# Molecular distinction of cell wall and capsular polysaccharides in encapsulated pathogens by in situ magic-angle spinning NMR techniques

Alons Lends, Gaelle Lamon, Loic Delcourte, Aude Sturny-Leclere, Axelle Grélard, Estelle Morvan, Muhammed Bilal Abdul-Shukkoor, Mélanie Berbon, Alicia Vallet, Birgit Habenstein, et al.

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Alons Lends, Gaelle Lamon, Loic Delcourte, Aude Sturny-Leclere, Axelle Grélard, et al. Molecular distinction of cell wall and capsular polysaccharides in encapsulated pathogens by in situ magic-angle spinning NMR techniques. Journal of the American Chemical Society, 2025, 147 (8), pp.6813-6824. 10.1021/jacs.4c16975 . hal-05386051

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# Molecular Distinction of Cell Wall and Capsular Polysaccharides in Encapsulated Pathogens by In Situ Magic-Angle Spinning NMR Techniques

Alons Lends, Gaelle Lamon, Loic Delcourte, Aude Sturny-Leclere, Axelle Grélard, Estelle Morvan, Muhammed Bilal Abdul-Shukkoor, Mélanie Berbon, Alicia Vallet, Birgit Habenstein, Erick J. Dufourc, Paul Schanda, Vishukumar Amanianda,* and Antoine Loquet*

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Cite This: https://doi.org/10.1021/jacs.4c16975

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**ABSTRACT:** Pathogenic fungal and bacterial cells are enveloped within a cell wall, a molecular barrier at their cell surface, and a critical architecture that constantly evolves during pathogenesis. Understanding the molecular composition, structural organization, and mobility of polysaccharides constituting this cell envelope is crucial to correlate cell wall organization with its role in pathogenicity and to identify potential antifungal targets. For the fungal pathogen *Cryptococcus neoformans*, the characterization of the cell envelope has been complexified by the presence of an additional external polysaccharide capsular shell. Here, we investigate how magic-angle spinning (MAS) solid-state NMR techniques increase the analytical capabilities to characterize the structure and dynamics of this encapsulated pathogen. The versatility of proton detection experiments, dynamic-based filters, and relaxation measurements facilitate the discrimination of the highly mobile external capsular structure from the internal rigid cell wall of *C. neoformans*. In addition, we report the *in situ* detection of triglyceride molecules from lipid droplets based on NMR dynamic filters. Together, we demonstrate a nondestructive technique to study the cell wall architecture of encapsulated microbes using *C. neoformans* as a model, an airborne opportunistic fungal pathogen that infects mainly immunocompromised but also competent hosts.

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## INTRODUCTION

Pathogenic fungi play deleterious roles in many living organisms, e.g., fungal infections cause approximately 6.5 million cases per year and 3.8 million deaths,¹ costing about $12 billion per year. Severe fungal infections mainly occur in critically ill patients (i.e., those suffering from burns, immunocompromised diseases such as AIDS, or those undergoing immunosuppressive interventions).² They are primarily caused by species belonging to the genera of two yeasts (*Candida* and *Cryptococcus*) and one filamentous fungus (*Aspergillus*).

*Cryptococcus* causes cryptococcosis and cryptococcal meningitis (accounting for ~200,000 deaths per year) that occur primarily by inhalation of the propagules, leading to pneumonia-like symptoms and, in the worst case, meningitis when the central nervous system is affected.³ *Cryptococcus neoformans* is the major pathogen of this fungal genus. The cell envelope of *C. neoformans* is unique in the fungal kingdom as it possesses, besides the cell wall, an additional structure called the capsule. This capsule is secreted to surround the pathogen and is highly sensitive to the external environment.⁴⁻⁶ Different variants

(serotypes⁷) of *C. neoformans* have been identified on the basis of the immunologic properties of capsular polysaccharides. The capsule forms a dynamic organelle that acts as a crucial virulence factor,⁸ while mutants lacking capsular structure are avirulent.⁹,¹⁰

The antifungal repertoire to treat *C. neoformans* infection is limited to amphotericin B, flucytosine, and fluconazole; the first two are toxic and expensive, whereas there is the emergence of fungal resistance against fluconazole.¹¹ Echinocandins target specific enzymes responsible for the synthesis of fungal cell wall component β-(1,3-glucan);¹² however, they fail to efficiently counter *C. neoformans* *in vitro* as well as *in vivo*.¹³ Developing analytical tools to investigate the structural integrity and dynamics of the *C. neoformans* cell wall and its

