precisely the precession of their nuclear magnetic moments) in the cellular milieu. Three main factors can make a particular population of nuclei distinguishable from others in the same sample: i) among nuclei, each nuclear species (isotope) generates magnetic resonances in a relatively narrow and precise range of frequencies due to its different gyromagnetic ratio $γ$ ($ω_0=-γB_0$ [49] defining the precession frequency $ω_0$ of a nuclear spin having a gyromagnetic ratio $γ$ in a magnetic field $B_0$); ii) among populations of the same isotope, nuclei having the same chemical environment will contribute signals at a very specific
Glossary

<table><tr><td>ADP</td><td>Adenosine DiPhosphate</td></tr><tr><td>AMP</td><td>Adenosine MonoPhosphate</td></tr><tr><td>α-Syn</td><td>α-Synuclein</td></tr><tr><td>ATP</td><td>Adenosine TriPhosphate</td></tr><tr><td>BIRD</td><td>Bi-linear Rotation Decoupling</td></tr><tr><td>BOLD</td><td>Blood Oxygenation Level Dependent</td></tr><tr><td>cAMP</td><td>Cyclic Adenosine MonoPhosphate</td></tr><tr><td>CEST</td><td>Chemical Exchange Saturation Transfer.</td></tr><tr><td>CP-MAS</td><td>Cross-Polarization Magic Angle Spinning</td></tr><tr><td>CPMG</td><td>Carr-Purcell-Meiboom-Gill (pulse sequence)</td></tr><tr><td>CPP</td><td>Cell-Penetrating Peptide</td></tr><tr><td>CRINEPT</td><td>Cross Relaxation-enhanced INEPT</td></tr><tr><td>CSA</td><td>Chemical Shift Anisotropy</td></tr><tr><td>cryo-EM</td><td>Cryogenic Electron Microscopy</td></tr><tr><td>cryo-ET</td><td>Cryogenic Electron Tomography</td></tr><tr><td>DCP</td><td>Double Cross-Polarization</td></tr><tr><td>dDNP</td><td>dissolution Dynamic Nuclear Polarization</td></tr><tr><td>DNA</td><td>DeoxyriboNucleic Acid</td></tr><tr><td>DNP</td><td>Dynamic Nuclear Polarization</td></tr><tr><td>DOTA</td><td>1,4,7,10-tetraazacyclododecane-1,4,7,10-tetraacetic acid</td></tr><tr><td>DP</td><td>Direct Polarization</td></tr><tr><td>DPG</td><td>DiPhosphoGlycerate</td></tr><tr><td>DQF/TQF</td><td>Double quantum filter / Triple quantum Filter</td></tr><tr><td>ECM</td><td>ExtraCellular Matrix</td></tr><tr><td>EDTA</td><td>EthyleneDiamineTetraacetic Acid</td></tr><tr><td>EPR</td><td>Electron Paramagnetic Resonance</td></tr><tr><td>FID</td><td>Free Induction Decay</td></tr><tr><td>FLIM</td><td>Fluorescence Lifetime Imaging Microscopy</td></tr><tr><td>FRET</td><td>Förster or Fluorescence Resonance Energy Transfer</td></tr><tr><td>FT</td><td>Fourier Transform</td></tr><tr><td>GFP</td><td>Green Fluorescence Protein</td></tr><tr><td>GPCR</td><td>G-Protein-Coupled Receptor</td></tr><tr><td>GSH</td><td>Glutathione</td></tr><tr><td>HMQC</td><td>Heteronuclear Multiple Quantum Coherence</td></tr><tr><td>Ht</td><td>Haematocrit</td></tr><tr><td>HR-MAS</td><td>High-Resolution Magic Angle Spinning</td></tr><tr><td>HSQC</td><td>Heteronuclear Single Quantum Coherence</td></tr><tr><td>IDP</td><td>Intrinsically Disordered Protein</td></tr><tr><td>IDR</td><td>Intrinsically Disordered Region (of a protein)</td></tr><tr><td>INEPT</td><td>Insensitive Nuclei Enhanced by Polarization Transfer</td></tr><tr><td>MD</td><td>Molecular Dynamics</td></tr><tr><td>NAD+</td><td>Nicotinamide Adenine Dinucleotide+</td></tr><tr><td>NADP+</td><td>Nicotinamide Adenine Dinucleotide Phosphate</td></tr><tr><td>NMR</td><td>Nuclear Magnetic Resonance</td></tr></table>

<table><tr><td>MAS</td><td>Magic Angle Spinning</td></tr><tr><td>MRI</td><td>Magnetic Resonance Imaging</td></tr><tr><td>mRNA</td><td>messenger RiboNucleic Acid</td></tr><tr><td>MRS</td><td>Magnetic Resonance Spectroscopy (in vivo, spatially localized biochemical analysis)</td></tr><tr><td>NOE</td><td>Nuclear Overhauser Effect</td></tr><tr><td>NOESY</td><td>Nuclear Overhauser Effect Spectroscopy</td></tr><tr><td>NUS</td><td>Non-Uniform Sampling</td></tr><tr><td>PCS</td><td>Pseudo-Contact Shift</td></tr><tr><td>PFG</td><td>Pulsed Field Gradient</td></tr><tr><td>PFT</td><td>Pore-Forming Toxin</td></tr><tr><td>PG</td><td>PeptidoGlycan</td></tr><tr><td>PHIP</td><td>ParaHydrogen Induced Polarization.</td></tr><tr><td>ppm</td><td>parts per billion</td></tr><tr><td>ppm</td><td>parts per million</td></tr><tr><td>PRE</td><td>Paramagnetic Relaxation Enhancement</td></tr><tr><td>PTM</td><td>Post-Translational Modification.</td></tr><tr><td>RBC</td><td>Red Blood Cell</td></tr><tr><td>RDC</td><td>Residual Dipolar Coupling</td></tr><tr><td>REDOR</td><td>Rotational-Echo DOuble-Resonance</td></tr><tr><td>RNA</td><td>RiboNucleic Acid</td></tr><tr><td>rRNA</td><td>ribosomal RiboNucleic Acid</td></tr><tr><td>SLO</td><td>Streptolysin O</td></tr><tr><td>smFRET</td><td>Single-molecule FRET</td></tr><tr><td>SOD1</td><td>SuperOxide Dismutase 1</td></tr><tr><td>SOFAST-HMQC</td><td>Selective Optimized Flip-Angle Short-Transient Heteronuclear Multiple Quantum Coherence</td></tr><tr><td>SNR</td><td>Signal-to-Noise Ratio</td></tr><tr><td>ssNMR</td><td>solid-state Nuclear Magnetic Resonance</td></tr><tr><td>ssRNA</td><td>single-strand RiboNucleic Acid</td></tr><tr><td>STD</td><td>Saturation Transfer Difference</td></tr><tr><td>STDD</td><td>Saturation Transfer Double Difference</td></tr><tr><td>τ<sub>c</sub></td><td>Correlation time for overall (isotropic) tumbling of the protein</td></tr><tr><td>TANGO</td><td>Testing for Adjacent Nuclei with a Gyration Operator</td></tr><tr><td>TRNOE</td><td>TTransferred NOE</td></tr><tr><td>TR-NUS</td><td>Time-Resolved Non-Uniform Sampling.</td></tr><tr><td>TROSY</td><td>Transverse Relaxation-Optimized Spectroscopy.</td></tr><tr><td>VAS</td><td>Variable Angle Spinning</td></tr><tr><td>WATERGATE</td><td>Water suppression by Gradient-Tailored Excitation</td></tr><tr><td>waterLOGSY</td><td>Water-Ligand Observed via Gradient Spectroscopy</td></tr><tr><td>WATERGATE</td><td>WATER suppression by GrAdient Tailored Excitation</td></tr><tr><td>WT</td><td>Wild-Type</td></tr><tr><td>WTA</td><td>Wall Teichoic Acid</td></tr></table>

fic resonance frequency, the so-called chemical shift ($\delta=(\omega-\omega_0)/\omega_0$); iii) among populations with similar chemical shifts, nuclei with relatively high abundance will stand out. Further distinctions can be exploited that are based on the selective manipulation of nuclei bound to a determined nuclear isotope (e.g. protons covalently bound to $^{15}$N or $^{13}$C).

NMR signal intensities are proportional to the number of nuclei resonating at the corresponding frequency. However, a large population does not guarantee its detectability: conformational and interaction dynamics, sample homogeneity, and other intrinsic properties of the observed nuclei have consequences on the signal linewidth, all of which are decisive for NMR signal intensity as we will often see below.

1.3. Which nuclei?

We must recall that only nuclei having a non-zero spin magnetic moment can be observed. This excludes, unfortunately, the

naturally predominant isotopes $^{12}$C, $^{16}$O and $^{32}$S, which are thus "NMR invisible". Other abundant isotopes like $^{14}$N have unfavorable nuclear spin quantum numbers ($I > 1/2$) that generate fast relaxation, hence very broad signals that are impractical to detect (except for very small, symmetric molecules like $NH_4^+$). NMR spectroscopists have circumvented and even transcended this difficulty by incorporating $^{13}$C- or $^{15}$N-labeled species in cells and then detecting those in a specific fashion. The stable isotopes $^{13}$C and $^{15}$N have low natural abundances (1.1% and 0.4%, respectively) and their NMR detection generates thus low background signal intensities from the endogenous material.

Besides null spins, we can classify nuclei in two categories. The most useful isotopes are spin $1/2$ nuclei, like $^{1}$H, $^{13}$C, $^{15}$N, $^{19}$F, $^{31}$P. These have manageable mono-exponential "transverse" relaxation in solution on timescales of the order of 1 to 100 milliseconds. This holds true for homogeneous pools of molecules tumbling in the 10 nanoseconds range or less, that do not experience interactions in the micro- to milli-second time scale. Indeed, such "intermediate
time scale" binding events can cause line broadening, which makes the signals more difficult to detect. Importantly, in the cellular context, detecting molecules involved in complexes larger than ~30 kDa often necessitates more advanced NMR strategies, whose success is not guaranteed.

Other isotopes are “quadrupolar” nuclei, like $ ^2H$, $ ^7Li$, $ ^{14}N$, $ ^{17}O$, $ ^{23}Na$, $ ^{25}Mg$, $ ^{27}Al$, $ ^{39}K$, $ ^{43}Ca$ and almost all metals with non-zero nuclear spin: their non-spherical distribution of nuclear charges generates a quadrupolar moment interacting with the surrounding electric field, which can cause a strong relaxation of nuclear magnetic resonances in the case of a non-symmetric local environment, as typically occurs when transient binding to macromolecules occurs in cells. There, such quadrupolar nuclei show fast, multi-exponential relaxations, whose rates depend on the quadrupole strengths. This severely limits their detectability. We will show that these quadrupolar nuclei can still be exploited, even though they sometimes gave rise to misinterpretations in early studies.

## 2. Applications of NMR spectroscopy on living cells.

Anfinsen, who established the deterministic relationship between amino acid sequence and three-dimensional structure (at least for small folded proteins), emphasized that "a protein molecule only makes stable, structural sense when it exists under conditions similar to those for which it was selected" [50]. This thought is clearly at the heart of many biophysicists' concerns: to what extent do molecules behave the same or differently in the test tube and in cells? There is no question about the importance of *in vitro* studies using purified material: they are key to exploring the properties and the manipulation of biological molecules. However, presenting a complete picture in molecular biophysics requires characterizing molecules and their interactions in their native environment. Let's push an open door: To fully understand molecular phenomena occurring in cells, one has to study them in cells. NMR has been instrumental in this task. Here, we discuss the various questions that NMR can tackle.

### 2.1. Cellular solvent

NMR spectroscopy offers the possibility to manipulate and detect selectively the chosen nuclear spins, because every isotope has a different gyromagnetic ratio (see Section 1.2.2.). A broad spectrum of analysis is thus achievable on the composition of the cellular solvent. NMR is familiar for studies focusing on metabolites and proteins, but it can provide information on more fundamental properties of cells: this starts with water of course, but a number of exotic nuclei can be studied, as well as pH or redox potentials, as we will show in the paragraphs below.

#### 2.1.1. Water: intracellular and transmembrane diffusion, interactions and correlation times

Water combines many advantages where NMR spectroscopy is concerned: it is a fast-tumbling, symmetric, abundant small molecule containing two (equivalent) protons. $ ^1H$ is the highest gyromagnetic ratio nucleus (except for the radioactive $ ^3H$), whose spin ½ is the simplest to manipulate and detect. All these parameters have beneficial consequences for NMR relaxation rates, simplicity of spin manipulation and sensitivity. The T1 and T2 relaxation rates of water are sensitive to the chemical environment, and are sufficiently slow to be measured accurately. They are also slow enough to be affected in a substantial and observable manner by many solutes. Hence, because of its convenient NMR behavior, detection of water signals has been the basis for some of the most successful applications of NMR, water-based MRI being a particularly clear example.

After a slow, error-prone start (see Section 3.1.1.), NMR of cellular water became useful in the following three main categories: measurements of i) transmembrane diffusion, ii) intracellular diffusion, iii) the correlation times of bulk water and of water molecules associated with cellular macromolecules.

The water permeability of erythrocytes has been thoroughly studied since the 1970's. T2 relaxation measurements have been carried out in the absence or presence of extracellular, impermeable paramagnetic agents (notably $Mn^{2+}$), which greatly enhance the extracellular T2 rate but not the intracellular one (similar to the contrast agents used in MRI). The global T2s of water and their differences in absence/presence of paramagnetic agents give access to transmembrane exchange kinetics (see the previous reviews for the relevant equations [51,52]). This was a good alternative to methods using radioactive tritiated water from the late 1950's. It permitted measurement of transmembrane water exchange of erythrocytes in varying osmolarity, temperature or pH conditions [53-59], upon exposure to chemicals [57,60], from patients with various illnesses [61,62] or from a great number of animals [63]. Hence, by the end of the 1980's, NMR had already made an important contribution to the field [51,64,65]. However, the extracellular paramagnetic species used at that time was mostly $Mn^{2+}$, which actually penetrates erythrocytes at high concentrations, hence diminishing the apparent water exchange rate and introducing discrepancies among the published reports [51,66]. MRI-related $Gd^{3+}$-chelators are more reliable in this regard [67].

Another approach was proposed, which did not require use of an external paramagnetic agent. It relied on the possibility to measure diffusion coefficients using Pulsed Field-Gradient (PFG) NMR, a method developed in the 1950-60's. The extensive presentation of its principles is out of our scope; a number of reviews are available [68-72]. Briefly, PFG NMR relies on i) the imposition of a spatial gradient of magnetic field for a short time, which causes a loss of spatial coherence of the NMR signals, ii) waiting for a defined delay, and iii) applying a second gradient pulse of opposite sign: hence, the coherence is fully recovered for immobile species, but only partially recovered for species that diffused in the delay inserted between the two gradient pulses (Fig. 1). The resulting observable NMR intensities are anti-correlated to the diffusion coefficients and to the time between the two gradient pulses. Thus, measuring these intensities gives access to NMR diffusometry. PFG NMR has been successful in measuring molecular displacements from 10 nm to 100 µm in a non-invasive fashion. This concept was used in the early 1970's to measure water diffusion in frog and rat muscles, which was found to be on average about half the rate seen in distilled water [73,74].

Because intracellular water diffusion is restricted by cell walls, long delays between the pulsed gradients cause a progressively attenuated signal decay. This would reach a plateau if the intracellular pool of water was trapped forever, since the cellular dimensions limit how far a water molecule can diffuse, but in reality, transmembrane exchange causes further decay, resulting in a measurable weighted average of the intra- and extra-cellular diffusion coefficients (Fig. 1). Theoretically, this effect allows estimation of intra- and extra-cellular self-diffusion coefficients of water, the cellular size defining the restricted space of diffusion, and the transmembrane water exchange rate [52,69]. It has been used by Andrasko in 1975 to measure simultaneously the transmembrane water exchange rate in erythrocytes and the average intracellular diffusion coefficient of water [82]. These measurements neglected the effects of magnetic susceptibility inhomogeneities in cell samples, which generate intrinsic field gradients and thus additional signal decay during PFG NMR experiments (These can themselves be used to measure intracellular diffusion coefficients of water in live and fixed cells [83,84]); the effects of such inhomogeneities on NMR signals are rather complicated and are treated in
<figure><img src="image_5.png" /><figcaption>**Fig. 1.** Measurement of molecular diffusion in presence of intra- and extra-cellular compartments. **A)** Sketch of a simple Pulsed Field Gradients Spin Echo (PFGSE) pulse sequence used to measure molecular translational diffusion by NMR, where the z-gradient pulses have a duration d, a power g and are separated by a delay Δ; **B)** Brief schematic explanation showing the z-dependent magnetization (red arrows) before and after the first pulsed field gradient, and showing the final magnetization, magnetization precession (free-induction decay, FID) and the signal intensities after Fourier Transform (FT); **C)** Scheme introducing the parameters to fit, D<sub>i</sub>/D<sub>e</sub>, the diffusion coefficients, P<sub>i</sub>/P<sub>e</sub>, the molecular populations, τ<sub>i</sub>/τ<sub>e</sub>, the average residency times, k<sub>i</sub>/k<sub>e</sub>, the transmembrane exchanges of intra- and extra-cellular molecules, respectively; In the case of free Brownian motion, the final acquired signal intensity is proportional to S(2τ)=M<sub>0</sub>exp(-2τ/T<sub>2</sub>)exp[-D<sub>i</sub>γ<sup>2</sup>(Δ-d/3)], with M<sub>0</sub> the equilibrium magnetization, T<sub>2</sub> the transverse relaxation time, D the diffusion coefficient and τ, Δ and d the PFGSE-defined parameters; a number of improved variants of this pulse sequence have been proposed [75-79], notably the Stimulated Echo sequences, where diffusion is affected by T1 and not T2 (T1 is often much longer and more favorable than T2 in cells and tissues); **D)** Theoretical graph of the PFGSE attenuated signal intensity of a solute diffusing in two compartments in exchange, in the model situation corresponding to packed cells or tissues, where P<sub>i</sub>>P<sub>e</sub> and D<sub>e</sub>>D<sub>i</sub>; (1)=exp(-Δ/2τ<sub>i</sub>); (2) = 1/(τ<sub>i</sub>P<sub>e</sub>D<sub>e</sub>); (3) = exp(-Δ/τ<sub>i</sub>); (4) = exp(-Δ/τ<sub>i</sub>(1+D<sub>i</sub>/P<sub>e</sub>D<sub>e</sub>)); slope1 = exp(-Δ/τ<sub>i</sub>)*exp(-KΔ(P<sub>e</sub>D<sub>e</sub>+D<sub>i</sub>)); slope2 = exp(-KΔ(P<sub>e</sub>D<sub>e</sub>+D<sub>i</sub>)). Adapted from Waldeck et al. 1997 [69,72,80,81].</figcaption></figure>

**Section 3.2.** They were also ignored in the later PFG NMR studies on cells published since the 1990's, which aimed at measuring trans-membrane water exchange in erythrocytes with varying cholesterol contents [85] and in the Gram positive *Corynebacterium glutamicum* [86], or self-diffusion coefficients of water in cultured mammalian cells through the cell cycle [87] and in camel erythrocytes [88]. PFG NMR experiments have also been used to measure simultaneously both water self-diffusion coefficients and trans-membrane exchange in various systems: in yeast at different growth phases [89], in unicellular algae *Chlorella* [90], in erythrocytes [91-94], in cultured mammalian cells [95-97], or in plant systems like banana tissues [98], lupin roots upon exposure to lead [99], in maize roots of different lengths, or upon oxidative stress or at varying CO₂ concentrations [100-102]. Interestingly, MRI contrast agents were shown to inhibit transmembrane water exchange in this latter system [103].

Recent improvements have been proposed to this technique [75-79]. An interesting example takes advantage of the full signal decay of species in the extracellular medium, which experience unrestricted diffusion: a pair of strong pulsed gradients generate a “diffusion-filter” that leaves only the species in the restricted intracellular volume observable [75,76]. Their magnetization can be subsequently manipulated for any purpose, including diffusion measurements. Such a diffusion-filter has also been used for in-cell structural biology: it helped to filter out the leaking population of proteins and thus to observe intracellular proteins selectively [104].

In a rough summary, NMR spectroscopy has helped to evaluate the mean intracellular lifetime of water molecules to be about 10-20 ms in erythrocytes, and about 100 ms in HeLa cells. It also permitted quantification of intracellular water self-diffusion: it is about half as fast in cells as it is in pure aqueous buffers, and this number varies only slightly depending on cell types and conditions.

Concerning water rotational dynamics, NMR spectroscopy and more specifically water T1 relaxation- has been used to study them, notably in the context of macromolecules hydration since the 1960's [105-109]. Many mechanisms combine to influence ¹H T1 relaxation in biological samples: solute-water cross-relaxation, solute-water proton exchange and the broad diversity of water interactions, which provoke many magnetic interactions and relaxation pathways [106,107,110,111]. Hence, interpretation of ¹H T1 relaxation often remains ambiguous. Fortunately, water dynamics can be studied using NMR spectroscopy of ¹⁷O- and ²H nuclei, whose low gyromagnetic ratios and quadrupolar nature of these nuclei are better adapted to water dynamics studies. The ¹⁷O and ²H bulk water relaxation times are largely immune to the phenomena of cross-relaxation and spin-diffusion, and are also only little affected by chemical exchange. These relaxation times are dominated by quadrupolar interactions, which are mostly sensitive to tumbling dynamics. ¹⁷O and ²H bulk water relaxation rates are fast enough to allow monitoring of water dynamics from the picosecond to the nano-microsecond timescale. Measuring magnetic relaxation at varying fields provides “relaxation dispersion”
profiles reflecting the exchange rates between "bound" and "free" water molecules.