Received: November 28, 2024

Revised: February 6, 2025

Accepted: February 12, 2025
capsule is of primary importance to understand antifungal drug resistance and how surface polysaccharides interact with host immune factors. During the past three decades, the standard protocols used for the extraction of cell wall polysaccharides have relied on successive chemical extraction, lyophilization, and sometimes enzymatic treatments.¹⁴ Unfortunately, for encapsulated C. neoformans, the presence of the capsular shell requires more complex protocols to specifically extract capsular polysaccharides.¹⁵⁻²⁰ It is fundamental to note that capsular polysaccharides are not retrieved from the capsule itself but rather from culture supernatants.²¹ Therefore, there is a need to develop biophysical tools to monitor capsular polysaccharides in situ.

Here, we take advantage of magic-angle spinning (MAS) nuclear magnetic resonance (NMR) techniques to investigate the structure and dynamics of the C. neoformans yeast cell envelope. Importantly, our study was performed on intact encapsulated yeasts, avoiding the extraction of polysaccharides from the cell wall and capsule, to derive relevant information on the entire cell envelope under near-native conditions. We employed a minimal quantity (~500 ug) of yeast combined with fast MAS (>100 kHz) and direct ¹H detection. Our NMR data reveal that atomic resolution structural and dynamic information on cell wall polymers, lipid droplets, and capsule polysaccharides can be uncovered in a highly selective manner, suggesting that this technique could offer a new avenue to investigate the C. neoformans envelope regarding its infectivity and the effect of antifungal drugs at high resolution.

## ■ RESULTS AND DISCUSSION

**Predominance of Polysaccharides in the Rigid Core of C. neoformans Cell Envelope.** We used a liquid-culture minimal medium composed of ¹³C-labeled glucose and ¹⁵N-labeled NH₄Cl to grow C. neoformans yeasts (H99 isolate, var. grubii) under capsule-inducing conditions. The details are described in the *Materials and Methods* section. The fluorescence micrograph (Figure 1A) clearly indicated two separate compartments of a cell envelope, where cell wall chitin was stained with Calcofluor white (in blue), and the capsule was immunolabeled with anticapsular antibodies (in green). The schematic structure of the cell is shown in Figure 1B.

First, to probe the most rigid and immobilized polysaccharides of the cell wall associated with large dipolar couplings, we employed cross-polarization (CP) experiments. All the ¹H-detected pulse sequences used in this study are shown in Figure S1. The 2D CP hCH spectrum of C. neoformans cells (Figure 2B) recorded at a MAS rate of 100 kHz on an 18.8 T NMR magnet showed well-dispersed but rather broad lines for the ¹H dimension (full-width at half-height 460 ± 160 Hz) (Figure S2A). The chemical shift range indicates a strong predominance of carbohydrate signals from the cell surface. Despite the high MAS frequency, the ¹H signals were not sufficiently well resolved in the 2D CP hCH spectrum for precise analysis due to the severe overlap of many peaks. This is mostly the result of strong ¹H and ¹³C chemical shift overlaps for polysaccharides, for which substantial chemical modifications (nature of the side chains, change in the subunit branching, and presence of structurally different conformations) unfortunately often lead to small changes in chemical shifts, in contrast to proteins where ¹³C chemical shifts for aliphatic carbon spread over ~60 ppm. To improve the spectral resolution, we increased the dimensionality by acquiring a 3D spectrum to encode an additional J-coupling-

based ¹³C–¹³C polarization transfer with isotropic WALTZ-16 mixing.²²,²³ This 3D CP hCCH TOCSY spectrum (the 2D ¹³C–¹³C projection is shown in Figure 2D) led to a gain of spectral resolution and allowed us to distinguish several spin systems based on their distinct chemical shifts and correlation patterns. As illustrated with the 2D ¹³C–¹H strips presented in Figure 2B for β-1,3-glucan, the improvement in the spectral resolution in the indirect ¹³C dimension is clearly beneficial, and the correlation patterns observed in the hCCH experiment allowed us to connect ¹³C–¹³C pairs to their respective ¹H chemical shifts and to establish C₁–C₂–C₃–C₄–C₅–C₆ spin topologies. Overall, we identify three major spin systems belonging to the polysaccharide families: α-1,3-glucan, β-1,3-glucan, and chitin (vide infra for the detailed assignment procedure).

To identify the three main spin systems (the molecular structures are shown in Figure 2E,F), we investigated the patterns of the spin topologies observed in the 3D CP hCCH experiment (¹H–¹³C 2D slices are shown in Figure 2C,D and ¹³C–¹³C projection in Figure 2E), in combination with a manual inspection of ¹³C and ¹H chemical shift values. We started our assignment with the identification of anomeric carbons C₁ observed at 103.1, 105.8, and 106.2 ppm positions and connected them to their attached proton H₁. Taking advantage of the TOCSY polarization pathway, we connected them to the C₂ and C₃ positions and found two spin systems with C₃ chemical shifts at ~90 and ~75 ppm. Polysaccharides substituted at the C₃ position with the O–C linkage have C₃ downfield chemical shifts at ~87–91 ppm; thus, we assigned them to (1,3)-linked D-glucose moieties. Without branching, the C₃ resonances are shifted more upfield. To disambiguate the α- and β-configuration of the two spin systems with (1,3)-linked D-glucose moieties, we used C₁ chemical shifts, which are sensitive to the anomeric configuration, and assigned ¹³C shifts at ~103 and ~106 ppm for α-1,3-glucan and β-1,3-glucan, respectively. A third major spin system was identified

<figure><img src="image_10.png" /><figcaption>**Figure 1.** Molecular organization of the C. neoformans envelope. (A) Fluorescence microscopy of C. neoformans yeast cells; the cell wall chitin is stained in blue with Calcofluor white, and the capsule (in green) was immunolabeled using primary anticapsular antibodies (mouse monoclonal, E1) and secondary fluoresceine isothiocyanate-conjugated antimouse IgG. (B) Concentric schematic organization of C. neoformans yeast cells.</figcaption></figure>
<figure><img src="image_11.png" /><figcaption>**Figure 2.** C. neoformans cell wall polysaccharides form a rigid network. (A) Concentric schematic view of polysaccharides detected by CP-based ¹H MAS NMR detection. (B) 2D CP hCH spectrum recorded at 100 kHz MAS. (C) 2D ¹H–¹³C representative strips from the 3D hCCH experiment, exemplified by β-1,3-glucan resonance assignment. (D) 2D ¹³C–¹³C projection of the 3D hCCH experiment. Molecular structures of AGlc, BGlc (E), and Ch (F). (G) DQ-SQ ¹³C–¹³C spectrum recorded at 11 kHz MAS. AGlc, α-1,3-glucan; BGlc, β-1,3-glucan; and Ch, chitin; Cs, chitosan.</figcaption></figure>

and assigned to chitin based on its C₂ contribution, which shifted up to 55 ppm when bonded to an N-acetyl group. It has been reported that due to strong structural polymorphism, the use of ¹³C chemical shifts obtained with ¹³C-detected experiments at moderate MAS frequencies still poses a challenge to distinguish and identify unambiguously the structural forms of chitin in fungal cell walls.²⁴ To evaluate whether chitin ¹H signals derived from the 3D CP hCCH could help investigate the local structural polymorphism, we compared our ¹H chemical shifts obtained *in situ* on C. neoformans yeasts with those reported earlier on crystallized α- and β-chitin samples,²⁵ for which the major shift difference is associated with the H₁ position (−0.7 ppm between α- and β-chitin). Here, the line width for H₁ is large (~580 Hz, i.e., ~0.72 ppm), which reinforces the idea that the native chitin organization found in the cell wall of microorganisms such as fungi or yeasts undergo intense structural polymorphism, with chemical shift values covering both α- and β-structural forms.