The most abundant isotope of oxygen, ¹⁶O, has a null spin and is thus useless from an NMR standpoint. The only "NMR-visible" isotope of oxygen, ¹⁷O, is rare (< 0.04 % natural abundance), but can be enriched and incorporated in biological systems, where it becomes a potent NMR tracer. Although not trivial to manipulate, its spin 5/2 can be fruitfully exploited to delineate water populations that interact with cellular species. Succinctly, the quadrupolar nature of ¹⁷O causes faster relaxation of water molecules "bound" to macromolecules. The very large field of ¹⁷O-NMR studies is beyond our scope, dedicated reviews are available [112,113]. Early work in the 1970's used ¹⁷O relaxation measurements to show that multiple water populations were constantly exchanging in cells, notably a minor fraction bound to nucleic acids or proteins, and a major "free" fraction [55,114-117]. In publications from the 1990's, this helped to estimate that below 1% of the cellular water molecules could be considered as "strongly bound", i.e. hydrogen-bonded, with correlation times of about 5-10 ns [118]. This number could reach 2% in mitochondria, but ¹⁷O NMR confirmed that the great majority of cellular water molecules had correlation times in the picosecond timescale [119]. This dismissed the existence of any substantial population of "ordered" water molecules in these highly crowded organelles, although they contain about 400 g/L of macromolecules. In bacteriophage Pf1, 4% of the water molecules are bound to macromolecules according to ¹⁷O relaxation measurements, which corresponds to about 4 water molecules per phage nucleotide [120].

In the late 2000's, Halle and colleagues measured ²H and ¹⁷O magnetic relaxation dispersion in *E. coli*, in the halophile archaea *Haloarcula marismortui* and in *Bacillus subtilis* spores [121-123]. In both *E. coli* and *H. marismortui*, they showed that ~85% of water molecules had correlation times of about 1.5 ps, 15% had correlation times of about 25-30 ps corresponding to the hydration-shells of cellular macromolecules, and only 0.1 % of water molecules were trapped in buried sites on the microsecond time scale. Even in the dense core of *B. subtilis* spores, the hydration water molecules had tumbling times of only 50 ps and buried water represented ~0.15 % of the total. These numbers were in line with those measured on *in vitro* dilute proteins samples.

We stress again the fact that these experiments do not report the translational behavior of water molecules, which can be measured by neutron scattering [124-126] and NMR measurements of diffusion (see above). The numbers above are consistent with results from more recent studies using other techniques like time-resolved infrared spectroscopy or dielectric relaxation spectroscopy [127].

Altogether, NMR spectroscopy has thus contributed greatly to the progressive understanding of water dynamics in cells.

### 2.1.2. Quantification and interactions of metal ions:

Metal ions are key components of cellular life, involved in modulating solvent properties, membrane potentials and thus neuron or muscle excitability, and in enzyme catalytic sites. They find applications in imaging or radiation therapy. They all have one or multiple isotopes with non-zero magnetic moment, but these are not necessarily found at high natural abundance (see [128] or https://www.webelements.com).

⁶Li⁺ and ⁷Li⁺, ²³Na⁺, ²⁷Al³⁺, ³⁹K⁺, ⁸⁵Rb⁺ and ⁸⁷Rb⁺, ¹¹³Cd²⁺ and ¹³³Cs⁺ have all been studied by direct NMR detection in cells. They have also been studied using molecules that change their chemical shifts according to whether they are in extra- or intra-cellular media, the so-called "shift reagents" (Fig. 2). These contain paramagnetic ions that shift the observed NMR frequencies of species in the same sample compartment. Hence, they make it possible to distinguish intra- and extra-cellular species. These shift reagents

are similar to contrast agents used for MRI, but induce less NMR relaxation of the solvent signals. Some "smart" contrast agents have been designed more recently by coupling basic contrast agents to metal-ion-chelating scaffolds: upon ion binding of the second moiety, the paramagnetic core of the contrast agent is exposed to the solvent (Fig. 3). These ion-activated contrast agents can thus reveal the presence of free cellular metal ions by enhancing the water NMR relaxation.

Mg²⁺, Ca²⁺, Fe²⁺, Zn²⁺ have active NMR isotopes at very low abundance, which precludes sensitive direct NMR detection. Moreover, paramagnetic states prevent NMR direct detection of Mn(II), Mn(III), Fe(III), Cu(II), Co(II) to name only a few. The abundant and diamagnetic isotopes of metal ions are however not easy to handle either. Their high spin ≥ 3/2 causes fast, multi-exponential relaxation, and thus intrinsically weak NMR signals. Their low gyromagnetic ratios can further plague their NMR detectability, which is typically the case for ³⁹K. Hence, reporters have been designed that incorporate sensitive nuclei (e.g. ¹⁹F), whose chemical shifts change when chelating the cation of interest. These will be called "labeled chelators/sensors" (Fig. 4).

We describe the strategies used in NMR studies of the main ions in the sections below. Some further pieces of information on these nuclei are given in Section 3.3.1. Other more exotic species have been studied sporadically by NMR in cells, e.g. ¹⁰B/¹¹B or ¹⁹⁵Pt [146-152]. Overall, NMR has been very useful in understanding the dynamics of all these ions in cells. NMR of metal ions is unable to provide spatial information at the (sub)micrometric scale of the subcellular level. However, it is still a very versatile technique that we suggest may find further applications in nutrition or toxicology studies on living microorganisms, plants, small animals or organoids, where it would also offer millimetric imaging capacities. All the studies below may be reinterpreted/readapted with these capabilities in mind.

### 2.1.3. Li⁺

Lithium is only found in trace amounts in living organisms, but it is commonly used therapeutically in treatment of bipolar disorders and may be effective against some cancers or neurodegenerative diseases [153-155]. The mechanisms behind its beneficial and adverse effects are still not well defined: they may be related to competition with Mg²⁺ in binding to kinases or phosphatases, competition with Na⁺ in GPCR binding, or membrane binding [156-158]. Both ⁶Li and ⁷Li have non-zero spins, but ⁷Li has been almost the only species used in NMR cellular studies, because it is much more abundant and shows better relaxation properties than ⁶Li [159]. Important results came from solution NMR investigations using direct ⁷Li one-dimensional spectroscopy. In a first study in the 1970's, Andrasko used the PFG-NMR strategy (see above in Section 2.1.1.) to reveal the restriction of Li⁺ diffusion in erythrocytes and measure their transmembrane exchange [160]. Then, in the 1980-90's, the use of "shift reagents" and solution ⁷Li-NMR permitted non-invasive quantification of Li⁺ cellular uptake, and detection of Li⁺ interactions with nucleic acids or phospholipids. Monitoring of Li⁺ cellular uptake was performed in yeast [161] and in erythrocytes [162-164] and astrocytomas [165]. An improved, more membrane-impermeant Tm-DOTP "shift reagent" was used in the 1990's to study Li⁺ cellular uptake of erythrocytes [166] and neuroblastoma cells (Fig. 5) [167,168]. ⁷Li has also been used as a surrogate for studying Na⁺ efflux in rat hearts [169-171].

Measurements of ⁷Li NMR relaxation times helped to show that it was binding preferentially to the inner phospholipid layer of erythrocytes (and not to sialic acid, actin or hemoglobin) [172-175]. Li⁺ phospholipid binding was later observed also in neuroblastoma cells [65]. Li⁺ competes with Mg²⁺ for binding to phospholipid in erythrocytes, as shown by fluorescence microscopy and ³¹P NMR [176] and later by ⁶Li and ⁷Li NMR relaxation measurements
<figure><img src="image_6.png" /><figcaption>**Fig. 2.** Operating principle of "shift reagents", A) Chemical structures of the main shift reagents used in cellular studies (adapted from Szklaruk et al. 1990 [129]): these chelators are typically loaded with transition metals (most often Mn²⁺) or lanthanide ions. B) Schematic ¹H NMR signals from mouse tissue water and fat, adapted from Harnden et al. 2019 [130]: the extracellular chelated paramagnetic center provokes important changes in chemical shifts of extracellular species, here water, or C) ²³Na⁺ NMR signals from human erythrocytes in suspension (the reference signal is generated from a capillary containing a pure aqueous NaCl solution), adapted from Sherry et al. 1988 [131]. Paramagnetic induced chemical shifts are mostly due to the merging effects of hyperfine shifts and bulk magnetic susceptibility (see [130,132,133] for more comprehensive explanations). Shift reagents also accelerate relaxation times and thus signal linewidths of species in the same sample compartment.</figcaption></figure>

<figure><img src="image_7.png" /><figcaption>**Fig. 3.** Operating principle of ion-dependent ("smart") contrast agents, adapted from Barandov et al. 2019 [134]. A) Schematic chemical structure of a Ca²⁺-activated, Mn²⁺-contrast agent: a Mn²⁺ chelator is linked to an acetomethoxyl-BAPTA moiety; the acetomethoxyl ester groups are removed upon cell internalization by cellular enzymes, which traps the complex in cells. B) In presence of free Ca²⁺, the BAPTA moiety chelates Ca²⁺ preferentially and does not chelate any longer the Mn²⁺ paramagnetic center, which, in turn, is more exposed to solvent; this induces a faster NMR relaxation of solvent signals. The insets show the resulting ¹H-water signal observed by MRI. A number of chemical scaffolds have been proposed with the same purpose [135-141]. These can be used for NMR studies on cells; the final goal is often to obtain probes for functional MRI.</figcaption></figure>

<figure><img src="image_8.png" /><figcaption>**Fig. 4.** Operating principle of "labeled chelators/sensors" that report the presence of metal ions in cells, adapted from Bar-shir et al. 2013 and 2015 [142,143]. We show here one of the most commonly used scaffolds in the literature, the cell-penetrant BAPTA chelators. Their acetoxymethyl ester form (nF-BAPTA-AM) freely enters into cells, where it is converted to an impermeable form (devoid of acetoxymethyl ester function, nF-BAPTA) by intracellular esterases [144,145]. Being fluorinated, these molecules are observable by ¹⁹F-NMR spectroscopy, which does not suffer any background signal in cells. Different fluorination schemes result in different affinities and exchange rates for every divalent metal ion. A) Chemical structure of 5,5',6,6'-TF-BAPTA and B) the 1D ¹⁹F NMR obtained in presence of Fe²⁺ or Zn²⁺, showing the 5,5'–¹⁹F and 6,6'–¹⁹F signals in purple and green, respectively. C) Chemical structure of 5F-BAPTA and the exchange between free and chelating situations, whose exchange rates D) cause different ¹⁹F signal linewidths in 1D ¹⁹F NMR spectra (recorded at 11.7 T, k<sub>ex</sub> is the exchange rate, and Δω is the difference of NMR frequencies between the signals from the free and chelating forms of nF-BAPTA). Other labeled sensors have been proposed, which can also use ¹³C- or ¹⁵N- labeling. The sensitivity of these sensors can be enhanced by Chemical Exchange Saturation Transfer (CEST) methods, allowing detection levels of metal ion concentration in the submicromolar range [142,143].</figcaption></figure>
<figure><img src="image_9.png" /><figcaption>**Fig. 5.** A) 1D $^{7}$Li NMR spectra showing the Li⁺ influx in SH-SY5Y human neuroblastoma cells perfused by a culture medium supplemented with the shift agent HTmDOTP⁴⁻ and with LiCl at 15 mM after (a) 11 min, (b) 39 min and (c) 133 min; B) 1D $^{7}$Li NMR spectra showing the Li⁺ efflux from Li⁺-loaded SH-SY5Y human neuroblastoma cells perfused by a culture medium supplemented with the shift agent HTmDOTP⁴⁻ and with LiCl at 5 mM after (d) 75 min, (e) 90 min and (f) 120 min. Adapted from Nikolakopoulos et al. [168].</figcaption></figure>

[177]. $^{7}$Li relaxation was also used to demonstrate another competition of Li⁺ with Mg²⁺ in chromaffin cells and in lymphoblastoma and neuroblastoma, where faster relaxation was measured than in erythrocytes [178,179]. This was interpreted as being due to supplementary interactions of Li⁺ with unknown targets, or to changes in composition of cellular phospholipids upon exposure to Li⁺ [180,181]. Li⁺ does indeed compete with Mg²⁺ at many levels [156]: it binds to ATP [182], to the kinase Gsk3β, and to the inositol phosphatase [183], provoking an increase of phosphoinositol content. The precise mechanisms of Li⁺ actions are thus not yet elucidated, but in-cell NMR has made important contributions in the field. These studies have moreover paved the way for $^{7}$Li-MRI approaches, which have helped improve understanding of the distribution of Li⁺ in brain upon lithium treatments [159,184,185].

### 2.1.4. Na⁺

Na⁺ is less abundant in cells (~10-30 mM) than in extracellular fluids (~100-150 mM). The mechanisms ensuring low Na⁺ intracellular concentrations are of great importance, notably because they are essential to the proper functioning of excitable cells and of Na⁺-dependent membrane proteins [186]. Because $^{23}$Na represents 100% of sodium and has a relatively favorable gyromagnetic ratio (about ¼ of $^{1}$H), $^{23}$Na NMR has long been attractive for NMR studies.

Early $^{23}$Na⁺ NMR measurements by Cope in the 1960's reported large NMR-invisible populations, representing about 60% of intracellular $^{23}$Na⁺ [187,188], similar to the situation for $^{39}$K⁺ that we will describe later (see below and Section 3.1.2.). The "invisibility" of this fraction of the intracellular $^{23}$Na⁺ population was interpreted by some authors as indicating that it was bound or immobilized [187-193]. An exception was found in the very "watery" plant vacuoles, where few interactions with macromolecules exist [194]. However, Civan and Shporer showed in the 1970's that these early measurements had to be re-interpreted using an appropriate theory of quadrupolar nuclei relaxation, which led to the

conclusion that actually less than 1% of $^{23}$Na⁺ was bound and immobilized in cells [195-197]. Parallel investigations on $^{39}$K reinforced this theoretical framework (see below). T2 relaxation of cellular $^{23}$Na⁺ has two components, one fast (~0.5-2.5 ms) and one slow (~10-50 ms) corresponding to 60% and 40% of the total signals, respectively. In the early times, the experimental set up was not likely to permit detection of the fastest relaxation components of quadrupolar ions in cells – hence the invisibility of 60% of the signal.

Improved equipment such as faster electronic systems and higher field spectrometers (>5 T) paved the way to studies focusing on Na⁺ transmembrane exchange in the 1980's. In 1982, Springer, Gupta and their coworkers proposed independently the use of extracellular shift reagents, i.e. dysprosium and thulium complexes, to distinguish NMR signals from intra- and extra-cellular $^{23}$Na⁺ (Fig. 6) [161,198,199]. Recording a time series of simple 1D $^{23}$Na NMR spectra was thus sufficient to monitor the influx and efflux of Na⁺ in a non-invasive fashion, in real-time with a time-frame of one minute. At that time, other methods were used for such measurements, such as atomic absorption spectrometry, flame-emission photometry, radioisotope tracing and ion-selective microelectrodes. These were destructive, time consuming methods, which did not come to consistent conclusions on cation transport and dynamic equilibria [200-204]. $^{23}$Na NMR was thus seen as an interesting opportunity in the field, not requiring prior separation of the cells or their culture medium. This strategy using shift reagents was adopted by many groups in the 1980's to monitor intracellular levels of $^{23}$Na⁺ in yeast [161,200], in bacteria [205-214], in frog oocytes [215], in amoeba [216], in erythrocytes [203,217-222], in animal tissues and organs such as perfused hearts [223-230] and in living rats or rabbits [231-235]. These NMR experiments were combined with $^{31}$P-NMR to measure intracellular pH in parallel, and sometimes also with $^{39}$K NMR. Among others things, this permitted studies of the regulation of Na⁺ levels in microorganisms submitted to osmotic shocks
<figure><img src="image_10.png" /><figcaption>**Fig. 6.** 1D $^{23}$Na NMR spectrum of human erythrocytes (27% hematocrit) resuspended in a medium containing the shift agents Tm(PPP)$_2$ necessary to separate intra- (at 0 ppm) and extra- (at ~2 ppm) cellular $^{23}$Na$^+$ NMR signals. The dashed lines in the inset show the two fitted Lorentzian lines of the intracellular $^{23}$Na$^+$ signal, with a fast and a slow relaxing component representing 40% and 60% of the integrated intensity, respectively. The spectrum was recorded at 8.46 T (frequency of $^{23}$Na: 95.245 MHz) in ~9 minutes in 1984. Adapted from Pike et al. 1984 [242].</figcaption></figure>

[200,205,206,211,213], evaluation of the selectivity of cation-carrying chemicals penetrating Gram positive bacteria [209,221,222,236], and measurement of the activity of the Na⁺/K⁺ ATPase pump in situ [224,228,229,237]. The assumed stability and inertness of the shift reagents used was however disproved in the late 1980's [238-241].

Another approach has been adopted to observe the signal from intracellular $^{23}$Na⁺ selectively: selecting the double-quantum transitions of $^{23}$Na has a filtering effect, which permits observation of only those $^{23}$Na⁺ ions that experience the bi-exponential relaxation caused by interactions with macromolecules [243-245]. Even though much attenuated, the extracellular $^{23}$Na⁺ signal can also immediately be recognized in such Double-Quantum filtered (DQF) spectra, because of interactions with exterior cellular membranes or proteins in the extracellular medium [246,247]. Addition of a low concentration of extracellular paramagnetic Gd³⁺ was necessary to quench this remaining signal. Triple-Quantum filtering (TQF) was shown to perform slightly better in isolating the intracellular $^{23}$Na⁺ signal [248,249], but was also not capable of canceling the extracellular $^{23}$Na⁺ signal completely (Fig. 7) [250-252]. In the mid 1990's, these DQF or TQF methods were still proposed to monitor intracellular Na⁺ levels upon ischemia in cultured cells [253,254] or in perfused rat heart [255], because the DQF/TQF $^{23}$Na⁺ signal variations were only due to changes in intracellular Na⁺ levels. TQF $^{23}$Na NMR has been applied to study the intracellular sodium levels in tumors implanted in mice or rats [256,256-259], and even to map intracellular sodium in human brain [260,261]. TQF suffers from significant signal-to-noise losses as compared to Single-Quantum $^{23}$Na NMR, and other $^{23}$Na MRI strategies have been developed, which we can only mention here (see [262-265]). Recent improvements in pulse sequences permitted detection of TQ and SQ signals simultaneously with improved sensitivity, and reporting of the minute-timescale evolution of intracellular $^{23}$Na⁺ levels in just ~15 million cells [266-268].

### 2.1.5. K⁺, and Rb⁺ as a surrogate

Potassium is the least receptive of the alkali metals in NMR studies. Indeed, although found in high natural abundance (93.3%), $^{39}$K suffers from its low gyromagnetic ratio and its quadrupolar nature, yielding 180-fold lower in vivo sensitivity than

$^{23}$Na. $^{39}$K was first observed in cells in 1970 using halobacteria that contain intracellular K⁺ concentrations as high as 4-5 mol/L [269]. Importantly, $^{39}$K NMR has also helped in understanding cellular ion mobility. Similar to water proton and $^{23}$Na NMR studies (see above), erroneous conclusions were drawn in the late 1960's from data obtained with early spectrometers and incorrect interpretations, namely that the weak 30-40% NMR visibility of cellular $^{39}$K was due to the remaining K⁺ being immobilized in a semitrystilline cellular milieu [269]. Very soon afterwards, in the 1970's, studies using more advanced equipment and NMR theory showed that only at most a few percent of $^{39}$K⁺ were actually transiently interacting with cellular molecules [270-273]. We give more details on this controversy in Section 3.1.2. Starting from the early 1980's, shift reagents permitted distinction of extra- and intra-cellular $^{39}$K, and thus allowed them to be quantified [274], to monitor transmembrane ion transport in erythrocytes [218], in halobacteria [275], in yeast [200], in E. coli [276], and later in frog [277], rat [225,278] or guinea pig hearts [279], and rat salivary glands [280]. Because potassium homeostasis is linked to heart injuries or cellular membrane potential in brain, $^{39}$K MRI studies were attempted from the late 1990's [281]. Use of higher magnetic fields [282,283] and more sensitive spectrometer probes [284] progressively pushed the sensitivity limits of this approach.

Rubidium has for long been established as a good surrogate for cellular potassium uptake [285], and used in quantitative studies primarily in its radioactive form $^{86}$Rb [286]. Its stable isotope $^{87}$Rb was considered in the late 1980's for achieving NMR studies of potassium uptake in intact rat kidney and heart [287-289], in mammalian cultured cells [290], in erythrocytes [291,292], in rat salivary glands [293] and in skeletal muscles of living rats [294]. Sensitivity assessment in erythrocytes indeed showed that direct $^{87}$Rb NMR detection was likely to provide about 19 times more signal than $^{39}$K [291]. However, as with $^{23}$Na or $^{39}$K, the quadrupolar nature of $^{87}$Rb causes fast, bi-exponential relaxation in cells [295,296]. This was initially used to distinguish two overlapped $^{87}$Rb peaks, one broad signal that would correspond to intrakidney $^{87}$Rb⁺, and one narrow revealing the extracellular signal [287,288]. Fitting to the sum of these Lorentzian signals helped to quantify $^{87}$Rb uptake in perfused organs, where the $^{87}$Rb signal was so broad that shift agents in the extracellular milieu could
<figure><img src="image_11.png" /><figcaption>**Fig. 7.** A) 1D $^{23}$Na NMR spectra of a perfused rat heart: One pulse SQ NMR (upper spectrum, 24 scans, t$_{acq}$~5 s) and TQF (lower spectrum, 960 scans, t$_{acq}$~200 s) spectra; B) 1D $^{23}$Na NMR spectra of a perfused rat heart in a medium containing the shift reagent [Dy$^{3+}$]-(tripolyphosphate)$_2$: One pulse SQ NMR (upper spectrum) and TQF (lower spectrum) spectra, showing intra- (peak at the left) and extra- (peak at the right) cellular $^{23}$Na$^+$ signals; C) Plotted intensities of signals from intra- (square) and extra-cellular (circles) $^{23}$Na$^+$ in 1D $^{23}$Na TQF NMR spectra of perfused (triplicates) rat hearts in a medium containing Tm(DOTP)$^{5-}$ as a shift reagent, before and after exposure to the inhibitor of the Na$^+/K^+$-ATPase pump ouabain, leading to an increase of intracellular Na$^+$. A) and B) are adapted from Jelicks et al. 1993 [250], the spectra were recorded at 9.4 T in 1993; C) is adapted from Dizon et al. 1996 [255].</figcaption></figure>

not separate extra- and intra-cellular peaks [297]. A strategy was proposed to cancel the extracellular $^{87}$Rb signal using lanthanide contrast reagents [298]. In later studies on animal tissues, an inter- nal reference was used to quantify $^{87}$Rb$^+$ concentration, which con- sists of a capillary containing $^{87}$Rb in a KI saturated solution, generating a $^{87}$Rb signal shifted by 55 ppm [294]. $^{87}$Rb NMR spec- troscopy evolved towards functional studies on Na$^+/K^+$ ATPase activity in injured hearts [299] and brain potassium imaging [300,301]. We have not found any study using $^{87}$Rb NMR published in the last ten years. The limited sensitivity and natural abundance of $^{87}$Rb requires high concentrations of Rb$^+$ for measurement, resulting in partial replacement of K$^+$ in cells, with reported cardiac consequences [299].

### 2.1.6. Mg$^{2+}$

With an intracellular concentration in the range of 10 mM, Mg$^{2+}$ is the second most abundant cation in cells after K$^+$ [302]. It is essential in many regards, notably for the proper functioning of many enzymes and channel proteins [303]. It can be considered an integral part of many RNA and DNA structures [304-306]. Understanding the impact of intra- and extra-cellular Mg$^{2+}$ levels in many pathologies has been a longstanding aim. The only NMR visible isotope is $^{25}$Mg, which has a very low intrinsic sensitivity because of its poor natural abundance (~10%), its low gyromag- netic ratio and its spin 5/2. It has thus been studied mostly via its chelating partners and their chemical shift changes upon Mg$^{2+}$ binding.

The main example of this is ATP, whose $^{31}$P NMR chemical shifts follow the Mg$^{2+}$ binding equilibrium [202,307,308]. ATP proton chemical shifts of the ribose also depend on Mg$^{2+}$ chelation [309,310]. As we will see in Section 2.3.2. on metabolites, $^{31}$P NMR is convenient in cells because of its relatively good sensitivity when present in small molecules and the small number of observ- able signals. Hence, $^{31}$P NMR and the survey of ATP chemical shifts has been used since the late 1970's [311,312] to measure the intra- cellular free Mg$^{2+}$ and the ATP:Mg$^{2+}$ population in erythrocytes [202,312-317], lymphocytes [318], Ehrlich ascites tumor cells [319], cultured mammalian cells [320,321], animal tissues [322- 327], in worms, amoebae or sea snail [328-330], and more recently in MRI/MRS studies [331-335]. These $^{31}$P NMR studies have regu- larly reported that the concentration of free intracellular Mg$^{2+}$ is only ~0.3-0.6 mM in mammalian cells, while the total concentra- tion of 10-20 mM in most cells [336] (with an exception: only

~2 mM in erythrocytes that contain low quantities of nucleic acids). They also revealed that more than 90% of the cellular ATP (intracellular concentrations between 2 and 10 mM) is found in a Mg$^{2+}$ bound form. These measurements could be ambiguous, because of the uncertain amount of invisible ATP that would bind cellular macromolecules, because of the possible compartmental- ization of ATP:Mg$^{2+}$, because of the salt and pH dependency of ATP:Mg$^{2+}$ association constant, and because the ATP $^{31}$P chemical shifts are themselves strongly dependent on pH and temperature, all of which aspects were progressively taken into account as mod- els developed [202,307,308,326,337-343]. Diphtosphoglycerate $^{31}$P chemical shifts are also dependent on pH and Mg$^{2+}$ chelation, and it was thus used with ATP in a complementary fashion to mea- sure free Mg$^{2+}$ and pH in erythrocytes (Fig. 8) [344]. The $^{31}$P peak linewidths of ATP have also been proposed for quantification of Mg$^{2+}$ binding [345,346], although this type of analysis may be affected by field homogeneity [347]. In any case, these measure- ments were non-invasive and extremely useful, by making it pos- sible to study the evolution of ATP:Mg$^{2+}$ and cellular free Mg$^{2+}$ levels under different stresses. They provided a complementary method to the early fluorescence measurements, which tended to overestimate the free Mg$^{2+}$ levels. The studies based on fluores- cence spectroscopy used reporters that had their flaws in terms of selectivity (notably towards Ca$^{2+}$) and leakage, in addition to the potential that they might compete with endogenous binders of Mg$^{2+}$ [202,307,348]. Improved fluorescence sensors have since been tailored, although the perfect sensor is still elusive [349,350].

To complete this section, we present a few direct detection NMR studies that use $^{25}$Mg: by measuring the $^{25}$Mg NMR peak linewidth at increasing concentrations of ATP, ADP, DPG, or in presence of actin filaments, these studies have provided direct proof of the formation of ATP:Mg$^{2+}$ and (ATP)$_2$:Mg$^{2+}$ (Fig. 9) [351] and of the very loose binding of Mg$^{2+}$ to actin filaments acting only like polyelectrolytes and not as a cognate partner [352]. The chelated Mg$^{2+}$ is in fast exchange with free Mg$^{2+}$ and is never trapped in these complexes unless actin filament bundles form. $^{25}$Mg$^{2+}$ has also been shown to have weak affinities for a number of proteins [351,353-355]. An interesting feature is its regular competition with Ca$^{2+}$ and not with other cations [353,354]. All these $^{25}$Mg NMR *in vitro* studies most probably recapitulate some cellular aspects. We did not find any published in-cell $^{25}$Mg studies. Still, these studies show how much information NMR has yielded that shaped understanding of cellular Mg$^{2+}$.
<figure><img src="image_12.png" /><figcaption>**Fig. 8.** A) 1D ³¹P NMR spectrum of human erythrocytes (~35-50% hematocrit, recorded at 310K and 9.4 T in 2002, 2048 scans, tₐcq~30 min), showing the ³¹P signals from the two main species diphosphoglycerate (or 2,3-bisphosphoglycerate) and ATP; B) Chemical structures of DPG and ATP; C) *Up*: Plot of the chemical shift (absolute values) of the β phosphate of ATP in function of pH and of [Mg]ᵢₐcq; *Bottom*: Plot of the chemical shift of the 3-phosphate of DPG in function of pH and of [Mg]ᵢₐcq. Adapted from Willcocks et al. 2002 [344].</figcaption></figure>

### 2.1.7. Ca²⁺

Ca²⁺ is involved in an very wide variety of cell signaling pathways, from cell proliferation to apoptosis, thymocytes activation or synaptic transmission and cell contraction in metazoans [356], and in many adaptive reactions in plants [357]. These all rely on spatiotemporal variations of cellular concentrations of Ca²⁺. Resting cells keep their cytosolic Ca²⁺ at about 100 nM and store Ca²⁺ in the Golgi apparatus, endoplasmic reticulum (ER) and sarcoplasmic reticulum of muscle cells at 0.5-1 mM. These organelles eventually trigger calcium signaling upon stimulation via Ca²⁺ release that enhances its cytosolic concentration to 200-500 nM (extracellular Ca²⁺ is at 2 mM) [356,358]. Direct NMR detection of calcium in solution is impractical: the only detectable isotope ⁴³Ca is present at very low natural abundance (0.14%) and yields low and broad signals because of its quadrupolar nature [359]. Very few solution ⁴³Ca NMR studies have been published, to the best of our knowledge, and ⁴³Ca NMR is probably more of interest for solid-state applications [360].

This limitation has motivated NMR spectroscopists to test calcium detection via calcium-sensitive probes in the early 1980's: inspired by existing fluorescent probes, Ca²⁺ chelators were designed to integrate ¹⁹F atoms in their chemical structures, which

permitted ¹⁹F-NMR detection and quantification of cytosolic Ca²⁺ in thymocytes [144]. The free and chelating forms of these ¹⁹F-labeled sensors indeed do show two different chemical shifts, and the area of the two peaks are proportional to the corresponding populations (Fig. 4). This principle has been very widely exploited. It has been applied to Ca²⁺ measurement in aging or sickle erythrocytes [145,361-363], platelets [364], synaptosomes [365], leukemic cells [366], osteoblasts [367,368], hear cells [369], kidneys [370], perfused hearts [371-373] and brain [374-376] in the native ~50-100 nM range. The initial sensors have moreover been modified for tuning cell delivery and for quantifying other metals: ¹⁹F-containing chelators have different affinities and ligation kinetics for every metal, and are also associated with different specific chemical shifts [137]. The initial advance was the use of acetoxymethyl ester 5F-BAPTA-AM, which freely enters cells and is converted to the cell-impermeant 5F-BAPTA by intracellular esterases [144,145]. The 5F-BAPTA sensor was actually able to report simultaneously cellular Ca²⁺ and Zn²⁺ concentrations [365,377,378] or Ca²⁺ and Pb²⁺ [367], and later Ca²⁺, Cd²⁺ and Zn²⁺ [379]. It has for example been used recently to monitor Ca²⁺ uptake in erythrocytes upon mechanical distortion [380]. Other sensors for transition metals quantification or pH measurement were developed (see below). Moreover, because 5F-BAPTA ¹⁹F resonances were also recognized to be pH-sensitive, pH sensors were designed in a similar way (see below).

In a series of derivatives, the ~500 nM Ca²⁺ affinity of 5F-BAPTA was modified down to ~50 nM for DiMe-5F-BAPTA, which permitted lower sensor load, hence less perturbation of Ca²⁺ pulses in perfused beating hearts [381,382]. These improvements were of course motivated by the opportunity to carry out *in vivo* Ca²⁺ studies, which started in the 1990's [383]. It was however hampered by the low sensitivity of 5F-BAPTA ¹⁹F-NMR spectroscopy, estimated to be 10⁵ fold less than standard proton MRI [384].

Chemical Exchange Saturation Transfer (CEST) approaches (Fig. 4, Fig. 14) have been proposed that enhance 5F-BAPTA-mediated Ca²⁺ detection by a factor 100 [142,143]. To further increase sensitivity, sensors incorporating multiple equivalent ¹⁹F nuclei were designed and combined with the use of CEST; similarly, ¹⁵N- or ¹³C-containing sensors were hyperpolarized and used in CEST strategies as well; Ca²⁺ sensors coupled to cryptophane cages recognizing hyperpolarized ¹²⁹Xe were also proposed in combination with CEST. None of these methods have been tested yet in cells to the best of our knowledge [137,385].

<figure><img src="image_13.png" /><figcaption>**Fig. 9.** 1D ²⁵Mg NMR spectra of samples containing A) 2 mM ²⁵MgCl₂, 100 mM KCl, 20 mM Tris-HCl, 20% ²H₂O at pH7.2, B) supplemented with 2 mM ATP and C) 16 mM ATP. Spectra were obtained from A) 256 scans (tₐcq~100 s), B) 4096 scans (tₐcq~1600 s) and C) 29109 scans (tₐcq~11600 s) at 9.4 T (²⁵Mg frequency of 24.485 MHz) in 1991. (Adapted from Bock et al. [351])</figcaption></figure>
Another approach was developed, which takes advantage of
molecules combining a Ca²⁺ chelator fragment that modulates
the relaxivity of a paramagnetic fragment (Fig. 3): the first of these
"smart contrast agents" was designed in the late 1990's, and
exposed the paramagnetic Gd³⁺ to the solvent in presence of Ca²⁺
[386]. This strategy adopts the common MRI strategy of contrast
agents, which accelerates water ¹H resonance relaxation, revealed
by lower signal intensities. It was progressively improved by a
number of groups, tested in cells [136,387], and recently used in
MRI studies of extracellular Ca²⁺ dynamics in the brain interstitial
space [388] and during cerebral ischemia [389]. Cell-permeable
Ca²⁺-sensor contrast agents were also designed, which have shown
MRI compatible sensitivity in cultured cells [138] and even in liv-
ing rats [134]. However, none of these developments circumvent
the inability of NMR to assess the functional inhomogeneity of
intracellular Ca²⁺, which are accessible by fluorescent probes for
example [358,390].

2.1.8. Transition metal ions Mn²⁺, Fe²⁺/Fe³⁺, Co²⁺, Ni²⁺, Cu⁺/Cu²⁺ and Zn²⁺, Pb²⁺

Because of their paramagnetic properties, transition metal ions Mn²⁺, Fe²⁺/Fe³⁺, Co²⁺, Ni²⁺, Cu²⁺ are not directly observed by NMR. They are mostly chelated and involved in enzymatic catalytic sites, and their concentration as free ions is very low in living organisms. Copper is present in cells in its diamagnetic ionic form Cu⁺, but it is also completely coordinated in proteins or by glutathione in prokaryotic and eukaryotic cells, resulting in sub-femtomolar concentrations of free Cu⁺ [391-395]. The quadrupolar ⁶³Cu and ⁶⁵Cu would be expected to provide very broad NMR signals and poor sensitivity. In-cell NMR spectroscopy is thus not very well adapted to probing the intracellular behavior and exchange of transition metals. However, they all bind the ¹⁹F-labeled Ca²⁺-sensor 5F-BAPTA and its derivatives with submicromolar affinities [382], with which they each show different chemical shifts, allowing them to be detected and identified simultaneously. Hence, using ¹⁹F NMR spectroscopy, authors have reported the parallel monitoring of Ca²⁺ and Pb²⁺ [367] or Ca²⁺, Cd²⁺ and Zn²⁺ in cultured cells [379]. However, we found only a limited body of literature on NMR studies in cells focusing on these ions. Other sensors have been suggested, and Ca²⁺ sensors or fluorescence reporters may be redesigned for NMR studies of these metals. Paramagnetic metal ions may help to accelerate T1 relaxation and S/N per experimental time in the future, as shown *in vitro* [396-401]. Gd³⁺ or V³⁺ complexes may be more suitable because they would avoid binding competition from endogenous divalent metal ions. Transition metal ions are also useful to establish MRI contrast agents in association with chelator molecules to induce faster water relaxation, or in combination with fluorinated sensors also for use in MRI [137]. Re-designing such agents to create cell-permeable forms, or adapting Ca²⁺-sensors, should be feasible in some cases [134].

Zn²⁺ is an integral structural component of about 3000 human
proteins, among which are included many enzymes or small pro-
tein domains, notably the well-known family of zinc-fingers
[402]. In total, cells contain Zn²⁺ at hundreds of micromolar con-
centrations, but only pico- to nano-molar concentrations of free
Zn²⁺. However, zinc may also have signaling functions and "Zn²⁺
waves" occur [403]. Zn²⁺ is not paramagnetic but its only NMR-
observable isotope ⁶⁷Zn is quadrupolar, has a low gyromagnetic
ratio and is present at low natural abundance (~4%). Given also
its low concentration, it is overall poorly amenable to NMR studies
via direct detection, and consistent with this, it has not been much
studied in cells using ⁶⁷Zn NMR, to the best of our knowledge. Like
other divalent ions, Zn²⁺ can still be detected through the NMR sig-
nals of chelating agents. Zn²⁺ has been recognized to bind the Ca²⁺
chelator 5F-BAPTA since the very first use of this reporter in the
early 1980's [144]. Zn²⁺ had actually a much better affinity than

Ca²⁺ for 5F-BAPTA, i.e. 1 versus 500 nM, respectively [382]. Fortu-
nately, Ca²⁺ and Zn²⁺ binding result in different ¹⁹F chemical shifts
of 5F-BAPTA, and the two ions have been detected in parallel in
Ehrlich ascite tumor cells [378], in perfused ferret hearts [377],
in guinea-pig cortical slices [376,404], and in cultured neuroblas-
toma cells [379]. Another BAPTA derivative, 5,5',6,6'-tetrafluoro-B
APTA (TF-BAPTA) has been to shown to chelate Ca²⁺ poorly, and
thus it is potentially a good sensor of intracellular Zn²⁺/Fe²⁺
[405]. Other ¹⁹F-labeled, ¹¹B-labeled or hyperpolarized ¹³C-
labeled or ¹²⁹Xe-cage ion sensors have been designed that can
report and distinguish the presence of Zn²⁺ and Ca²⁺ by NMR, but
with few applications in cells up to now [137]. We can also men-
tion that many groups have recently proposed Zn²⁺-activated
MRI contrast agents to probe extracellular Zn²⁺ in brain or prostate,
where it may have amyloidogenic properties or diagnostic poten-
tial, respectively [140,141,406,407]. Upon Zn²⁺ chelation, these
molecules either expose a paramagnetic center or exchangeable
protons to the solvent, or show altered chemical shifts of ¹⁹F
moieties.

Hence, NMR spectroscopy can report on the presence of free
divalent metal ions in cells, using cell-permeable sensors and their
chemical shift variations upon chelation. The subcellular localiza-
tion information is lost most of the time, but the chemical shift
variations are metal dependent. This ability to monitor and distin-
guish simultaneously multiple metal ions is not always perfectly
executed by fluorescence probes.

2.1.9. More highly toxic species: Al³⁺, Cd²⁺ and Cs⁺

NMR spectroscopy can help in studies focusing on toxic metal
species: the tissues or cells of interest can be characterized without
organelle separation and without solute extraction by hydropho-
bic/hydrophilic solvents. Moreover, they do not require radioactive
isotopes. Chemical shifts can reveal the type of chelation or the
storage organelle. Hence, NMR spectroscopy of species like Al³⁺,
Cd²⁺ and Cs⁺ can be highly complementary to other techniques.

Several ²⁷Al-NMR studies have been carried out on plants. Tea,
buckwheat, hydrangea or eucalyptus can accumulate high alu-
minum concentrations despite its usual toxicity. Detoxification
mechanisms have been characterized using solution ²⁷Al-NMR of
intact plant leaves. Direct one-dimensional ²⁷Al-NMR experiments
provided spectra of Al³⁺ stored in leaves of the organisms studied,
revealing chemical shifts identical to those of Al:citrate and Al:ox-
alate complexes [408-416]. This led to the conclusion that these
plants stored aluminum under these forms. In rape plant, Al:-
malate complex was also detected [417]. Hence, NMR was instru-
mental in describing these non-covalent storage strategies adopted
by plants.

Like ²⁷Al, ¹¹³Cd has permitted NMR studies of cadmium detox-
ification in plants, more specifically in the cadmium-accumulator
plant *Thlaspi caerulescens*. After growth in a ¹¹³Cd(NO₃)₂ enriched
nutrient medium, one-dimensional solution NMR of leaves using
direct ¹¹³Cd-detection revealed a ¹¹³Cd chemical shift identical to
that of the Cd:malate complex [418]. ¹¹³Cd NMR sensitivity is poor
though, so a ¹⁹F-labeled chelator probe has been used to report
simultaneously Cd²⁺ and Zn²⁺ uptakes and to measure their effects
on cellular Ca²⁺ concentration [379]. Cd²⁺ interaction with diphos-
phoglycerate (DPG) also gives rise to shifts of diphosphoglycerate
(DPG) ³¹P NMR signals, but these could not be detected in erythro-
cytes [419].