To estimate/compare the degree of structural homogeneity for the three main polysaccharides, we compared the ¹H and ¹³C line widths of the H₁–C₁ signals from the 2D CP hCH (Figure S2A). We found line widths of 192 Hz ± 6 Hz, 264 Hz ± 8, and 240 Hz + 7 Hz (¹H) and 388 Hz ± 12, 488 Hz ± 15, and 580 Hz ± 17 Hz (¹³C) for α-1,3-glucan, β-1,3-glucan and chitin, respectively, suggesting a comparable local structural homogeneity between the three biopolymers. An additional complexity of chitin analysis is the putative presence of chitosan, the deacetylated form of chitin (Figure S3A). Previously, chitosan has been identified as a minor but crucial component of the cryptococcal cell wall.²⁶,²⁷ Although an unambiguous assignment of chitosan was not possible solely based on ¹H-(¹³C)-¹³C spin systems, a careful inspection of a 2D ¹³C–¹³C INADEQUATE (Figure 2G) led to the identification of a minor contribution for chitosan, at ¹³C chemical shifts previously reported for chitosan identified in the fungus *Rhizopus delemar*.²⁸ Our results reveal a lower molar ratio of chitosan compared to chitin, in line with previous
<figure><img src="image_12.png" /><figcaption>**Figure 3.** MAS NMR detection of mobile capsular polysaccharides on *C. neoformans* yeast cells. (A) Concentric schematic view of polysaccharides detected by INEPT-based ¹H MAS NMR. (B) Molecular structures of Manp, Xylp, and GlcpA from GXM. (C) Relative intensity ratio between CP, INEPT, and direct polarization ¹³C signals, for *C. neoformans* and *A. fumigatus*. (D) 2D scalar hCH experiment recorded at 100 kHz MAS. (E) 2D ¹³C–¹³C DQ-SQ experiment showing spectral overlap for GlcpA and Xylp for their C₁–C₂ correlations.</figcaption></figure>

reports on fungal and yeast cells,²⁹,³⁰ including recent chitin/chitosan quantification in *Aspergillus fumigatus* by MAS NMR,²⁴,³¹ but differ from previous absorbance measurements at 585 nm on *C. neoformans*,³² for which chitosan was found to be in higher amount than chitin. A strict comparison of these studies is challenging because in our work, we estimated the relative amount of chitin/chitosan based on NMR intensities from *in situ* measurements on cells, while Banks et al.³² used multiple steps of chemical treatments to derivatize polysaccharides and further hydrolyze them to GlcNAc.

The ¹⁵N chemical shift is a useful spectroscopic reporter to disambiguate chitin to chitosan signals, based on distinct chemical shift values observed for amine (~33 ppm) and amide (~120 ppm) groups, respectively, for chitosan and chitin. However, such spectral analysis based on the comparison of ¹⁵N peak intensities could lead to an overestimation of chitin content since the amide region might also contain signals from proteins in the cell envelope. To analyze this, we recorded a 2D CP hNH spectrum (Figure S3B) using fast MAS to inspect nitrogen-containing molecules in more detail. We found a broad distribution of HN correlations in the range of ~7–10.5 ppm, suggesting that the spectral contribution observed at a ¹⁵N chemical shift of ~120 ppm might also encode for minor amino acid signals. Indeed, the cell wall of *C. neoformans* contains mannoproteins,⁵ and most likely the spectrum comprises peaks arising from the mixture of indistinguishable protein and chitin NH resonances. A similar ¹⁵N chemical shift range was observed for proteins in the cell walls of zygomycotic fungus and corynebacteria.³³,³⁴ This finding was confirmed by a careful inspection of the 2D CP hCH experiment (Figure 2B), which showed several minor signals in the chemical shift ranges of 4.5–5 ppm (¹H dimension) and 55–65 ppm (¹³C dimension), a spectral area where the amino acid signals are observed. We employed an additional MAS NMR approach to test whether amino acid signals significantly contributed to the rigid NMR signals observed in the CP-based experiments. We recorded two 2D versions of the H–N–C correlation experiments (3D

hCaNH²⁵ and hNCaH²³), which are typically employed for protein backbone chemical shift assignments (pulse sequences are shown in Figure S1C,D), where magnetization is transferred between the ¹⁵N and ¹³Cα spins. The resulting peak distribution showed a wide range of ¹³C chemical shifts with ¹³C peaks spanning outside of the previously reported chemical shift region for the C₂ position of chitin species (2D ¹H–¹³C projections from the H–N–C correlation spectra are shown in Figure S3C),²⁴,³⁶,³⁷ strengthening the hypothesis of the detection of immobilized proteins in the cell wall of *C. neoformans* using CP-based ¹H detection.

To estimate the relative amount of each polysaccharide in the *C. neoformans* cell wall, we took advantage of the ¹H dimension and extracted 1D ¹H traces from the 2D CP hCH. We established the relative molar fractions of α-1,3-glucan, β-1,3-glucan, and chitin to be, respectively, 41, 46, and 13% (±3%) in the cell wall. Such composition clearly differs from our previous observations of *A. fumigatus* conidia³⁷ (21, 67, and 12% of α-1,3-glucan, β-1,3-glucan, and chitin, respectively). Although *A. fumigatus* possesses three α-glucan synthases (i.e., AGS1, AGS2, and AGS3),³⁸,³⁹ only one cryptococcal enzyme responsible for the production of α-1,3-glucan is known (AGS1⁴⁰). The high molar ratio in favor of α-1,3-glucan, also associated with narrow lines suggesting structural homogeneity, indicates the predominant role of this polysaccharide in the molecular stability of the cell wall. Indeed, *C. neoformans* lacking α-1,3-glucan (*ags1Δmutant*) showed temperature sensitivity, difficulty in growth, capacity loss to produce a capsule as well as failed to grow in a mouse model of infection.⁴¹

β-1,3-Glucan is the most abundant polysaccharide in the fungal cell wall, and targeting its synthesis via the inhibition of specific synthases is a common route to develop antifungal drugs.⁴² Controversies exist in the literature on the amount of this polysaccharide in the cell wall of *C. neoformans* and the relevance of its detection in the diagnosis of cryptococcal meningitis.⁴³⁻⁴⁵ In our study, we measured the relative amount of β-1,3-glucan to be ~46%. Studies on glucan quantification in
<figure><img src="image_13.png" /><figcaption>**Figure 4.** MAS NMR detection of *C. neoformans* GXM and GXMGal. (A) 2D hCcH TOCSY experiment recorded at 100 kHz MAS, providing polarization transfers to connect the C₁–H₁ signals to their neighboring ¹³C sites. (B) Polarization transfer in hCcH for C₂H₂ and C₄H₄ of Xylp. (C) Two low-intensity C₂–H₂ correlations, encoding two Manp units in two different O-substitutions. (D) Two Gal spin systems were assigned in the 2D hCcH TOCSY experiment.</figcaption></figure>

cryptococcal cell walls are extremely scarce. The presence of β-
1,3-glucan has often been assessed by qualitative measure-
ments such as using anti-β-1,3-glucan antibodies,⁴⁶,⁴⁷ although
these antibodies might directly affect fungal growth and
capsule formation.⁴⁷ Our NMR measurements provide an
unprecedented tool to estimate the molar ratio of β-1,3-glucan
and distinguish this polysaccharide from α-1,3-glucan and
chitin/chitosan. Doering and co-workers reported a semi-
quantitative analysis of alkali-soluble (AS) and alkali-insoluble
(AI) fractions of the *C. neoformans* cap59 mutant after several
steps of chemical extractions,⁴¹ with an unequilibrated
distribution of α-1,3-glucan only detected in the AS fraction
and β-1,3-glucan in both AS and AI fractions. Our results
suggest comparable rigidity between the two glucans and
different molar ratios in the cell wall. Our observations are
understandable as (1) the cap59 mutant is an acapsular strain
while our study focuses on a capsular wild-type/reference
strain, (2) chemical analysis after cell wall extraction can
underestimate glucan amounts;⁴⁸ and (3) there is no direct
correlation between the molecular mobility of the poly-
saccharide and their preferential presence in AS or AI
fractions. Similar assumptions were elaborated based on
solid-state NMR studies complemented with chemical analysis
of AI and AS fractions of *A. fumigatus*.⁴⁸

**C. neoformans** Capsular Polysaccharides Undergo Fast Molecular Motions. Early work led to the identification and structural characterization of the two polysaccharides as the main components of the *C. neoformans* capsule (Figure 3A). Upon ethanol extraction and further purification by ion-exchange chromatography after various chemical treatments, previous studies using solution NMR<sup>16,49–53</sup> identified glucuronoxylomannan (GXM), a complex polysaccharide, as

the major component in the capsule (Figure 3A). GXM
contains (1,3)-α-D-mannopyranan (Manp) backbone substi-
tuted with β-D-glucopyranosyl-uranic acid (GlcpA) in β-1,2-
linkage and β-D-xylopyranosyl (Xylp) units with β-1,2 and β-
1,4 linkages varying with the serotype.<sup>16,54</sup> The chemical
structure of Manp, Xylp, and GlcpA are shown in Figure 3B. In
addition to GXM, glucuronoxylomannogalactan (GXMGal) as
a minor capsular polysaccharide was deciphered using liquid-
state NMR and GC/MS,<sup>19,21,55</sup> consisting of 1,6-linked α-
galactan (Galp) with substitution at O-3 to branched Manp
and at O-2 with galactofuranose (Galf).<sup>56,57</sup>