Living cells assimilate cesium primarily through potassium
transport systems [420]. The stable isotope ¹³³Cs has been
exploited as an NMR-practical ³⁹K congener to study potassium
cellular influx and efflux and intracellular distribution. Moreover,
cesium has a mild toxicity in regular circumstances, but is very
deleterious in the form of the radioactive isotope ¹³⁷Cs, which is
produced and released by military and civil nuclear activities. It
would thus be of interest to understand the toxicity mechanism of ¹³³Cs, and to characterize cesium integration in the food chain by plants [421]. ¹³³Cs has much better NMR characteristics than ³⁹K and ⁸⁷Rb: i) it represents 100% of stable cesium; ii) it has a small quadrupole moment yielding moderately fast relaxation, hence reasonably high and sharp NMR signals; iii) it shows a large chemical shift range and a high sensitivity to its chemical environment, especially to its counter anions [422]. Hence, two separated peaks from intracellular and extracellular media were early observed in halobacteria using one-dimensional direct ¹³³Cs NMR detection [423], which occurs through an active translocation mechanism resulting in cesium accumulation [424]. Several NMR studies on erythrocytes provided similar spectra showing two resonances from intra- and extracellular ¹³³Cs without the need of adding extracellular shift reagents. Cesium ions were shown to compete with potassium ions in the cellular uptake process, cesium rates being one third of that of potassium [425,426]. ¹³³Cs⁺ was thus used to study K⁺ transport in cultured cells of perfused rat hearts [427-430]. Magnetic susceptibility effects were shown to be only a minor cause of intracellular chemical shifts, while non-ideal water solvation and interactions with phospho-anions, notably diphosphoglycerate (DPG), had larger contributions [431-433]. This interaction with DPG appears to displace the DPG:deoxyhemoglobin interaction. Consistently, various ¹³³Cs chemical shifts were observed in various rat tissues organs [434,435], but the measured relaxation rates argued for a common Cs⁺-free state in fast exchange with anion-binding Cs⁺ [436]. Even more interesting, ¹³³Cs proved to be a good subcellular probe: two intracellular ¹³³Cs NMR peaks were detected in maize root tissue, corresponding to cytosolic and vacuolar compartmentation [437], and two peaks also appeared from cytoplasmic and mitochondrial ¹³³Cs internalized in hepatocytes [438]. Because the ¹³³Cs resonance is also very sensitive to nitrate concentration, the two subcellular ¹³³Cs reporter peaks permitted monitoring of in vivo nitrate incorporation into maize root compartments [439], and cesium/potassium exchange in *Arabidopsis* [440]. To the best of our knowledge, this interesting ¹³³Cs sensing of the chemical environment has been only sparsely used in the last 15 years. Because of its preference for the intracellular space, it has been less popular than ⁸⁷Rb in K⁺ translocation studies [300], although Rb⁺ has also been shown to be a better generator of K⁺ for influx than for efflux studies [291]. Moreover, the rather low sensitivity of ¹³³Cs NMR (γ(¹³³Cs)~γ(¹H)/8) requires too high intracellular concentrations given its intrinsic toxicity. Using DNP-sensitivity enhancement, recent studies achieving ¹³³Cs hyperpolarization followed by dissolution and mixing with live cells have reported cesium translocation rates measurements in yeast [441] and erythrocytes [442]. This strategy necessitates measuring the NMR signal before ¹³³Cs polarization relaxes: ¹³³Cs fast T1 relaxation (< 10 s) may limit the number of applications of ¹³³Cs hyperpolarization.

### 2.1.10. Common anions

Phosphate ions are treated in Section 2.3.2.

NH₄⁺ and NO₃⁻ are treated in the Section 2.3.3.

Cl⁻ is an abundant anion in cells, but its concentration can vary significantly depending on the culture medium, the cell type, the growth phase or the organelle: it can reach concentrations between 10 and 100 mM in bacterial and animal cells [443-447]. ³⁵Cl is the most abundant isotope of chlorine, and also the best adapted to NMR studies. It is however a spin 3/2 nucleus, whose quadrupolar nature has hindered cellular investigations, similar to what we have seen with ¹⁷O (Section 2.1.1.), ²³Na and ³⁹K (Sections 2.1.4. and 3.1.2). Since the 1970's, publications have shown that the ³⁵Cl signal is considerably broadened by its transient interactions with proteins [448,449]. A mean binding lifetime between 10 and 1000 nanoseconds had been calculated for Cl⁻ interacting

with hemoglobin. In these conditions, the electric field gradients felt by ³⁵Cl⁻ contacting proteins, combined with the slow protein tumbling, lead to T1 and T2 values of about 10 ms at only 4 g/L of BSA or hemoglobin. In cells, the NMR signal of ³⁵Cl⁻ was so broad that it was undetectable in early studies [450,451], while the extracellular ³⁵Cl⁻ generated sharp signals. Hence, ³⁵Cl NMR was used in the 1980's to measure cellular volumes as inferred from the intensity of the extracellular peak of ³⁵Cl⁻ [451-454]. In the early 1990's, ³⁵Cl⁻ NMR detection was reported from algae and tobacco suspension cells, probably from the watery vacuoles, and in erythrocyte ghosts (i.e. emptied cells) [455]. ³⁵Cl⁻ ions were finally observed in erythrocytes in the mid-1990's and a double exponential relaxation was measured (T2slow=1.2 ms, T2fast=0.09ms) [444], in agreement with the theoretical behavior of spins 3/2 interacting transiently with macromolecules. Using Co²⁺-chelating shift agents, the peak from extracellular ³⁵Cl⁻ was shifted sufficiently to be distinguished from the intracellular peaks in erythrocytes and fibroblasts, and the exchange of intracellular Cl⁻ with extracellular H₂PO₂ could be observed by ³⁵Cl NMR in erythrocytes [456,457]. The sensitivity was poor according to the authors and we have not found any other attempts to study intracellular Cl⁻ using ³⁵Cl NMR in the literature since the early 2000's. ³⁵Cl⁻ NMR has been explored for use in MRI in the 2010's: because it is very sensitive to the protein concentration, it could be useful to localize liquid accumulation upon strokes or edema [445,458-460]. The bi-exponential T2 rates of ³⁵Cl are in the millisecond range in these extracellular environments, and are still not very sensitive.

Altogether, the contribution of NMR spectroscopy to characterizing the intracellular behavior of Cl⁻ was not substantial up to now.

### 2.1.11. Viscosity, pH, redox potential

Intracellular viscosity is a complex, multifaceted property, and its measurement by NMR spectroscopy is treated separately in Section 3.2.1. Viscosity can be derived from two types of molecular mobilities, namely molecular tumbling and translational diffusion. These can both be determined by NMR, but they should be interpreted carefully: i) interactions with intracellular species can affect both tumbling and translation; ii) organelles, cytoskeleton, and cellular compartments cause restricted diffusion; iii) the apparent cellular viscosity is size-dependent, small and large molecules being differently retarded by the cellular meshwork.

In contrast, pH is a physical parameter that is usually very accessible by NMR spectroscopy: it needs a protonatable molecule carrying an “NMR-visible” isotope, whose chemical shift is intrinsically dependent on the protonation state [461]. Studies reporting intracellular pH values from an observed metabolite are too numerous to list. We mention a few below. ³¹P-NMR has been used to measure intracellular pH since the early 1970's [462]: it does not require any isotopic enrichment, and biological phosphate-containing molecules are numerous, commonly with pKa values around 6.5-7, so that their ³¹P chemical shifts are good pH sensors. Inorganic phosphate, DPG, ATP, etc. have thus been used regularly to monitor intracellular pH in animal, plant or microbial cells [463-465]. A few lines on this topic appear in Section 2.3.2, focusing on ³¹P-NMR monitoring of cellular metabolism. The recent commercial multi-channel probes designed for high-sensitivity ¹H and ³¹P detection are certainly interesting in this regard [465,466]. In addition, ¹H-, ¹³C-, ¹⁵N-NMR or ¹³C/¹⁵N-edited/¹H-detected NMR can faithfully report on pH through observation of metabolites that show pH-dependent protonation states with pKa close to 7. Notable examples include histidine (or imidazole-containing molecules), citrate, carnitine and bicarbonate [461,467-470]. However, ¹H-NMR pH-dependent chemical shifts are often problematic for real-time monitoring of cellular metabolism, because a number of peaks can shift over time and possibly cross or overlap (see
<figure><img src="image_14.png" /><figcaption>**Fig. 10.** A) Time series of ¹H HR-MAS NMR spectra (restricted spectral window) of *N. crassa* showing the variations of chemical shifts of citrate, succinate and glutamate, consistent with pH changing from 6.3 to 5.7 (adapted from Judge et al. 2019 [474]); B) NMR peak separation of GSH and GSSG at the position Cβ, ¹³C-labeling originates from ¹³C-cysteine supplementation in the growth medium of cultured mammalian cells (adapted from Mochizuki et al. [475]).</figcaption></figure>

Fig. 10). ¹⁹F-labeled probes have also been designed for intracellular pH measurements [461,471-473]. These can be convenient, notably because of the absence of ¹⁹F-background in cells.

Redox potential is measurable via the thermodynamic equilibrium of the main cellular redox couples NAD+/NADH, NADP+/NADH and GSSG/GSH (GSH= reduced glutathione). ¹³C-labeled NAD+/NADH, NADP+/NADPH have been observed in *E. coli* and yeast upon incorporation of ¹³C-labeled nicotinate [476,477]. The redox reaction of the nicotinamide moiety generates two sets of chemical shifts. Unfortunately, the chemical shifts are very similar in the NAD+/NADP+ (or NADH/NADPH) forms [477], so that monitoring the two couples correctly by NMR is not generally possible. The glutathione redox equilibrium has been monitored by ¹H-¹⁵N or ¹H-¹³C NMR after supplementing the culture medium with ¹⁵N-glycine [478] or ¹³C-cysteine [475,479], respectively: these two amino acids can enter cells and are incorporated in cellular glutathione. GSH and GSSG ¹H-chemical shifts overlap, therefore ¹⁵N or ¹³C-labeling is necessary to distinguish these two forms [475,478-480]. The ratio [GSSG]/[GSH]² enables measurement of a redox potential between ~250 and -100 mV, which corresponds to the intracellular variations between proliferative and apoptotic cells (~270 to -170 mV, respectively) [481]. However, these values should be interpreted cautiously: i) because the reaction occurs between GSSG and 2\*GSH, the equivalence in terms of redox potential is GSH concentration-dependent [481,482]; ii) intracellular concentrations of GSH vary between different organelles (concentration ranges from 1 to 15 mM, and redox potential down to -340 mV in mitochondrial matrix) [483,484]; iii) like all other redox couples in cells (measured by NMR or any other technique), the GSSG/2\*GSH ratio does not necessarily exactly reflect the real cellular redox potential, because spontaneous redox reactions are much slower than enzyme-catalyzed reactions in cells, hence the measured ratio may not correspond to a redox equilibrium [483,485]. The redox ratios of ¹³C-labeled ascorbic acid and dehydroascorbic acid have also been measured, but these compounds are constantly exchanged between the intra- and extra-cellular media [486-488].

NMR spectroscopy can thus provide information on a range of intracellular biophysical parameters. It reports averaged information on observable metabolites, with no subcellular details. Hence, it does not compete with up-to-date fluorescence microscopy approaches for functional investigations. However, NMR measurements of viscosity, pH and redox potential can be very valuable to monitor cell viability or overall populations of metabolites in real-time, possibly in parallel with other NMR characterizations.

## 2.2. Cell wall composition and dynamics

Plant, algae and microorganisms often establish dense polymeric layers surrounding their lipid membranes: these are carbohydrate-

rich networks, which help in shaping cells, in adapting them to environmental variations, and in escaping host immune defenses or pathogen attacks. The detailed characterization of biological carbohydrate-containing polymers can help for example in designing vaccines [489,490], in understanding pathological microorganisms features and capacities [491-496], or in exploiting their potential for medical and industrial purposes [497-499]. In most cases, the chemical composition of these polymers has been established since a long time, but the knowledge obtained from chemical extraction, purification and degrading chemical analysis has of course severe limitations. Supplementary techniques such as *in situ* NMR spectroscopy were necessary to learn about their three-dimensional structure, dynamics, packing, grafting ratios, degradation mechanisms and kinetics, interactions with antibiotics etc. We discuss below the contributions made by NMR spectroscopy of intact cells in understanding these coating polymers.

We will only briefly mention here the existence of NMR studies on biofilms [500-503]: although they are closely related to live-cell material and not truly inert, we classified biofilms as being out of the scope of this review.

### 2.2.1. Plant cell walls

Somewhat similar arguments could be given about the inert nature of plant cell walls, but we have included a more extensive discussion for them: solid-state NMR spectroscopy has made highly valuable contributions over the last 40 years in characterizing plant cell walls, using progressively more native material, including intact plant stems, in recent years.

Related studies also represented important steps in the development of NMR methods: it was with the aim of characterizing wood (together with polysulfone and ivory) that Stejskal and Schaefer reported the first use of Cross-Polarization Magic Angle Spinning (CP-MAS), a technique that has since had enormous importance for solid-state NMR [504,505]. This is a point not only of intense interest to spectroscopists: solid-state NMR spectroscopy revealed the different conformations of cellulose [506-514] and lignin [511,515-517] and how the crystalline and amorphous ratios were correlated to soft- and hard-woods [518,519]; it also helped in monitoring their evolution through rot, drying/rewetting and industrial processes [516,520-529].

The more mobile pectin polymers were also characterized from the mid-1990's in non-dried samples using various NMR strategies, either taking advantage of variable cross-polarization efficiency or ¹H relaxation times associated with different mobilities and hydration regimes [530-532], or using direct polarization (DP) MAS [533]. Solid-state NMR approaches progressively showed that pectins i) were distributed among different pools showing different mobilities, more or less associated to cellulose [534-539], and ii) became more mobile upon ripening of fruits while cellulose did not [540-543]. CP and DP are now regularly combined to achieve
a more integrated description of both rigid and mobile fractions of plant cell walls, respectively [544-546].

NMR studies on plant cell walls were carried out mostly on natural abundance samples until 2010. In the last 10 years, uniform ¹³C-labeling permitted comprehensive NMR signal assignments of plant cell wall polymers, notably by Mei Hong and her coworkers [547-550], and Kikuchi and his coworkers [551]. This led to renewed studies of the intermolecular contacts between cellulose, lignin and the complex hemicellulose and pectin heteropolysaccharides: indeed, it permitted detection of interatomic spatial contacts as revealed by magnetization transfer [552,553].

Very recently, this research became more “in-cell related” NMR spectroscopy with the analysis of intact material: using fresh ¹³C-labeled *Arabidopsis* stems, Dupree and his coworkers have shown that a specific, twofold helical population of hemicellulosic xylan binds to cellulose fibrils [554-556]; they observed the same xylan-cellulose interaction in spruce wood, showing also an additional binding of hemicellulosic galactoglucomannan to cellulose [557]. In a complementary fashion, using DNP-enhancement on intact ¹³C-labeled maize, rice, switchgrass and *Arabidopsis* stems, Wang and his coworkers showed in 2019 that lignin binds to the various populations of xylan through electrostatic interactions [558]. This last remarkable study also identified an impressive number of contacts between the different polymers and their level of interpenetration, which completely changed the accepted structural model of lignocellulose: lignin forms nanoaggregates and xylan bridges them with cellulose microfibrils.

Wang and Hong had also used DNP previously to characterize the trace occurrences of interactions between the lysis protein expansins and cellulose [559]. ¹H-detected fast MAS NMR spectroscopy also promises improved sensitivity in the future [550], so that advances may become achievable on sporadic but important connections [560], also on natural abundance material [553,561].

NMR spectroscopy has thus made fundamental contributions to both early and recent advances in understanding plant cell wall organization [560,562-565], which is of great value in solving the so called “biomass recalcitrance” to degradation for industrial purposes, including in the field of “green bioenergy” [566,567].

### 2.2.2. Peptidoglycans

NMR spectroscopic studies have also provided important information about bacterial peptidoglycan (PG), the branched glycan-peptide heteropolymer from which bacterial cell walls are composed [496,568]. Chemical structures of PGs have often been known for decades, but classical approaches use radiolabels or digestion and separation steps, which work better on Gram-negative bacteria PG: their Gram-positive counterparts are more difficult to solubilize, densely cross-linked and thus more difficult to analyze. Consistently, most NMR spectroscopy studies on PG have focused on Gram-positive bacteria using solid-state NMR. Several questions remain: the architecture of intact PG tridimensional is still under debate [568,569] and understanding interactions between PG and PG-targeted antibiotics is of vital importance [570-572].

To appreciate the contributions of NMR in the field, it is necessary first to present some elements of PGs' structure (Fig. 11). PG is based around a polymer built from a disaccharide repeated unit, namely a N-acetylglucosamine and a N-acetyl muramic acid, the latter being linked to a 4-5 amino acid stem [573,574]. Gram positive PG chains contain L-Ala, D-Ala, L-Gly, L-Lys, iso-Gln or diaminopimelic acid (DAP), which can be synthesized from L-Asp. Covalent crosslinks between peptide stems are established after integration in the PG layer through peptide bridges. Antibiotics can thus inhibit either the transglycosylases of the carbohydrate backbone, or the transpeptidases that establish crosslinks between

the peptidic stems. This can occur through enzyme inhibition or PG stems interaction and sequestration. Antibiotics can also capture lipid II, the precursor of PG, which contains the PG building block tethered to a pyrophosphoryl-undecaprenyl membrane anchor, whose copy-numbers are very limited in cells [575-577].

As early as 1977, Irving and Lapidot showed that the pentaglycyl bridge of *Staphylococcus aureus* PG was in a random coil conformation in intact cells, by using ¹⁵N-glycine selective labeling and ¹⁵N-detected solution NMR [580]. They used the efficiency of ¹H-¹⁵N magnetization transfer to characterize cell wall components' mobility. Importantly, they showed that PG mobility was higher in isolated *E. coli* cell-envelope than in whole cells, demonstrating a need for characterization of intact cells [581]. The first atomic-scale information was also obtained on the differentiated modes of action of the antibiotic vancomycin: it reduced the mobility of the surface polymer (teichoic acid) in *Bacillus licheniformis*, and that of the PG C-terminal D-Ala residue in *Micrococcus lysodeikticus* (which lacks teichoic acid) [582]. At that time, although providing only low sensitivity, ¹⁵N and its low gyromagnetic ratio were giving access to resolved solution NMR spectra of semi-mobile polymers, yielding distinguishable PG spectral signatures from species to species [583].

As for plant cell walls, the more solid fraction of the PG became observable using CP-MAS solid-state NMR [504,505]. And again as for plant cell walls, isotopic ¹³C/¹⁵N labeling was necessary to assign NMR resonances, identify covalent and spatial proximities between residues, and establish strategies to filter out the bacterial backbone. Such a rationale has been employed from 1983 by Schaefer and colleagues on *Aerococcus viridans*. In this bacterium, PG peptide stems show a lysyl-amide peak at 95 ppm when cross-links exist between lysine N-epsilon and D-alanine carbonyl, while non-bridged stems show a lysyl-amide peak at 5 ppm. By recording CP-MAS spectra on bacteria fed with ¹⁵N-lysine, ¹⁵N-NMR signals were observed for lysine ¹⁵N-epsilon in both free and D-Ala-linked states, which allowed the quantification of cross-linking upon exposure to penicillin [584]. This method was convenient and safe, avoiding extensive purification processes and the use of radiolabeled amino acids. It demonstrated that penicillin causes the production of larger amounts of peptidoglycan but with fewer cross-links [585], and that no C-terminal D-Ala or D-Ala-D-Ala existed in the vicinity of Lys-D-Ala cross-links [586].

Schaefer and his coworkers improved their approach from the early 1990's, by using their recent developments, such as DANTE ¹³C-¹³C and REDOR or TEDOR ¹³C-¹⁵N editing filters (see [570,587-591]). This improved both sensitivity and the removal of cellular background signals from natural abundance ¹³C and ¹⁵N [592,593]. Hence, a number of PG cross-links and bridge-links could be analyzed upon amino-acid specific ¹³C/¹⁵N-labeling, which produced ¹³C-¹³C and/or ¹³C-¹⁵N covalent bonds associated with specific chemical shifts. The degree of cross-linking between PG peptide stems was measured on isolated cell walls of *Bacillus subtilis* fed with [1-¹³C]-D-Ala and [1,2-¹³C-¹⁵N]-Asp as a precursor of DAP, revealing changes upon exposure to the antibiotic cephalotin [594-596]. Similar strategies and analysis have been carried out in the 2000's, using various D-Ala, L-Gly, L-Lys ¹³C or ¹⁵N-labeling schemes with *Staphylococcus aureus* and *Enterococcus faecium* to analyze the impact of antibiotics (vancomycin, amphotycin, plusbacin, oritavancin) on cross-links, bridge-links and stems [597-601], to detect PG lysine succinimidation [602], to characterize PG structures of *S. aureus* mutant strains [603,604] and in different growth phases and nutrient media [605].

Moreover, REDOR gave access to internuclear distance measurements [606]. This has been used fruitfully in the years 2000-2010, notably by combining ¹³C- and/or ¹⁵N-amino acid specific labeling and ¹⁹F-containing antibiotics. Indeed, ¹³C-¹⁹F, ¹⁵N-¹⁹F, and even ³¹P-¹⁹F distance measurements became feasible between
<figure><img src="image_15.png" /><figcaption>**Fig. 11.** A) Chemical structure of two PG stems from *S. aureus*, showing isotope labeling obtained from growing cells in a media supplemented with D-[1-¹³C]Ala and ¹⁵N-Gly and with an alanine racemase inhibitor. Other labeling schemes have been used, e.g. ¹⁵N- or ¹³C-Lys, ¹⁹F-side chain Lys, ¹⁵N-Ala, which permitted dissection of PG structures in various conditions (adapted from Cegelski et al. 2008 [578]). B) Chemical structure of TA, composed of polyribitol phosphate chains, whose ³¹P nuclei permit natural abundance ³¹P-NMR and ³¹P-REDOR filters; the scheme shows a possible D-[¹⁵N]Ala labeling pattern that has been used in the literature; anomeric carbons are indicated by red dots (adapted from Romaniuk et al. 2018 [579]). C) One-dimensional ¹³C and ¹⁵N CPMAS solid-state NMR spectrum of uniformly ¹³C- or ¹⁵N-labeled *S. aureus* cells treated with (red) or without (black) 1 µg/mL tunicamycin. The resolved peaks that permit the quantification of peptidoglycan (PG) and wall teichoic acid (WTA) are shown with arrows (adapted from Romaniuk et al. 2018 [579]).</figcaption></figure>

PG residues and the antibiotics, and between bacterial phospholipids (exploiting the 100% natural abundance of ³¹P) and the antibiotics. This helped Schaefer and his coworkers to progressively elucidate the interaction modes of ¹⁹F-labeled derivatives of antibiotics (vancomycin, oritavancin, eremomycin, chloroeromomycin, Plusbacin A3) with *S. aureus* or *E. faecium* PGs [578,600,607-611] (see for example Fig. 12). In complete contrast to a widely accepted belief amongst the community from earlier in vitro studies [612-614], vancomycin and oritavancin were never found to form dimers *in situ* according to REDOR analysis [578,579,607,609,611]. Deuteration of the lipid moiety of Plusbacin A3 also permitted ²H-¹³C and ²H-³¹P REDOR distance measurements, showing that this lipid side chain was essential for the antibiotic action, but did not insert itself into the bacterial lipid membrane [615].

To finally derive a model of the tridimensional PG architecture, Schaefer and colleagues had to create one more solid-state NMR experiment called CODEX, which was designed for measurement of ¹³C spin diffusion, giving access to ¹³C-¹³C distance information between pentaglycyl bridge and the *S. aureus* PG glycan moiety in isolated intact cell walls [616]. In the 2010's, the introduction of L-[5-¹⁹F]-lysine together with use of D-¹³C-Ala, L-¹³C-Ala, L-¹⁵N-Ala or L-¹³C-Gly in the culture medium generated PG in which ¹³C/¹⁵N-¹⁹F labels were combined so as to facilitate use of REDOR to derive distances in intact cell walls of *S. aureus* and some mutant strains [589,617,618]. These combined studies allowed advanced models of the PG mesh of *S. aureus* to be established [574].

NMR of whole bacterial cells also helped to characterize teichoic acid (TA), another important surface polymer of bacterial cell walls. TA contains D-Ala residues, which had been observed to be quite mobile by ¹⁵N-NMR in the late 1970's [582,583]. However, TA D-Ala is sufficiently constrained to allow solid-state cross-polarization and to yield ¹³C-CP-MAS spectra, but its ¹³C resonance is severely overlapped with signals of other D-Ala residues in peptidoglycan [598]. Spectral deconvolution permitted comparison of the TA-associated D-Ala content in different species and upon exposure to antibiotics [589,598,615,619,620]. In contrast,

¹⁵N-NMR peaks of D-Ala from TA and PG are well separated, and also helped to monitor TA D-Ala in various conditions [589,599,620,621]. Uniform ¹³C- and ¹⁵N- labeling was used recently to evaluate the contents in TA ribitol and D-Ala in intact cells (Fig. 11) [579]. This represents a convenient approach for rapid characterization of antibiotic mechanism. Finally, ¹³C-¹⁹F or ¹⁹F-³¹P REDOR filtering proved the absence of contact between the antibiotics ¹⁹F-oritavancin or ¹⁹F-Plusbacin A₃ and TA [600,611].

Hence, in their very extensive work on the characterization of PG and the impact of PG-interacting antibiotics, Schaefer and colleagues provided an impressive body of results, but also very significant developments of solid-state NMR approaches. However, to the best of our knowledge, we must note that these NMR experimental advances are not so well integrated into the common toolset for PG analysis [568,572,622]. This is probably due to the sophisticated NMR instruments and the highly advanced expertise that are required. Also, the poor sensitivity of NMR hampers the study of rare, but functionally relevant events that affect PG upon antibiotic exposure, for example.

Such issues may be solved in the future by ¹H-detected solid-state NMR, which has been carried out *in vitro* on purified PG sacculi of *B. subtilis* [623], or by DNP-enhancement using intact cells [624]. These two techniques have already led to proposals of structural models for the interaction between two antibiotics (nisin and teixobactin) and the essential, low-copy number lipid II molecule, using bacterial membrane extracts [625,626] (see Section 2.4.2 for more details).

The carbohydrate moieties of PG and cell wall teichoic acids have not been extensively studied by in-cell NMR, to the best of our knowledge. NMR spectroscopy of carbohydrates is not straightforward, because most of the carbohydrate resonances are in crowded regions of the spectra, except those from the anomeric protons and carbons. Appropriate carbohydrate isotopic labeling may help to observe the GlcNAc and MurNAc moieties contained in PG and TA.
<figure><img src="image_16.png" /><figcaption>**Fig. 12.** Structural information obtained from REDOR distance measurements in S. aureus whole cells incubated with the ¹⁹F-labeled glycopeptide drug LCTA-1110. Top right: Schematic cross-section of the PG tertiary structure of S. aureus (grey balls: GlcNAc-MurNAc glycan backbone; green rectangles: PG stems; red rectangles: PG bridges; LCTA-1110 is represented by a grey cartoon); Top left: ¹³C-¹⁵N, ¹³C-19F, and ¹⁵N-¹⁹F measured distances in S. aureus grown in a medium containing D-[1-¹³C]Ala and L-[¹⁵N]Ala; Bottom left: chemical structure of LCTA-1110; Bottom right: Model of the interaction between LCTA-1110 and PG based on measured REDOR distances. Adapted from Kim et al. 2013 [589].</figcaption></figure>

### 2.2.3. Lipopolysaccharides and beta-glucans

Cell wall polymers of higher flexibility such as bacterial or yeast polysaccharides are also observable by NMR in intact cells. The so-called High-Resolution Magic-Angle Spinning (HR-MAS) NMR technique [627-631] has been instrumental in characterizing polysaccharides in situ, allowing their analysis without any solvent extraction or separation steps, during which information could be lost [632,633].

Because of their density in the membrane context, these polysaccharides form a loose meshwork that restricts molecular motion to some extent. Added to the sample heterogeneity between local environments (intracellular, extracellular and membrane), this restricted mobility causes broad NMR signals for surface polysaccharides. This prevents a precise, atomic-scale characterization using classical liquid-state NMR. Surface polysaccharides still conserve high degrees of local mobility, which can generate sharp NMR peaks once the sample is rotated at the so-called magic-angle (54.7°). Restricted motion and sample heterogeneity do cause broadening effects, which are observable and can be assessed in the shape of NMR peaks in non-spinning conditions (see Section 3.2.2. for more extensive explanations). Magic-Angle Spinning (MAS) averages out these effects when spinning rates are faster in Hz than the observed deleterious peak broadening, i.e. a few hundreds of Hz for cell surface polysaccharides. The resulting high-resolution spectra obtained under MAS gave rise to

the acronym HR-MAS (see Section 3.3.4. for specific explanations on HR-MAS).

Surface polysaccharides are often produced in high quantities and in rather homogeneous populations by microorganisms, so that their NMR signals surpass those of the other cellular species. ¹H-HR-MAS permits the detection and analysis of surface polysaccharides from milligram quantities of live bacteria (~10-50 mg wet weight, ~1 mg dry weight), thus avoiding the need for large scale production and supplementary purification [634]. The HR-MAS NMR analysis of surface polysaccharides in live cells allows observation of the effects of drug exposure, environment, growth phase and gene mutations on microorganism cell walls, while requiring only moderate manipulations of the biological material. HR-MAS requires spinning the samples at kHz frequencies in cylindrical rotors, whose diameters are in the millimeter range. This generates forces that do not affect microorganism viability [635] but can often damage mammalian cells or tissues [636]. We consider HR-MAS NMR studies on mammalian cells further in a later section, which focuses on real-time monitoring of live-cells metabolism (Section 2.3.5.).

¹H-HR-MAS has been applied from the late 1990's to verify the integrity of the extracted and purified polysaccharides that were used for "structural characterization" (in the carbohydrate community, this corresponds to the equivalent of protein primary structure determination, i.e. finding the sequence and linkage of
monosaccharide units). We should mention that ¹³C-HR-MAS has also been used in one publication to study the three-dimensional features of cell wall glucans from *Saccharomyces cerevisiae* [637]. Kenne and coworkers applied ¹H-HR-MAS to Gram-negative, opportunistic pathogenic bacteria to establish the structure of their surface-exposed O-specific polysaccharides [638,639]. At the same time, Wieruszeski, Lippens and their coworkers showed that the osmo-regulated cyclic glucans of a plant pathogenic bacterium i) adopted the same three-dimensional arrangement in vitro and in live cells and ii) had reduced diffusion in their native crowded environment, i.e. the periplasm [640,641].

The ¹H-HR-MAS approach was next applied to the analysis of *Campylobacter jejuni*, an infamous gastrointestinal pathogen: structures of its capsular polysaccharides and lipo-oligosaccharides (LOS) were compared upon gene mutation [642,643], between clonal variants [644,645] and strains [646,647]. ¹H-HR-MAS similarly permitted elucidation of the structures of the surface polysaccharides from Gram-negative [648-651] and Gram-positive bacteria [652], mycobacteria [653], and yeast [654]. These studies were carried out either to detail the function of genes involved in polysaccharide biosynthesis, or simply to verify the absence of degradation during the purification steps used for other analyses. Kinetics of acid incorporation [655], growth-phase and pH-dependent O-acetylation [656,657] were also monitored, which may help to understand the adaptation of bacteria to their environment.

More generally, all these studies have shown that microorganism surface polysaccharides conserve similar three-dimensional structures and a high conformational flexibility when isolated or in the native membrane meshwork. This is an important, fundamental piece of information.

### 2.2.4. Lipid dynamics in membranes

We address here NMR studies focusing on lipids in cellular membranes. Approaches aiming at measuring transmembrane diffusion are considered in Section 2.1, and measurements of cytoplasmic lipids are mostly treated in Section 2.3.5.

¹H-, ²H-, ¹³C-, ¹⁹F- and ³¹P-NMR spectroscopy was applied to study lipid dynamics in the early 1960's, complementing other spectroscopies already in use (Raman, Infrared, light scattering methods, fluorescence probes, spin label probes for EPR spectroscopy...) [658,659]. Early ¹H NMR spectra of erythrocytes, mitochondria and sarcoplasmic reticulum membranes had revealed that only the high mobility choline head group was observable, while signals of the acyl chains were extremely broad in the absence of detergent [660-663]. Hence, lipid dynamics in cellular membranes were first investigated by ¹³C NMR because of its favorable relaxation characteristics. The first ¹³C NMR spectra of native membranes by Metcalfe and coworkers in the early 1970's showed very broad signals, either from human erythrocytes or from the small bacterium *Acholeplasma laidlawii* (a species devoid of cell-wall and thus very practical for lipid membrane studies) [664]. The information on lipid dynamics came first from the characterization of membranes among the most fluid membranes, i.e. those of the sarcoplasmic reticulum [665] and the retinal rod outer membranes [666], which contain high proportions of unsaturated acyl chains. These studies reported ¹³C (natural abundance) T1 values for the acyl chains similar to those measured for purified lipid vesicles, while ¹H NMR signals were very broad. Hence, native membranes probably had similar viscosities, but produced more complex environments due to the presence of membrane proteins [659]. Similar conclusions came from ¹³C NMR studies on the choline head group of less fluid membranes from virions or cultured cells [667,668]. The same ¹³C signal from choline head groups was also used to monitor the distribution of ¹³C-labeled phosphatidyl-choline in the inner- and outer- leaflet of sarcoplas-

mic reticulum samples purified from rat muscles [669,670]; the addition of a shift reagent (here Dy³⁺, see Section 2.1.2. for further explanation of shift reagents) in the extracellular medium permitted the distinction of choline groups pointing towards the intra- or extra-cellular sides of the membrane.

Hence, ¹³C T1 relaxation and linewidth analysis was providing scarce and rather qualitative information on cellular lipid membrane dynamics. Moreover, strong dipolar interactions and magnetic susceptibility gradients were producing broad ¹H NMR signals, which were also poorly informative. ³¹P-NMR has been extremely useful for *in vitro* studies on lipid phases, because it yields very different spectra from bilayers, hexagonal, vesicular and micellar phases, without any need for isotopic enrichment [671,672]. It has been much less informative for intact cellular membranes [673-681], for several reasons: i) there is no need to prove they are bilayers; ii) different phosphate ester functions have overlapping resonances; iii) intense signals from cellular phosphate-containing species (ATP, etc..) can obscure the broad signals from membranes.

With the advent in the mid-1970's of pulse schemes (namely the quadrupolar echo) adapted to the measurement of ²H quadrupolar splittings [682], more quantitative measurements of cellular membrane dynamics could be achieved. Indeed, the quadrupolar nature of the ²H nucleus revealed itself to be extremely useful here (Fig. 13) [683,684]: i) the quadrupolar peak splitting is related to the order parameter of the carbon-deuterium bond vector $S_{CD}$, ii) it is very large, spanning up to dozens of kHz in the case of restricted dynamics, iii) it is usually wider than the ²H linewidth (the low ²H gyromagnetic ratio results in weak dipolar interactions and low sensitivity to field inhomogeneity).

Because of the simplicity of its single membrane and the capacity to produce it at gram scales, the bacterium *A. laidlawii* became an interesting model for ²H NMR investigations. In the early 1970's, perdeuterated or 16-²D₃ (tri-deuterated at the end of the acyl chain) palmitate was successfully inserted in *A. laidlawii* membranes by supplementation in the growth medium [689,690]. The end-chain ²H-labeling permitted measurements of its ²H residual quadrupolar splitting in freeze-dried, rehydrated *A. laidlawii* membranes, giving access to the end-chain order parameter [690]. Repeating this experiment with specific ²H-labeling at every position of the acyl chain successively, Stockton and colleagues reported the order parameter through the entire thickness of intact cellular membranes [691]. This revealed a behavior very similar to that observed with single phospholipids in the lamellar liquid crystalline phase at 42 °C. Similar measurements with decreasing temperatures showed the progressive shift of an equilibrium between liquid crystal and gel phases: the two populations exchanged with rates lower than 10⁻⁴ Hz, and the *A. laidlawii* membrane showed an almost complete motional freezing at 1 °C (micro-millisecond timescale) [688]. The impact of cholesterol insertion on *A. laidlawii* membrane dynamics was also evaluated, as well as the dynamics of saturated fatty acids and those of various fatty acid headgroups incorporated into the same membranes [687,692-695]. *Mycoplasma caprilocum*, which is naturally dependent on fatty acids and cholesterol supply, was also analyzed by ²H-NMR: the intact cells had membrane dynamics similar to those of model lipid membranes, and only those grown under low cholesterol supplementation showed a liquid-crystalline-gel phase transition [696]. All these results were very consistent with those obtained from purified lipids, and thus proved the suitability of *in vitro* approaches in the field [697]. It also emphasized that membrane proteins had little effect on average membrane lipid dynamics.

Similar studies were carried out almost in parallel on *E. coli* in the late 1970's. Deuterated lipids were incorporated in auxotrophic *E. coli* strains defective in certain lipid production steps. The ²H-NMR analysis was essentially performed on membrane vesicles,
<figure><img src="image_17.png" /><figcaption>**Fig. 13.** Principle of ²H-NMR quadrupolar splitting analysis. The chemical bond C-D is represented with carbon as a grey ball and deuterium as a blue ball. Most of the original work described in the main text was carried out following the rationale presented here. However, a new approach taking advantage of Magic-Angle Spinning (MAS) has been proposed in recent years, which offers better sensitivity [685]. A) The ²H nucleus has a spin I=1, giving rise to two different energy transitions due to the electric quadrupolar interaction, hence two resonance frequencies $v^+$ and $v^-$; moreover, these two quadrupole frequencies are dependent on individual spin orientations, and the density of orientation states determines a powder pattern for a non-oriented sample (with the typical “Pake doublet” shape); B) The residual quadrupolar splitting $\Delta v = v^+ - v^-$ is proportional to the square of the order parameter (adapted from Molugu et al. Chem Rev 2017 [686]); C) Oleic acid deuterated at various positions was incorporated in the small bacterium *A. laidlawii* and position-specific quadrupolar splitting of oleyl chains measured in intact membranes at 25 °C (adapted from Rance et al. 1980 [687]); D) Perdeuterated palmitic acid was incorporated in *A. laidlawii* and palmitoyl chains observed by ²H-NMR at varying temperatures, showing progressive loss of the powder pattern due to the liquid crystalline phase (adapted from Smith et al. 1979 [688]).</figcaption></figure>

and showed that i) these membranes had a similar dynamic finger-
print to that of model liposomes, although they were 10-20% less
ordered probably because membrane proteins slightly “destruc-
ture” the liquid-crystalline lipid bilayer [698-701], ii) *E. coli* mem-
branes were >97% in liquid crystalline phase at 37 °C. The inner and
outer membranes were also separated in two samples, which
revealed that the inner membrane was less ordered and had a 7
°C lower liquid-crystalline-gel transition than the outer one
[700,702,703]. In the 1980's, deuterated DPPC was incorporated
by incubation with erythrocyte ghost membranes: similar dynam-
ics were observed in these membranes and in pure DPPC vesicles,
which also showed no transition to the gel phase down to 5 °C
[704]. Brains of rats fed with deuterated choline headgroups were
also analyzed by ²H-NMR, revealing similar lipid dynamics in
intact tissues to those in model vesicles [705]. Later ²H-NMR stud-
ies on intact *E. coli* cells in the 1990's showed that acyl chains in
their membranes were indeed slightly less ordered than in model
liposomes [680] and that glycerol or trehalose had little effect on
these dynamics [681]. More recently, in the late 2000's, similar
conclusions were drawn from ²H-NMR studies of intact sea urchin
sperm cells incubated with deuterated phospholipids [706].

Since the 2010's, some applications of ²H-NMR have been
reported that focused on the effects of antimicrobial agents on bac-
terial membranes in intact cells. Even at sublethal concentrations,
some antimicrobial peptides were shown to cause a decrease of
order on the acyl chains of deuterated lipids, which were previ-

ously integrated by a lipid-auxotrophic *E. coli* [707]. In contrast,
other antimicrobial agents provoked the appearance of a gel phase
in intact *E. coli* cells similarly supplemented with deuterated lipids
[708]. In this latter study, Marcotte and coworkers managed to
integrate deuterated fatty acids from the growth medium into a
wild-type *E. coli* strain, where they were incorporated in phospho-
lipids. Magic-Angle Spinning (MAS) was finally used on such a sys-
tem only in 2015, to the best of our knowledge: ²H-MAS NMR
spectra of a cell pellet (about 80 µL) acquired in 10-20 minutes
gave access to the order parameters of *E. coli* membranes, with
the same precision as without MAS but ten times faster [685].
Warschawski, Marcotte et al. could show that *E. coli* membranes
were mostly in a gel phase when grown only with saturated palmi-
tic acid, while the addition of unsaturated oleic acid ensured a fluid
phase at every temperature. Deuterated palmitic acid was again
incorporated in the Gram-negative *E. coli* and in the Gram-
positive *Bacillus subtilis* to compare their reaction to antimicrobial
peptides: disordering effects of these peptides occurred at different
concentrations on the two bacterial strains [709,710]. The same
authors incorporated deuterated palmitic acid in the pathogenic
marine *Vibrio splendidus*, and could show that its membrane fluid-
ity varied through the different growth phases, dependent on the
saturated/unsaturated lipid content [711].

Altogether, NMR spectroscopy has been instrumental in quanti-
fying the dynamic features of membranes at the atomic scale, and
in assessing their validity in intact cells, notably using ²H-NMR
[686,712]. It established fundamental knowledge in the 1970-80's, using methods that required rather long acquisition times and often membrane separation. The recent implementation of deuterated lipid incorporation and of MAS has created new opportunities: the functional monitoring of membranes by NMR becomes faster and more straightforward. This may help in mechanistic studies of membrane binders like antimicrobial agents, for example [713,714].

## 2.3. Metabolic activity

We will not address metabolomics of body fluids and plants. This is now an immense field in its own right [38-46], which mostly deals with samples extracted from the studied organisms or tissues. Still, a number of developments have focused on measuring metabolome evolution of living biological material in an NMR spectrometer, and the corresponding experimental procedures and technical possibilities are typically in the scope of this review.

Magnetic field inhomogeneities and slow molecular tumbling generally hamper NMR spectroscopy, and these are exactly the conditions met in cells. As shown below, it was soon recognized, in the early 1970's, that ¹³C- and ³¹P-NMR were likely to yield analyzable spectra of free metabolites in cells. This was possible because these were fast tumbling species showing slow NMR relaxation and thus sharp NMR signals, even though broadened by sample inhomogeneities and the multiple, transient interactions with cellular species [715-717]. Due to their low cell mobility, macromolecules like proteins [715] or DNA [716] appeared at that time to be intractable, yielding very broad, low intensity resonances. The observation of low gyromagnetic ratios nuclei like ¹⁵N represented a possible solution for obtaining narrow line widths, because they are less susceptible to dipolar line broadening and thus generate slower relaxation. Although providing the highest sensitivity, ¹H NMR became useful only later, once specific editing strategies and techniques to narrow ¹H signals were available. Every nucleus is thus adapted to the observation of different metabolites and different concentrations or timescales. We discuss these various aspects below.

### 2.3.1. The early days: studying carbon fluxes using ¹³C-NMR

In the early 1970's, different groups had progressively established the use of ¹³C labeling as a powerful method to decipher metabolic pathways and carbon fluxes: cells were grown on natural abundance (98.9% ¹²C, 1.1% ¹³C) media supplemented with precursors carrying a ¹³C atom at a specific position; products of interest were extracted, isolated and analyzed by ¹³C-NMR spectroscopy, which permitted identification of the ¹³C-labeled final positions [718,719]. This relied on the scarce ¹³C natural abundance and the approximation that all detected ¹³C nuclei found their origins in the supplemented ¹³C-precusors. It was also possible because the cellular ¹³C-signal was low enough to generate simple, low background spectra.

Soon after the observation of dissolved ¹³CO/¹³CO₂/¹³CN in erythrocytes [720,721], Matwiyoff and colleagues established the feasibility of observing global metabolites using one-dimensional ¹³C-detected NMR of Neurospora crassa grown with ¹³Cε-Methionine [722], or of Candida utilis grown on ¹³C-labeled acetic acid [715]. In this latter publication, the authors also traced ¹³C atoms from [1-¹³C]-glucose, showing the concomitant disappearance of the anomeric ¹³C-peak and the appearance of the C2 signal of ethanol, followed later by signals of unknown species. Hence, the weak natural abundance of ¹³C (~1.1%) made isotopic enrichment necessary for carbon metabolism studies, but it was immediately recognized as an advantageous feature allowing the use of ¹³C-tracers of metabolic activity: introducing a ¹³C-labeled compound permitted

¹³C-detection of the successive metabolic products in a mostly ¹²C-containing background. Only small, fast-tumbling molecules were NMR observable, even though they showed broader line widths than in vitro. But they were even observable in living animals: in 1981, in vivo ¹³C-detected NMR of the rat abdomen on feeding with [1-¹³C]-glucose was used to demonstrate its progressive conversion into [1-¹³C]-glycogen by recording spectra every 30 minutes [723].

Shulman and coworkers took advantage of these capabilities, reporting in 1978 a real time NMR study of the first steps of glycolysis in E.coli, observing the progressive ¹³C transmission from [1-¹³C]-glucose to fructose biphosphate, lactate, succinate, acetate and amino acids [724]. Less than 5 years later they published similar studies focusing on gluconeogenesis in rat liver cells exposed to [1,3-¹³C]- and [2-¹³C]-glycerol [725,726] or to [3-¹³C]-alanine and [2-¹³C]-ethanol [727], on S. cerevisiae anaerobic glycolysis from [1-¹³C]- or [6-¹³C]-glucose [728] or aerobic metabolism of [1-¹³C]- and [2-¹³C]-acetate [729], and on the consumption of endogenous [3-4-¹³C]-trehalose and exogenous [1-¹³C]-glucose by Pichia pastoris spores upon germination [730,731]. To illustrate the variety of organisms and metabolic phenomena that ¹³C-NMR was able to tackle, in the meantime, Scott and colleagues characterized ¹³C redistribution in the biosynthesis of the more complex molecules like porphyrinogens or polyketides in living Rhodobacter sphaeroides or Penicillium urticae [732-734] and glucose catabolism in tsetse fly parasites Trypanosoma brucei [735]; Nicolay and his coworkers had monitored acetate metabolism in photosynthetic bacteria depending on light conditions [736,737]; Ashworth and colleagues had traced ¹³C nuclei incorporation from glycine, serine or glucose into serine, tryptophane and acetate by tobacco suspension plants or bacteria (using devices to ventilate the NMR sample or to pump fermenting agents in and out of the NMR tube from an extra-spectrometer fermentor) [738,739].

Then, London and colleagues found an interesting application in the measurement of oxidized and reduced populations of intracellular electron carriers NAD+/NADH and NADP+/NADPH [476,477]. These metabolites were ¹³C-labeled via the incorporation of ¹³C-nicotinate in E. coli and S. cerevisiae, and their reduced and oxidized states yielded different ¹³C-NMR signals. These enabled the simultaneous observation of a global redox ratio, but peaks from NAD+ and NADP+ or from NADH and NADPH could not be distinguished, the nicotinamide moiety being too far from the adenine phosphorylation site to result in resolvable shift differences.

Concerning microorganisms, we mention works on amino acid biosynthesis by Corynebacterium glutamicum, a bacterium used for industrial production of amino acids [740-742], glucose metabolism of the pathological S. aureus [743,744], of the yeast S. cerevisiae [745-748], of rumen bacteria [749-751] and of Lactococcus lactis [752-754], formaldehyde [755] and acetate [756] utilization by E. coli, and acetate, methanol or methylamine utilization by methylotrophic strains [757-760].

The early cellular studies paved the way towards future ¹³C studies carried out in vivo [761,762], which included studies of glucose metabolism of erythrocytes [763,764], acetate and glucose utilization in lymphocytes and breast cancer cells [765,766], the effects of 2-deoxyglucose in cancer cells [767,768], amino acid [769] and succinate [770] metabolism by renal cells, and glucose and glutamine metabolism by hybridomas [771]. After early reports on ethanol and alanine metabolism in perfused mouse livers in the late 1970's [725,772], ¹³C-MRS studies (i.e. NMR characterization of localized chemical composition and metabolism) included investigations of metabolism of plant seeds, brain, tumors, heart, but were limited by their low sensitivity [773-775]. Hyperpolarization can nowadays overcome these limitations in some cases, as we discuss in a later paragraph.
However, spectroscopists realized in the late 1980's that, for most ¹³C-labeled precursors, their insertion in intricate metabolic cycles resulted in the progressive scrambling of ¹³C atoms into numerous, low abundance species (for a thorough discussion, see [776]). Together with the low sensitivity of ¹³C-NMR and its low spectral resolution due to sample inhomogeneities resulting in broad signals, *in vivo* ¹³C-NMR was limited in its ability to carry out a fine analysis of metabolic pathways. Moreover, high quantities of ¹³C-labeled precursors necessary for real time NMR detection impacted cellular metabolism [776]. Among other difficulties, distinguishing NMR signals of intra- and extra-cellular populations of molecules was also a common concern: unlike those of ³¹P as we will see below, ¹³C chemical shifts often depend too weakly on pH or on the other phenomena that can vary between intra- and extra-cellular spaces [776]. Another concrete obstacle was the problem of maintaining cells in steady-state conditions. Indeed, many parameters influencing metabolism become unstable at the high cell density needed to obtain sufficient NMR signal-to-noise, e.g. oxygenation, nutrient levels, pH, osmolarity, (toxic) end products [740,776]. A number of bioreactors inside spectrometers were designed at that time (see Section 3.7.). These permitted continuous monitoring of cell metabolism, notably to characterize the impact of drug-treatment and irradiation on mammalian cancer cell metabolism by ¹³C- and ³¹P-NMR [768,777-794], or to study the effects of ethanol and various chemicals on hepatocytes [795,796], as well as metabolism in hybridoma [771,797]. However, in-cell NMR studies of metabolic activities appeared to be limited to cases involving the study of high concentration metabolites. It is striking that all these impediments to live cell ¹³C-NMR metabolomics were also faced 25 years later by the in-cell structural biology community.

Hence, in combination with gas- and liquid-chromatography mass spectrometry (GC-MS) since the early 1990's, NMR spectroscopy of extracted and isolated materials delivered the main body of atomic scale information on metabolic pathways for the so-called Metabolic Flux Analysis [39,762,798-800]. However, when adapted, specific advantages of real time measurements, like using a single sample and avoiding sample preparation, remained attractive at times to characterize simple metabolic transformations.

### 2.3.2. ³¹P-NMR of phospho-species, energy fluxes and cellular localization

Studies of ¹³C-NMR were early combined with those of ³¹P-NMR, the latter providing interesting information on cellular metabolic activity [746,801,802]. ³¹P was considered to be easier to detect by NMR because of its ~100% isotopic abundance, and one-dimensional ³¹P NMR spectra were reported from red blood cells a few months after the ¹³C-experiments described above [462]. Direct observation was thus possible of highly abundant molecules such as inorganic phosphate, ATP and diphosphoglycerate (DPG), which are important cellular metabolites. Hence, ³¹P-NMR showed early promise for monitoring metabolic activities or mechanisms. Large DNA and RNA molecules also contain significant amounts of ³¹P, but these are mostly undetected by solution NMR because of their slow tumbling, or by solid-state NMR because of their heterogeneity.

³¹P in phosphate groups often shows large chemical shifts variations with pH, and in the early 1970's DPG signals were proposed as useful pH reporters in erythrocytes [462]. DPG was also quantified in real time using ³¹P-NMR peak intensities upon inosine and pyruvate supplementation of erythrocytes [803]. Salt concentrations, notably [Mg²⁺], were later shown to also have an influence on ³¹P-phosphate pKa and chemical shifts [804,805]. The use of ³¹P-NMR for reporting on ATP or DPG binding to Mg²⁺ and quantifying cellular free Mg²⁺ concentration is extensively discussed in

Section 2.1.6. Suffice to mention here that about 90% of intracellular ATP is found to chelate Mg²⁺ in a fast exchange equilibrium.

Of course, it became apparent that tissue metabolism could be observed via the detection of the ³¹P NMR signals of cellular phosphate moieties *in vivo*: ³¹P-NMR studies of rat muscle showing well-resolved signals from phosphocreatine, ATP and inorganic phosphate were reported in 1974 [311]. This prompted ³¹P-NMR studies on the evolution of inorganic phosphate/ATP/ADP/phospho creatine levels in rat hearts [806,807], rat kidneys [808] and mouse livers [809] subjected to ischemia and frog muscles under fatigue [810-812], completed with intra-tissue pH monitoring using ³¹P-NMR [812-814]. The stage was thus set for *in vivo* ³¹P-MRS/MRI, which was developed from the late 1970's [815-818] also on human patients [819,820]: it has contributed important knowledge, notably on metabolism of healthy or diseased organs, under rest or effort, but such studies are out of the scope of this review; dedicated reviews on this field exist [821,822].

In parallel, ³¹P-NMR on whole cells yielded important advances in the description of intracellular organization. It clarified mechanisms of polyphosphate storage and its use in yeast [823,824], and permitted quantification of phosphate-containing metabolites, including ATP/ADP/NAD and phospholipid precursors (phosphoryl-choline and phosphorylethanolamine) in lymphoid, Friend erythroleukemia, and HeLa cells [716,825]. pH Measurements by ³¹P-NMR also showed multiple intracellular ³¹P-phosphate NMR peaks, revealing the existence of different intracellular phosphate pools: this permitted distinction between cytosolic and mitochondrial pH in rat liver cells [826], cytosolic and vacuolar pH in maize root tips [827-829], and cytosolic and vacuolar pH and phosphate stocks in various yeast strains [737,824,830,831] including in spores [832]. Similarly, ³¹P-NMR analysis of intact weeds showed that glyphosate-resistant species can store this herbicide in vacuoles and thus elude its inhibitory capacities [833-835].

Real time pH monitoring also became accessible, and gave rise to reports on i) ATP metabolism upon oxygenation or deoxyglucose supplementation in Ehrlich ascites tumor cells [836], ii) intracellular and extracellular pH under aerobic or anaerobic conditions in *E. coli* [837-840], and iii) light-induced pH variations in photosynthetic bacteria [841,842]. NMR spectrometers gained a reputation of being the most expensive pH meter in the world [461], but the non-invasive nature of the method was certainly appealing. Moreover, complex metabolic studies were also carried out on glycolytic mutant strains of *S. cerevisiae*, by reporting multiple phospho-metabolite levels upon addition of glucose or O₂ [843]. This demonstrated the ability of NMR to report multiple metabolite levels in order to support more system-wide characterizations.

Still more informative ³¹P-NMR experiments were developed once Shulman and colleagues showed that reaction rates could also be measured between orthophosphate and ATP in *E. coli*: upon saturation of either the resonance of ³¹Pγ-ATP or of phosphate, they observed peak intensity losses of phosphate or of ³¹Pγ-ATP, respectively [844]. Hence, it was not only possible to monitor phosphate and ATP levels, but also to measure rates of ATP hydrolysis or synthesis. This “saturation” method, developed in the 1960’s [845,846], relies on the selective destruction of magnetization of a chosen resonance, i.e. corresponding to a chosen atom in a chosen species, which thus acts as a tracer of chemical exchange: the saturated atoms transfer their loss of magnetization to the product of the studied chemical reaction, whose NMR resonance intensities are attenuated proportionally to the reaction kinetic rate (Fig. 14). Saturation approaches have later been used in ligand-observed NMR spectroscopy of ligand:protein interactions, as described in Section 2.5 on drug discovery.

Resonance saturation also underlies recent Chemical Exchange Saturation Transfer (CEST) studies for advanced protein conforma-
<figure><img src="image_18.png" /><figcaption>**Fig. 14.** 1D $^{31}$P NMR spectra without (grey) and with (black) saturation of the γ-phosphate of ATP at -2 ppm. $^{31}$P signals can be attenuated upon γ-ATP saturation either because i) they are close in space to γ-ATP (i.e. β- and α-ATP), or because ii) they are in chemical exchange with γ-ATP (i.e. phospho-creatine or phosphate). The signal intensities in these peaks provide quantitative information on the kinetic rates of the chemical reactions. Adapted from Befroy et al. 2012 [847].</figcaption></figure>

tional studies, notably on “invisible” minor species [848,849]. In the case of living cells, $^{31}$P-NMR saturation transfer was applied in the 1980's to quantify rates of inorganic phosphate or ATP consumption in E. coli [850], in human erythrocytes [851], in breast cancer cells [852] and also the fluxes towards glucose-6-phosphate or fructose-1,6-bis-phosphate in aerobic and anaerobic conditions in S. cerevisiae [746,853-855]. In parallel, a number of groups used $^{31}$P-resonance saturation transfer to measure ATP and phosphocreatine consumption and replenishment, hence measuring creatine or ATPase kinase activity in rabbit [856] and rat hearts [857-862], in frog [863] and cat muscles [864] and in rat brain [865]. These led to in vivo measurements and the description of altered metabolism in diseased tissues [821,847], i.e. the MRS/MRI field and the limits of our scope. Shulman and coworkers combined $^{31}$P- and $^{13}$C-detected NMR to describe ATP and glucose metabolism in an integrative fashion in E. coli and rat liver cells, and used NMR saturation transfer to evaluate phosphate transfer to ATP, i.e. ATP synthesis by ATPase in their conditions [801].

Lastly, regular $^{31}$P-NMR monitoring of intracellular ATP has shown its usefulness to check the health status of cells in the NMR tube, which in turn helped to establish the benefits of intra-spectrometer bioreactors (see Section 3.7.). As discussed above (Section 2.3.2.), in combination with $^{13}$C-NMR, $^{31}$P-NMR and bioreactors were used to study the effects of various drugs on mammalian cancer cell metabolism [768,777-794], of ethanol and various chemicals on hepatocytes [795,796], and the metabolism of hybridoma [771,797], and yeast immobilized in gel matrices [866]. More recently, $^{31}$P-NMR also provided interesting checks for cellular health during in-cell structural biology studies [867-869].

### 2.3.3. Nitrogen flux studied by $^{15}$N-NMR and $^{14}$N-NMR

$^{15}$N-NMR detection in living cells was explored from the mid-1970's, providing a whole cell pattern for a parasite fungus [870]. In a more extensive study, Lapidot and Irving reported $^{15}$N-detected spectra of uniformly $^{15}$N-labeled E. coli, S. cerevisiae and Friend leukemic cells, as well as S. aureus labeled with $^{15}$N-glycine [580]. They took advantage of the large signal increase generated by $^{1}$H-decoupling via the heteronuclear $^{15}$N($^{1}$H) NOE effect: fast tumbling small molecules show an enhancement ratio of up to -3.93, while slowly tumbling macromolecules show an enhancement ratio of approx. +1 (i.e. close to no effect) [871], and semimobile molecules can produce a null signal in these conditions. Lapidot and Irving demonstrated the high flexibility and $^{15}$N-

NMR detectability of phosphatidylethanolamine, lysine and arginine nitrogens as well as many other unidentified nitrogens in cells, and of glycines in S. aureus peptidoglycan. Because they are pH- and viscosity-dependent, related $^{15}$N relaxation times and peak linewidth of amino acid signals were also used to probe their local environments, either cytoplasmic or vacuolar [467,872].

Hence, $^{15}$N-NMR under $^{1}$H-decoupling and NOE enhancement provided clean spectra of high abundance $^{15}$N-containing metabolites produced from $^{15}$N-labeled precursors such as $^{15}$NH$_4^+$ or $^{15}$NO$_3^-$.
The rationale used earlier for $^{13}$C-NMR of carbon fluxes was applied similarly to nitrogen fluxes: living organisms fed with natural abundance nutrients (99.6% $^{14}$N, 0.4% $^{15}$N) are suddenly exposed to $^{15}$N-labeled precursors; $^{15}$N-NMR spectroscopy then reports the integration of $^{15}$N atoms in the first metabolic products, the natural abundance background signal being close to zero. Using this $^{15}$N-NMR approach, nitrogen metabolism was characterized in live microorganisms such as the fungus Neurospora crassa [873,874], C. glutamicum, a bacterium used for industrial production of amino acids [875,876], Pseudomonas [758], or cyanobacteria [877], and even once in mammalian cells [878]. From the late 1980's, most $^{15}$N-NMR studies on living organisms were concerned with nitrogen assimilation by plants, focusing on symbiotic bacteria [879] or fungi [880-882], algae [883], spruce buds [884] or cell cultures [885], maize root [886-888], duckweed [889], invasive weed [890,891] and even carrot cell suspensions [892,893]. $^{15}$N-detected solid-state NMR spectra were also carried out on lyophilized soybean cotyledon or leaves to determine nitrogen fluxes from ammonium, nitrate, N$_2$, asparagine, glutamine, allantoin or methionine [894-899]. Overall, these studies permitted characterization of the relative contributions of two complementary pathways (glutamine synthetase/glutamate synthase GS/GOGAT, and glutamate dehydrogenase GDH) in nitrogen assimilation, or comparison of NH$_4^+$ and NO$_3^-$ incorporation rates in various conditions [900,901]. $^{15}$N-NMR of cell extracts has found convenient applications to evaluate the impact of various stresses, since it does not require any separation steps [901-903], although it still demands a careful exploitation of the spectroscopic data [904]. $^{15}$N-NMR also permitted a few studies of secondary nitrogen metabolism of alkaloid precursors, notably using $^{15}$N-tropinone [890,891,900,901,905,906]. It has also provided quantification of the metabolism of the neurotransmitter glutamate in MRS studies of animal brains, which lies outside the scope of this review [907].

Similar limits to those encountered in studies of $^{13}$C-NMR carbon fluxes were also met in $^{15}$N-NMR metabolite studies: the low sensitivity of NMR compared to mass-spectrometry made it less attractive from the early 1990's [908]. $^{15}$N-direct detection is practically feasible only for species found at millimolar concentrations. Moreover, the use of the $^{15}$N($^{1}$H) NOE to enhance sensitivity coupled to the long T1 relaxation (due to the low $^{15}$N gyromagnetic ratio) make it difficult to analyze $^{15}$N-NMR spectra quantitatively [904]. It has however some advantages. $^{15}$N-detection is feasible at every pH and temperature, in contrast to indirect $^{15}$N-filtered $^{1}$H-detection, which became popular from the 1980's [908-910]: although it can yield a 10-100 fold better sensitivity theoretically, $^{1}$H-indirect detection is impossible for many protons attached to $^{15}$N that exchange with water protons at rates above 10-100 Hz under physiological conditions. Long-range, multiple bond magnetic transfers have been designed that partially circumvent this problem, but at the expense of sensitivity [911]. $^{15}$N-NMR is also convenient for the straightforward, direct observation of NH$_4^+$ and NO$_3^-$ *in situ* without extraction. This may seem an unsophisticated NMR application, but it is useful because the methods used for extraction prior to for mass-spectrometry analysis can lead to information losses [908,911].

However, for nitrogen NMR studies focused on quantifying NH$_4^+$ and NO$_3^-$, $^{14}$N-NMR can be also considered. The quadrupolar nature
of ¹⁴N is an obstacle to NMR measurement in most cases, but has less effect on ¹⁴N resonances of simple, symmetric, fast tumbling metabolites like NH₄⁺ and NO₃⁻ even inside cells [912]. Close to symmetric nitrogen sites such as those of urea, trimethylamines or betaine may also be observed using ¹⁴N-NMR [913]. It does not require any isotope enrichment, because ¹⁴N represent 99.6% of nitrogen nuclei at natural abundance, as mentioned earlier. ¹⁴N-NMR permits more reliable quantification than ¹⁵N-NMR, notably because of the faster ¹⁴N T1 relaxation [914,915]. It has been applied to various systems such as mammalian tissues [912,916,917], plant roots and leaves [914,918,919], algae [920], *E. coli* [913], and spruce seedlings [884,885,921,922]. Shift reagents have also been used to distinguish extra-cellular from intra-cellular ¹⁴N-signals [923,924]. These were used, in combination with saturation transfer methods similar to those presented above for ³¹P-NMR, for measurement of transmembrane fluxes of ammonia in erythrocytes [925]. Moreover, ¹⁴N-NMR signal linewidth and peak multiplicity of NH₄⁺ is sensitive to pH because of the varying exchange rate of protons with water: the local pH where the detected pools of NH₄⁺ are found can be measured [918,921]. The related measurements are less direct than ¹³C, ¹⁵N or ³¹P chemical shifts characterization of chosen metabolites whose protonation states are pH-dependent [926]. Nevertheless, ¹⁴N-NMR permitted quantification of NH₄⁺ present in the cytoplasm, or its storage in very acidic vacuoles of maize roots [918] or spruce seedlings [921,922].

### 2.3.4. Overcoming spectral congestion in ¹H-NMR spectroscopy for higher sensitivity: monitoring cellular species below millimolar concentrations in the timescale of minutes.

Macromolecules and molecules embedded in macroassemblies, such as lipids in cell membranes, contain innumerable NMR-visible protons. Proton resonance frequencies have a narrower dispersion than ¹³C or ¹⁵N nuclei, and species such as water, lipids or macromolecules generate important and broad ¹H-NMR signals that obscure ¹H resonances from metabolites. As far as NMR detection is concerned, the benefits of low natural abundance of ¹³C cellular contents, or of the sparsity of observable ³¹P signals do not apply for ¹H species. Hence, although being the highest gyromagnetic ratio nucleus and yielding theoretically the highest NMR signal intensities, ¹H-detected NMR was not the first-choice method in early studies. ¹H NMR necessitated development of signal-filtering strategies before it could become established for the observation of metabolites in living cells and organisms. Various solutions have been found for various applications that we detail below.

A prior requirement for studies on metabolites in live cells is always a robust suppression of the very large NMR signal from water protons. The studies mentioned below used contemporary methods that were developed in parallel. Dedicated reviews exist on this topic [927], which is much less problematic now than it once was (see also [928-930] for live material).

NMR signals from macromolecules were first removed using the so-called spin-echo technique to purge out resonances from slow-tumbling species (see Section 3.3.2. for some brief explanations). In the mid-1970's, Daniels and coworkers used a spin-echo pulse sequence to separate the sharp resonances of adrenaline from the cellular background of adrenal gland slices [931]. The resulting ¹H-detected one-dimensional NMR signal was the first experimental observation able to support the hypothesis of a homogeneous, fluid pool of adrenaline in this native context. The metabolite was selectively observed also because of its high concentration. The sharpest resonances from metabolite species present at millimolar concentrations were then used to study specific aspects of cellular metabolism. Using one-dimensional spin-echo ¹H-NMR, scientists in Oxford started to quantify erythrocyte metabolic

activities such as glutathione redox equilibrium, or glucose consumption [932]. They used the same technique to monitor D-Alanine or L-Alanine influx after adding a paramagnetic agent in the external medium to suppress NMR signals from outside erythrocytes [933]. A growing number of workers applied one-dimensional spin-echo ¹H-NMR, either to quantify Zn²⁺ complexation by glutathione in erythrocytes [934], or variations in levels of phosphorylcholines through the cell cycle in Friend Leukemia cells [935], choline levels in erythrocytes [936], the metabolism of malate, fumarate and lactate as a function of NAD+, pyruvate or oxalate concentrations in erythrocytes [937], amino acids levels and glucose consumption in HeLa cells [938], methylglyoxal metabolism and the stability of ergothioneine in erythrocytes [939,940].

An important aspect of spin-echo experiments is the evolution of scalar couplings between protons during the echo time. Within specific ranges of spin-echo durations, scalar couplings cause the inversion of certain proton signals (i.e. the signals become negative) (Fig. 15). Interestingly, if a proton-deuterium exchange occurs for one of the protons involved in the spin-spin coupling, the coupling is effectively removed (¹H-²H couplings are approx. 6.5 times smaller than the corresponding ¹H-¹H coupling, and the ²H nuclei are unaffected by ¹H pulses) and the spin-echo does not invert the resonance signs of the neighboring protons anymore. This concept permitted monitoring of the incorporation of deuterated glycine into glutathione [941], of specific enzyme activities that result in H/D exchange between water and pyruvate and lactate [942-944], and of lactate/pyruvate exchange upon supplementing cells with deuterated lactate or pyruvate [945-947], again in erythrocytes.

Unlike ¹²C nuclei (that have no spin), ¹³C spins are also scalar-coupled to covalently bonded protons. Integration of ¹³C nuclei into metabolites can thus be detected in ¹H-detected spectra in multiple ways: i) they cause a peak splitting in ¹H-detected spectra, the two new peaks being separated by the scalar coupling ¹J (¹H-¹³C)~100-150 Hz; ii) the signs of these peaks can be also manipulated specifically by ¹³C 180° pulses during the spin-echo (see Section 3.3.2.). Shulman and coworkers were the first to use this phenomenon to quantify ¹³C-incorporation from [2-¹³C]-acetate into glutamate or aspartate by S. cerevisiae [948]. A more advanced ¹³C-modulated ¹H spin-echo was exploited by Brindle and colleagues to monitor the ¹²C- and ¹³C-methyl content of alanine and pyruvate and thus quantify the activity of the alanine aminotransferase in erythrocytes [949]. Foxall and coworkers used and published a similar application of ¹H-¹³C spin echo difference to report the conversion of [1-¹³C]-glucose into [2-¹³C]-ethanol in yeast [950]. They showed that such indirect ¹H-detection of the presence of ¹³C provided a 22-fold higher sensitivity than direct ¹³C-detection. This strategy was also employed to detect ¹⁵N-incorporation from ammonium or glutamine in cultured mammalian cells, with similar sensitivity benefits [876]. Use of such spin-echo related filters was however abandoned by the majority of the community, and substituted by magnetization transfers allowing ¹H-detected, ¹³C- or ¹⁵N-edited 2D NMR spectroscopy. These spread the signals in a supplementary dimension and are thus likely largely to avoid peak overlaps (see below in Section 2.3.6.)

The combined use of multiple nuclei including ¹H-detection was explored from the early 1980's [946,951,952]. ¹H-detected NMR was soon considered to be complementary to ¹³C and ³¹P-NMR in the characterization of living tissue metabolism [761,953,954]. For example, ¹H- and ³¹P-NMR were combined to measure total creatine and creatine phosphate pools in frog muscles [951], and to report simultaneously pH and lactate levels in guinea pig brain slices using multiple channel NMR-probes [955,956]. In recent years, this complementarity between ¹H- and ³¹P-NMR was also applied to the simultaneous monitoring of
<figure><img src="image_19.png" /><figcaption>**Fig. 15.** A spin-echo can filter out signals from slowly tumbling molecules. We show here the theoretical schemes for the time-domain evolution of ¹H-signals acquired after a single 90° pulse for A) fast tumbling or B) slow tumbling molecules. The effective transverse relaxation T2* can be dominated by sample inhomogeneities and chemical shift distribution. C) A spin-echo refocuses the magnetization coherence lost due to inhomogeneities, but not that lost because of the intrinsic molecular T2. Hence, spin-echo ¹H-spectra show signal intensities modulated by the molecule-specific T2x2τ, which is much attenuated for D) slowly-tumbling molecules. E) One-dimensional ¹H-detected spectrum of a bovine adrenaline medulla slice; F) one-dimensional spectrum acquired after (2x 60 ms) a spin-echo of the same slice, revealing signals from adrenaline molecules (found at high concentration in this tissue). Signals from species having short transverse relaxation times T2 vanished. Because scalar couplings can evolve during the spin-echo, signals from CH₂ groups or from protons with a large, vicinal scalar coupling H₅/H₆ are inverted (highlighted in green). These spectra were recorded in 1976 at 270 MHz. Adapted from Daniels et al. [931].</figcaption></figure>

cellular metabolite levels, RNA synthesis or folding and protein interactions [466]. A number of cellular structural biology studies focusing on proteins took the simpler approach of using ³¹P-NMR to check cellular metabolic states from ATP levels, together with ¹H-detected NMR of proteins [867,868].

¹H-NMR acquisition can also be performed without background filters when monitoring metabolites supplemented at very high-concentrations, e.g. the metabolism of diluted *E. coli* cell suspension in ~100 mM glucose [957,958]. At such high concentrations, ¹³C-coupled ¹H-doublet signals are also easily observed, which allowed monitoring of ¹³C-glucose metabolism and the incorporation of carbon by the rumen cellulolytic bacterium *Fibrobacter suc cinogenes* [959]. We mentioned previously that lipids can often be seen as a problem, because their signals completely obscure parts of ¹H NMR spectra. These large signals can become interesting for monitoring the global lipid content of cells, and have been exploited to quantify live microalgae cultures in real time [960,961,962,963].

The observation of metabolite consumption is of course possible in the spent medium following culture of any cells, like plant, tumor spheroids [964] or mammalian cells grown to produce therapeutic proteins [965,966]. Extracellular metabolites generally provide sharp NMR signals that are good reporters of intracellular metabolism. A recent study of primary cancer cells permitted examination of their adaptation to hypoxic conditions: about 35 metabolites were identified and about a dozen monitored over 24 hours using samples containing 1.10⁷ to 5.10⁷ cells/mL embedded in agarose [967]. Similarly, a dozen metabolites were monitored in lactic bacteria growing over 24 hours in various glucose and pH conditions in the NMR tube [968]. Using this approach and up-to-date commercial NMR probes, one-dimensional proton spectra can be recorded every 5-10 minutes and provide metabolite quantification with ~10 µM precision for 500 µL sample volumes. Similar results were obtained from primary cancer cells embedded in methylcellulose, which allowed monitoring of extracellular levels of about 10 metabolites over 24 hours [969]. Although presenting practical advantages when compared to classical methods using quenching and extraction, this live-cell metabolomics approach using ¹H-NMR requires a careful deconvolution of signals because of progressive peak shifting and broadening [967,968]. The corresponding signals can thus be exploited with confidence only if their intensities surpass those of the other

signals with similar chemical shifts, which often means if the quantified species are found in the millimolar range. However, such applications fall in the area of metabolomics studies and are not considered here in greater detail.

### 2.3.5. HR-MAS ¹H-NMR spectroscopy: lipid filtering/editing and metabolome of intact cells, tissues and small living organisms.

Although spin-echo and T2-filters improved the ¹H NMR detectability of a number of metabolites, the observed peak linewidths were still too broad to allow analysis of the intracellular metabolome with much accuracy. Magnetic inhomogeneities or restricted motion of mobile species cause line-broadening of NMR signals, preventing deconvolution of signals. Moreover, resonances from lipids are often not completely removed by T2 filters: some intracellular pools of lipids exhibit largely liquid-like behavior and, even though constrained, maintain high mobilities. In general, the metabolites that could be analyzed with this approach were those found at high concentration, that were highly mobile, and producing NMR resonances far from those of lipids. Non-membrane lipids themselves, with their semi-restricted motional regime, were thus “in limbo” as far as accurate in-cell NMR description was concerned.

In an earlier Section (2.2.3.) on cell-surface lipopolysaccharides and glucans, we discussed briefly the deleterious effects of molecular restricted motion and non-homogeneous magnetic susceptibility, especially for ¹H NMR observation (see also Sections 3.2.2. for more detailed explanations). These broaden the NMR signals observed using classical liquid-state NMR approaches. However, high degrees of local mobility are often maintained for small molecules like metabolites in non-frozen samples, which can yield sharper peaks if Magic-Angle Spinning (MAS) is applied. Hence, high peak intensities and narrow linewidths (Fig. 16) can be obtained from mobile species, including lipids, contained in cell pellets, excised animal tissues or even small animals, e.g. *C. elegans*, small crustaceans or flies. These have to be loaded in a cylindrical sample container (the “rotor”) and spun between 1 and 4 kHz at an angle of 54.7°, with respect to the spectrometer magnetic field B₀.

This approach has been coined as High-Resolution-MAS (HR-MAS) NMR spectroscopy and has the obvious advantage of permitting the analysis of intact, non-processed samples (see Section 3.4.3. for a specific discussion on HR-MAS). It has been applied for more than 20 years in chemistry and food science,
<figure><img src="image_20.png" /><figcaption>**Fig. 16.** One-dimensional ¹H NMR spectra of cultured adipocytes under **A)** non-spinning conditions, and **B)** Magic-Angle Spinning (MAS) at 3.5 kHz. Both spectra were recorded at 11.7 T (500 MHz) in 42 minutes and twice as many cells were used in the non-spinning conditions. Adapted from Weybright et al. 1998 [987].</figcaption></figure>

but also to characterize the metabolome of intact cells or tissues in conditions such as those we describe below. Recent reviews can be found on this topic [970,971]. This approach is attractive because of its relative convenience: it requires ~5 million cells per measurement, i.e. 15-50 uL or mg of wet pellets, and no time-consuming chemical extraction. It benefits from the sensitivity of ¹H NMR and thus permits accurate quantification of metabolites present at ~0.1 mM concentrations [972]. To put this in context, more than 60 metabolites are present at 0.1 mM or above in growing *E. coli* [973]. Investigations on intact cells deliver only relative quantifications of the identified metabolites: they rely on semi-quantitative, comparative measurements of signal intensities in the various conditions tested. Absolute quantification can still be achieved using synthetic signals as an electronic reference [974,975]. The corresponding analysis falls in the category of largely classical metabolomics studies of quenched -although often "intact"- samples, which we do not cover extensively in this review. However, a number of recent developments permitted real-time observation of metabolic fluxes in small animals, for which we made an exception.

The benefits of Magic-Angle Spinning had been shown for drastically narrowing ¹H and ¹³C NMR signals of plant seeds at the end of the 1980's [630,976], and fruit tissues at the beginning of the 1990's [977-979]. The idea was taken up in the 1990's to compare the metabolome of normal and metastatic lymph nodes [980], and for the analysis of other animal soft tissues, e.g. kidneys [981,982], brain [983], adipocyte [984,985] and breast tissues [986]. Correspondingly, at the end of the 1990's, Singer and colleagues had shown that MAS at ~3-4 kHz permitted narrowing down ¹H NMR signals linewidth of intracellular lipids and metabolites by a factor ~10 in cultured adipocytes (Fig. 16) [987].

This helped to quantify cellular contents in various lipids from undifferentiated and differentiated adipocytes and to extend this type of characterization to benign and malignant liposarcoma tissues [988,989], and to measure intracellular diffusion coefficients of lipids [987,990]. Similar experiments were carried out more recently on cultured cells (exposed to cytotoxic agents) to derive

the diffusion coefficients of intracellular lipids on the millisecond timescale using HR-MAS, in a complementary fashion to fluorescence microscopy measurements on a time scale of seconds [991]. Because intracellular lipids conserve some mobility, ¹H HR-MAS NMR gave access to lipid analysis in intact cultured cells for various purposes: to distinguish neuronal and glial cells [992], to monitor phospholipid content in melanoma cells upon exposure to chemicals [993], fatty acid content variations in insulin secreting cells [994] and in mesenchymal stem cells [995]. These quantifications should however not be taken as absolute: increasing MAS spinning rates cause different degrees of ¹H NMR signal enhancement in different lipid pools, probably because of variable release of mobility restriction, as shown on glial cells [996]. Similar differences were observed for ¹H HR-MAS lipid signals of biopsies from different brain tissues [997].

¹H and ¹³C HR-MAS NMR also proved to be valuable for analyzing the whole-cell metabolic profile of algae in one single experiment [998-1000], which would usually have been carried out from separate water-soluble fractions and lipophilic extracts. Such a convenient profiling method of intact cells by HR-MAS NMR was presented as well suited to the analysis of algal content for nutritional and bio-fuel applications [960,1001-1003] and the identification of marine microorganisms [1004,1005]. Recent approaches favor flow-cell ¹H NMR in line with bioreactors to monitor the global lipid content of live microalgae cultures in real-time [960,961], also using benchtop NMR spectrometers [962,963]. These last characterizations do not require MAS and ensure a true non-invasive monitoring, although they provide less detailed information on the nature of the observed lipids.

Improved lipid editing/filtering capacities have been shown by diffusion editing methods: in combination with more classical T2-relaxation filters, they permit isolation of signals from species that present either unrestricted or restricted diffusion, semi-solid behavior and even those from true solid material (like chitin or lipid-protein matrices) if ¹³C-labeling and -detection are exploited [1006,1007]. This information can be useful to delineate the different pools of molecules, e.g. those that are free, and thus ready to be
incorporated in biosynthesis reactions, as opposed to those that are less mobile, and which are used for energy storage.

In the past decade, ¹H HR-MAS NMR has also been used to analyze the metabolome evolution of intact mammalian cultured cells previously exposed to chemicals: human cancer cells responding to chemotherapeutic agents [1008-1014], cultured cells reacting to pathway activators [990,1015,1016] or exposed to nanoparticles [1017], or expressing proteins with oncogenic mutations [1018], and reconstructed human epidermis exposed to skin allergens [1019]. In this last case, we can highlight the use of derivatives of ¹³C-labeled methanesulfonate, an allergenic reactive agent metabolized into new metabolites, which were quantified using ¹³C-filtered, ¹H-detected HR-MAS NMR [1020,1021]. Recently, HR-MAS has been applied to monitor the metabolic changes occurring within aggregating brain cell cultures, which may be promising for future neurotoxicology investigations according to the original authors [1022].

¹H HR-MAS was also used for studying microorganisms, e.g. to monitor the internalization and intracellular processing of a drug in mycobacteria [1023,1024], to compare planktonic and biofilm forms of *Pseudomonas aeruginosa* [1025,1026], and to characterize the metabolic consequences of reductase overexpression in the protozoan parasite *Giardia lamblia* [1027]. The metabolic activity of *G. lamblia* at confluency had been previously observed in real-time using HR-MAS at 3 kHz, which did not affect the stability of the trophozoites according to the authors [1028]. In a recent report, mycelia of *Neurospora crassa* were submitted to 6 kHz HR-MAS without apparent sedimentation and cell death, according to the authors, which permitted continuous monitoring of central metabolism pathways in aerobic and anaerobic conditions over 11 hours [474].

We stressed cell survival in the previous paragraph because it is a clear issue in HR-MAS experiments. Classical HR-MAS experiments require spinning rates in the range of 1-5 kHz to avoid the appearance of artefactual signals called "spinning sidebands". Using a standard disposable 4 mm diameter rotor, these spinning rates generate centrifugal acceleration between 8,000 and 100,000 g (gravity of Earth), which affects the viability of most mammalian cells and tissues [67] (see Section 3.3.4.). Certain cells are less sensitive to MAS at kHz rates, like erythrocytes [1029] or the above mentioned microorganisms. Improved pulse sequences and sample preparation protocols have been designed to make HR-MAS NMR feasible at slower MAS rates. Hence, a single lumpfish egg spun at only 400 Hz remained intact and provided NMR spectra showing promising resolution and signal-to-noise ratio for metabolic studies (Fig. 17) [1030]. Advanced pulse sequences permitted recording of highly resolved HR-MAS spectra of mouse and rat tissues using spinning rates down to 40 Hz and even 1 Hz in the early 2000's [1031-1033], but these approaches have so far not proved sufficiently practical for common use [929].

Slow MAS is essential for studies on live pluricellular organisms. These are promising approaches for understanding and evaluating the toxicity of environmental stressors on model animals like water fleas (*Daphnia magna*), fresh water shrimp (*Hyallela azteca*) and *C. elegans*. Continuous monitoring of single individuals or populations is an efficient approach to avoid the blurring impact of metabolic variability before starting the observation: real-time trajectories can greatly improve the interpretation and reduce the number of repeat experiments needed [1034]. Early studies in the 2010's had shown that anesthetized *D. magna* could be maintained alive at 2 kHz for days, giving access to the lipidic profile of juvenile and adult animals carrying eggs or not [1035]. Similarly, low-temperature anesthetized flies *D. melanogaster* were placed in HR-MAS rotors and survived 2 kHz MAS, the aim again being to compare the lipid content of individuals [1036]. Similar conditions were used to obtain a spatial map of high-concentration metabo-

lites of the head, thorax and abdomen of living, anesthetized flies [1037]. However, anesthetized animals are poor models for toxicity assays and spinning generates significant stress. The non-anesthetized fresh-water shrimp *H. azteca* survive about 40 and 6 hours at 0.5 and 2.5 kHz MAS rates, respectively [929,1038]. Above 2.5 kHz, their survival is affected in one hour [1006]. *D. magna* is even more sensitive to MAS and withstands only 24 hours at 50 Hz MAS [1038]. Simpson and coworkers have developed advanced NMR pulse sequences and sample preparation that made it possible to obtain quality spectra of *H. azteca* at these low MAS rates [929,1006,1038-1040]. These notably include labeling of animals by feeding with ¹³C, thus allowing heteronuclear correlation NMR, which we will describe in the next subsection. The development of slow (300 Hz) micro-HR-MAS systems by Wong and colleagues has even permitted acquisition of promising spectra from one single *C. elegans* worm [1041]. Using the same experimental design, they managed to analyze samples containing only 19 million yeast cells (~250 nL) and to distinguish their metabolome at various stages of growth or under osmotic stress [1042]. These recent low-spinning HR-MAS approaches may thus have interesting applications in the near future to study small organoids or animals.

### 2.3.6. 2D ¹H/¹³C-correlation NMR of live cells and small animals: metabolites at submillimolar concentrations on time-scales of minutes to hours

In the previous subsections, we presented the capabilities of one-dimensional NMR spectroscopy for monitoring the metabolite content of cells. The approach is always limited by peak resolution: in most cases, overlapping signals means loss of information, even though peak deconvolution can sometimes extract peak intensities from known resonances. As discussed above, although it is the most sensitive approach on paper, ¹H-detected NMR provides poorly dispersed and broad peaks in cells. We have seen that HR-MAS can narrow intracellular ¹H peaks, but this strategy has its own limits currently, particularly concerning its effects on living material.

Two-dimensional ¹H-detected ¹H/¹³C-correlation NMR offers additional spectral dispersion, which is clearly useful to avoid peak overlaps while preserving ¹H-detection sensitivity. In the late 1970's, Bodenhausen, Freeman, Ernst and their coworkers proposed pulse sequences correlating ¹H to covalently bonded lower gamma nuclei like ¹³C or ¹⁵N [1043-1047]. This not only gave access to ¹³C or ¹⁵N chemical shift measurements with the sensitivity of ¹H NMR, but being a two-dimensional technique, it could "disentangl[e] spectra that are complicated by extensive and multiple overlapping" as described by the aforementioned authors [1048]. This principle has been used for classical metabolomics studies of extracts and biofluids, originally under the name 2D ¹H-¹³C correlation spectroscopy [1049]. It is nowadays more often referred to as 2D ¹H-¹³C HSQC or HMQC spectroscopy and their derivatives (see for example [1050,1051]). Like one-dimensional NMR, ¹H-¹³C correlation spectroscopy is quantitative and several hundreds of metabolites have their signals assigned in 2D NMR spectra databases [1007]. It is however a bit more complicated to extract absolute quantities from 2D spectra than from 1D spectra (for a comprehensive discussion, see [1052,1053]).

In the context of classical metabolomics using extracts or biofluids, 2D ¹H-¹³C correlation spectra can be recorded using samples at natural isotopic: the 1.1% of ¹³C isotope can be sufficient to obtain a reasonable S/N ratio, provided that the sample is stable enough to remain intact during the recording of a large number of scans. When it comes to intact cells, ¹³C-enriched metabolites are supplemented in the culture media and their processing is monitored similarly to what was achieved using one-dimensional ¹³C NMR spectroscopy (see subsection 2.3.1): the assigned peaks
<figure><img src="image_21.png" /><figcaption>**Fig. 17.** One-dimensional ¹H NMR HR-MAS spectra of a single lumpfish egg (1 mm diameter) recorded at **A**) 4 kHz, or **B**) at 400 Hz, both using the PROJECT T2-filter suppressing the broad signals from macromolecular assemblies. The photographs show the lumpfish egg after MAS at 4 kHz (left) or 400 Hz (right). Both spectra were recorded in 4.5 minutes at 9.4 T (400 MHz). The higher S/N at 4 kHz is due notably to the higher fluidity of the leaked cytosol. Adapted from André et al. 2014 [1030].</figcaption></figure>

are quantified in time-series of spectra, which contains information about the fluxes of the supplemented ¹³C nuclei. The isotopic filter works as well as for one-dimensional ¹³C-spectroscopy, but 2D ¹H-¹³C NMR has been reported only since the 2000's, first in combination with HR-MAS [641,974,1015,1020], and then using "classical" solution NMR for metabolic monitoring of live cells or organisms, to the best of our knowledge.

Thus, Park and colleagues have monitored and compared the metabolism of normal and cancer cells in real-time by tracing the levels of alanine, glycine, serine, pyruvate, lactate, acetate, succinate and α-ketoglutarate upon ¹³C-glucose supplementation [1054]. They were also able to describe the dynamic changes of metabolism after subjecting cancer cells to galloflavin, an inhibitor of lactate dehydrogenase. In a later study, they showed how to measure metabolism in live mitochondria upon ¹³C-pyruvate supplementation, by monitoring 2D ¹H-¹³C signals from acetate, citrate, succinate, glutamate, acetate, acetyl phosphate and acetyl-CoA [1055]. These measurements were executed on wild-type and p53-KO cells, revealing their fundamental metabolic differences. With intact cells or isolated mitochondria, they recorded 2D ¹H-¹³C spectra every 5 minutes, which generated spectra showing an accuracy for concentration estimated (by the present authors) to be in the range in the range of ~20 µM.

In parallel, Simpson and coworkers studied two algal strains and one cyanobacterium cultivated in presence of ¹³CO₂, and established the ability of 2D ¹H-¹³C to identify and quantify 40-50 metabolites in freeze-dried whole cells [1056]. These were actually grown with the purpose of feeding water fleas *D. magna*: once placed in a flow-cell supplying oxygenated water, the living animals provided reasonable quality 2D ¹H-¹³C correlation spectra revealing differences between those fed with algae or cyanobacteria [1057], with natural algal bloom or laboratory control algae [1058], between pregnant and non-pregnant females [1059], or under hypoxic stress [1034]. In a similar fashion, using 2D ¹H-¹³C NMR, Park and colleagues have recently monitored the time-dependent changes of 24 metabolites in living *C. elegans* fed with ¹³C-labeled *E. coli*: two *C. elegans* strains engineered to knock out two different isotypes of a catalytic subunit of the AMP-activated kinase showed different metabolic reactions to starvation [1060]. We can also mention that Kazimierczuk and his coworkers monitored the metabolism of live *E. coli* exposed, or not, to ampicillin using 2D ¹H-¹³C correlation NMR [1061]. According to Simpson et al., the use of electronic referencing appears to make absolute quantification possible [1051]. Moreover, they have evaluated that a species at 100 µM would yield a signal-to-noise ratio of 10:1 in 2D ¹H-¹³C spectra recorded in about 5 minutes (at 500 MHz using

a cryoprobe) [930,1051]. 2D ¹H-¹³C correlation spectra are commonly recorded using ¹H-detection because of its higher sensitivity, but ¹³C-detection avoids problems from any unsuppressed water signal and thus can be complementary in peak detection and assignment: over 60 metabolites have been identified in living *D. magna* using both schemes [1062,1063].

Supplementary filters can be exploited. By selecting only ¹H-¹⁵N-¹³CO protons, it is possible to observe very specifically ¹³C/¹⁵N-labeled glutamine supplemented in a natural abundance culture medium and thus to monitor its hydrolysis in live mammalian cells [1064]. Simpson and colleagues have established amino-acid specific 2D ¹H-¹³C pulse sequences to monitor the free amino-acid concentrations [1065]. Hence, they measured their increase in *D. magna* upon exposure to a concentration ramp of bisphenol-A with a 1 minute time-frame allowing a very precise determination of the lowest concentration eliciting a response [1066]. They also introduced ¹²C-editing methods to monitor specifically the incorporation of ¹²C-molecules in animals grown under a ¹³C-labeled diet. They showed in a case study that they could use HR-MAS at 2.5 kHz to follow the conversion of nicotine into cotinine by *H. Azteca*, and the incorporation of ¹²C nutrients by *D. magna* in their ¹³C initial metabolome using an advanced strategy selecting signals only from ¹H-¹²C-¹³C groups [1067]. Finally, they also tested a strategy using ²H-¹³C labeling of algae, *H. Azteca* and *D. magna* to record 2D ²H-¹³C correlation spectra: while ²H-detection is intrinsically much less sensitive than ¹H-detection, Simpson and colleagues have shown that it can generate useful 2D spectra under 50 Hz HR-MAS of about 20 metabolites when very mobile, without any background issues from signals of water, lipids or macromolecules [1038].

All these approaches are clearly applicable not only for NMR studies of live animals, but also those on live cells and organoids. Although 2D ¹H-¹³C correlation NMR requires ¹³C-labeling, which can be expensive in some cases, a number of organisms can now commonly be obtained fully or partially ²H-, ¹³C-, and or ¹⁵N-labeled, notably for recombinant expression purposes. A number of culture media depleted in specific amino acids are commercially available for many cell types, and their supplementation with specifically labeled amino-acids can be relatively affordable. These may be applied for NMR in the near future for studies on metabolic and toxicologic responses to inhibitors or pollutants. In a complementary manner to classical quench- and end-point studies, the ability to monitor single samples in a continuous, non-destructive fashion over periods of days is of great interest for understanding biochemical pathways, toxic-mode-of-action or to firmly establish the lowest toxic concentrations.
Low-speed spinning methods have been developed for these applications linked to metabolomics [636,929,972,1030,1068,1069], but detailed discussion is beyond the scope of this review.

### 2.3.7. <sup>13</sup>C-hyperpolarization for carbon-flux quantification at low-millimolar concentrations and time scales of seconds

As mentioned earlier, the “NMR-visible” <sup>13</sup>C isotope has a low natural abundance. Hence, tracing <sup>13</sup>C-labeled species by <sup>13</sup>C-detected NMR can be a convenient approach to monitor metabolite incorporation and processing in cells that are themselves mostly made of “NMR-invisible” <sup>12</sup>C. However, the poor sensitivity of <sup>13</sup>C-NMR has been a major limitation. This is mainly due to the low extent of nuclear magnetic polarization at physiological temperatures, about 0.001% (that is to say the lower energy spin state has about 0.001% higher population than the higher) at common commercial magnetic field strengths, namely about 10 T. <sup>13</sup>C-hyperpolarization strategies have been proposed to enhance this population difference substantially.

Among such methods, dissolution-DNP (d-DNP) enhanced NMR has been the principal hyperpolarization approach used for achieving real-time monitoring of metabolic reactions in living cells. Pioneering d-DNP studies were reported in the early 2000's [1070,1071], making it possible to achieve 10,000-50,000 fold enhancements in signal, and this spurred a number of applications [1071,1072]. Concise explanations of this technique and others that are closely related are given in Section 3 (see also [1073,1074]), but the outline of a d-DNP experiment will also be described here: <sup>13</sup>C-enriched metabolites are hyperpolarized at cryogenic temperatures (~1K) in the presence of polarizing agents carrying free electrons by irradiating with microwaves at appropriate frequencies; importantly, this is usually done in a separate apparatus from the main NMR magnet. The <sup>13</sup>C nuclear magnetic polarization achieved can reach 10-50% within an hour. The metabolites are then submitted to fast dilution to quickly thaw the hyperpolarized samples and permit their entry into cells. The half-life of the hyperpolarization typically ranges from 1 to 50 seconds, depending on the metabolite and the conditions, which makes rapid transport to the magnet of the NMR spectrometer an absolute necessity. This also implies the technique is only useful for fast metabolic reactions that can be monitored in a few tens of seconds (Fig. 18).

The reasons that initially made <sup>13</sup>C-NMR appealing for MRI and MRS become still stronger if <sup>13</sup>C is hyperpolarized: i) <sup>13</sup>C-enriched, hyperpolarized molecules generate <sup>13</sup>C-NMR signals, which ensure their selective observation; ii) <sup>13</sup>C-enriched, hyperpolarized molecules that happen to be substrates of a chemical reaction transmit their <sup>13</sup>C-enrichment and hyperpolarization to the reaction products, which also become observable. The immediate proof-of-concept application of d-DNP by Ardenkjaer-Larsen and colleagues was a rat angiography study using hyperpolarized <sup>13</sup>C-urea [1076]. Following their previous work on <sup>13</sup>C-NMR/MRS, Brindle and colleagues, followed by a growing number of others, showed progressively that d-DNP hyperpolarized <sup>13</sup>C-metabolites (notably pyruvate [1077], but also lactate [1078], glucose [1079] or fumarate [1080], bicarbonate as a pH sensor [469] or ascorbic acid as a redox sensor [487]) were exquisite reporters of tumor-specific enzymatic activity for in vivo imaging applications [33,1081]. As shown by successful applications in humans, this approach may arguably compete with or be complementary to positron emission tomography (PET) for clinical purposes [1082-1086]. It also has potential for imaging heart or brain metabolism [1084,1087-1090]. Translation to clinical applications requires a number of difficulties to be overcome [1084,1091,1092], which we will not detail here.

With such promise and clinical potential, the d-DNP field and its application to investigations with cells multiplied greatly in the last 10 years. A number of hyperpolarized molecules have been

tested, often with use in imaging applications as a motivation. They gave access to real-time monitoring of cellular metabolism in a variety of situations. Among hyperpolarized <sup>13</sup>C-metabolites, pyruvate has probably been the most frequently used to characterize mammalian cells: it permits quantification of cancer-related glycolysis through pyruvate to lactate conversion by lactate dehydrogenase (LDH). This requires that cells be maintained in well-defined metabolic states, which led to approaches where cells are encapsulated in polymers, coupled with the use of a perfusion system: this permits cells to be kept in a constant flow of fresh medium while hyperpolarized substrates are injected at defined time points [1093,1094]. These first studies revealed that carboxylate pyruvate/lactate transmembrane transport by MCT1/MCT4 was as important as LDH activity for cancer cell metabolism. Advanced mathematical treatments integrating these parameters were recognized to be necessary for a proper quantification of pyruvate to lactate conversion [1095-1097], unless paramagnetic species like Gd-DO3A were added to quench the extracellular <sup>13</sup>C-pyruvate/lactate NMR signals [1098]. Hence, the effects of a PI3K inhibitor on the glycolysis pathway in murine lymphoma cells were attributed to a decrease in LDH expression [1095], while the exposure to a MEK inhibitor caused higher or lower expression of the pyruvate import protein MCT1 in prostate or breast cancer cells, respectively [1099]. Using varying flow rates of medium in the NMR tubes, Kurhanewicz and colleagues evaluated <sup>13</sup>C-hyperpolarized lactate efflux, and its importance was underlined by the high expression of lactate exporter MCT4 in metastatic renal carcinoma cells, as compared to non-metastatic ones [1094]. More recently, hyperpolarized <sup>13</sup>C-pyruvate influx due to MCT1 and lactate efflux due to MCT4 were shown to have a preeminent role in the oncogenic metabolism of pyruvate, as measured by d-DNP experiments [1100,1101]. Consistently, reduced expression of LDH, MCT1 and MCT4 caused lower pyruvate to lactate conversion rates in mutant IDH1 (isocitrate dehydrogenase 1) glioma cells [1102].

It is interesting to note that in a highly homogeneous field, two close but separated <sup>13</sup>C-NMR signals corresponding to extra- and intra-cellular lactate could be distinguished [1103,1104]. Alternative strategies were recently developed to distinguish intra- from extra-cellular hyperpolarized metabolites, which rely on the varying diffusion coefficients between the two environments [1105,1106]. These made it possible to detect pyruvate to lactate conversion together with their extra- and intracellular populations and diffusion rates in cultured cancer cells.

Hyperpolarized <sup>13</sup>C-pyruvate d-DNP experiments were combined with more classical metabolomics using <sup>1</sup>H-NMR HR-MAS and <sup>31</sup>P-NMR, which made possible a distinction between benign and malignant prostate cancer tissues [1107]. Another combination of d-DNP with more standard <sup>13</sup>C-NMR metabolomics was proposed, whereby cells were cultured in a medium containing [3-<sup>13</sup>C]-pyruvate and briefly exposed to hyperpolarized [<sup>1</sup>-<sup>13</sup>C]-pyruvate: this allowed quantification of the rapid conversion of [<sup>1</sup>-<sup>13</sup>C]-pyruvate into lactate and bicarbonate via the cancer-associated glycolytic metabolism and the slower integration of [<sup>3</sup>-<sup>13</sup>C]- or [<sup>1</sup>-<sup>13</sup>C]-pyruvate via the more standard Krebs cycle, also in the presence of Akt inhibitors [1108]. To apply hyperpolarized <sup>13</sup>C-pyruvate d-DNP experiments to mass-limited samples, Keshari and coworkers implemented a micro-coil apparatus, which made possible the real-time monitoring of pyruvate metabolism in ~10⁵ cells in a 1.05 T permanent magnet [1109] or in a 3 T MRI scanner [1110]. This allowed rapid assessment of the cancer-related glycolytic metabolism of various cancer cells upon drug exposure, which should now also be feasible for primary cells, organoids or biopsies.

Although monitoring glucose uptake and degradation would be more logical for quantification of glycolysis in cells, the fast T1 longitudinal relaxation of [<sup>U</sup>-<sup>1</sup>H,<sup>U</sup>-<sup>13</sup>C]-glucose (~1 second) precluded
<figure><img src="image_22.png" /><figcaption>**Fig. 18.** A) Operating principle of Dynamic Nuclear Polarization (DNP). Current technologies in common use employ intense microwaves to polarize nuclear spins in the presence of polarizing agents (particular molecules containing unpaired electrons, i.e. free radicals) at cryogenic temperatures; B) Operating principle of dissolution-DNP: hyperpolarization of nuclear spins is reached at cryogenic temperature under microwave (MW) irradiation; the hyperpolarized substrate is then dissolved in the final liquid sample for analysis (here suspension cells); C) Stacked one-dimensional ¹³C-NMR spectra showing real-time observation of glucose in *E. coli*, *S. cerevisiae* and cultured proliferative human cells: hyperpolarized ¹³C-labeled glucose is supplemented to the medium at time 0 and time series of 1D ¹³C-spectra are then recorded. These show the progressive integration of ¹³C-atoms into various metabolic products, together with a progressive fading of the signal due to T1 relaxation, ending with the loss of the hyperpolarization. (The ¹³C-glucose signals below 100 ppm are not shown here). Adapted from Lerche et al. 2015 [1075].</figcaption></figure>

such an approach: all hyperpolarization would be lost in just a few seconds. Several groups proposed use of uniformly deuterated ¹³C-glucose, which slowed this relaxation by a factor of about 10 [1079,1111-1114]. Hence, using d-DNP ¹³C-NMR, Frydman and coworkers could observe a number of intermediate products of the glycolytic pathways from supplemented hyperpolarized [U-²H,U-¹³C]glucose, such as dihydroxyacetone phosphate, 3-phosphoglycerate and lactate [1112]. They also revealed the presence of “invisible” intermediate products present at low concentrations: they applied saturation pulses at the NMR frequencies where these products resonate and then measured the resulting variations of signal from the downstream “visible” products that were more abundant. Such a saturation strategy is very similar to those used for ³¹P-NMR earlier (see above). Because of the multiple intermediate products generated in these carbohydrate pathways, it can be difficult to assign the emerging signals to single species [1112,1115]. Using an approach exploited earlier in standard ¹³C-NMR experiments [776], the cytosolic NAD+/NADH equilibrium has been measured from the ratio between intracellular ¹³C-pyruvate and ¹³C-lactate produced from hyperpolarized ¹³C-glucose [1113].

Dissolution-DNP experiments have also been used to study microorganism metabolism. Researchers affiliated to the Carlsberg Laboratory directed their investigations towards yeast glycolysis and CO₂/ethanol production: upon feeding *S. cerevisiae* cell suspensions with hyperpolarized (and deuterated) ¹³C-glucose or fructose, they observed the successive appearance of a dozen intermediate products from central carbon metabolism [1116], its inhibition by sulfite notably via pyruvate-sulfite adducts formation [1117], acetaldehyde accumulation upon acetate exposure [1118] and its impact on the intracellular redox status of the glycolysis pathway [1119]. They also used hyperpolarized ¹³C-acetate to monitor its influx: because the acetate chemical shift depends on pH, two peaks corresponding to extracellular (low pH) and intracellular (pH~7) acetate could be observed by NMR in real-time [1118]. They performed similar experiments on

*E. coli*, revealing subtle metabolic differences in different growth phases [1120] and strains [1111]. Later studies on hyperpolarized pyruvate metabolism revealed its conversion to acetate in *E. coli* and *S. aureus*, in an opposite fashion to pyruvate metabolism in human cells (see above) [1103]. d-DNP experiments were also carried out on *Trypanosoma brucei* (the sleeping-sickness parasite) to study pyruvate conversion to alanine, and revealed important differences between the form of *T. brucei* found in the insect vector and the form found in the bloodstream of the animal host [1121]. There have also been studies on pyruvate metabolism using d-DNP on motile human sperm [1122] and T lymphocytes [1123].

A number of other hyperpolarized metabolites were used to carry out d-DNP experiments [33,1081,1124,1125], e.g. fumarate to quantify fumarase activity associated with necrosis [1080,1126,1127], ascorbic acid to evaluate intracellular redox potential [487,488], α-ketoglutarate to quantify its conversion to 2-oxoglutarate by the isocitrate dehydrogenase 1 (IDH1) or to glutamate by the branched-chain amino acid transaminase 1 (BCAT1) in glioma cells [1128,1129], arginine to detect its degradation in urea by arginase in myeloid-derived suppressor cells [1130], and glutamine to observe oncogenic glutaminase activity [1131,1132]. A more complete list of hyperpolarized metabolites that have been used, including for *in vivo* applications, can be found in a recent review from Sando and colleagues [1133]. Other nuclei can also be usefully hyperpolarized. ¹³C-hyperpolarization can be progressively transferred to covalently bound protons after dissolution, which can be useful in cases where ¹³C-NMR signals from substrates and products overlap [1134]. ¹³³Cs has been shown to yield different peaks in the extracellular medium and in yeast [441] and erythrocytes [442]. Indeed, the ¹³³Cs⁺ chemical shift is sensitive to osmolarity, which could make it useful as a cytosolic sensor for this parameter. It was recently used to report cation channel activation in erythrocytes. ¹⁵N-labeled compounds have also been tested, such as choline or carnitine, whose slower T1 relaxation (about 5 times slower than that of pyruvate) render