Given that the NMR signals of the capsule could not be
detected with dipolar-coupling-based experiments, we explored
whether pulse sequences based on *J*-coupling transfer would
allow retrieving capsule signals of Manp, Xylp, and GlcpA. The
*J*-coupling-based INEPT experiment allows the detection of
sites undergoing large-amplitude motions, particularly if they
occur on submicrosecond time scales.<sup>58,59</sup> We employed *J*-coupling-based polarization transfer with ¹H detection for 2D
insensitive nuclei enhanced by polarization transfer (INEPT)
hCH<sup>60</sup> and 2D INEPT hCcH TOCSY (with DIPSI-2
homonuclear ¹³C–¹³C transfer<sup>22</sup>) experiments (pulse sequen-
ces are shown in Figure S1, full length spectrum in Figure S5).
First, to perform a global assessment of the presence of mobile
polysaccharides in *C. neoformans*, we compared the relative
intensity of 1D recorded with CP, INEPT, and direct
polarization (DP) transfer (Figure 3C). We found that,
compared to *A. fumigatus* conidia we investigated earlier,<sup>37</sup> C.
*neoformans* yeasts exhibit a much stronger INEPT contribution
with respect to the DP signal. This observation suggests that
the proportion of mobile biopolymers in the cryptococcal cell
envelope is higher than that in the *A. fumigatus* conidia, where
<figure><img src="image_14.png" /><figcaption>**Figure 5.** Detection of *C. neoformans* triglycerides by ¹H-detected MAS NMR. (A) Concentric schematic view of lipid droplets in *C. neoformans* putatively located in the endoplasmic reticulum. (B, E) Detection of triacylglycerols by ¹H-detected MAS NMR, except for the 2D hCCH spectra recorded at 100 kHz MAS. (C) ¹³C to ¹H connectivity for diastereotopic protons in TAG. The dashed lines indicate the CCH correlations. (D) 2D hCCH TOCSY spectrum. (F) The putative molecular formula of the detected lipid structure comprises a triacylglycerol group, at least one saturated double bond, and an aliphatic chain. The carbon position numbering was designed to follow the NMR signal assignments.</figcaption></figure>

mobile galactomannan and galactosaminogalactan were detected on the surface.³⁶,⁴⁸ Then, we recorded a 2D J-coupling-based hCH spectrum (Figure 3D), which led to the observation of five ¹³C-¹H correlations in the C₁ region. Importantly, the spectral pattern observed in the J-coupling-based 2D hCH does not match with the one obtained using the CP-based 2D hCH spectrum (Figure 2B), indicating that (i) α-1,3-glucan, β-1,3-glucan, and chitin are not observed as mobile species in the *C. neoformans* cell envelope and (ii) other distinct biopolymers are part of the cell envelope as highly mobile entities. All signals detected in the 2D J-coupling-based hCH spectrum exhibit very narrow line widths in the ¹H (~30 Hz) and ¹³C (~130 Hz) dimensions. Such narrow lines contrast with those that are much broader than those observed in the 2D CP hCH spectrum. This observation reflects an important dynamical behavior of the detected molecules in the J-coupling-based hCH due to the efficient averaging of dipole couplings for mobile molecules. The incorporation of an additional ¹H dimension is highly advantageous, as demonstrated by the two C₁H₁ signals of Xylp and GlcpA highlighted in Figure 3D (vide infra for the assignment procedure). These signals are well separated in the ¹H dimension, but undistinguishable in a 2D ¹³C-¹³C DQ-SQ INADEQUATE (Figure 3E) because their ¹³C chemical shift values are too close, even though the INADEQUATE experiment does not have a diagonal due to the DQ dimension. Analogous to the methodology presented for CP-based ¹H detection experiments, we added a J-coupling-based ¹³C-¹³C DIPSI-2 isotropic mixing to achieve an additional ¹³C-¹³C through-bond polarization transfer pathway,²² followed by ¹H detection. The resulting 2D hCcH TOCSY experiment (different parts of this spectrum are shown in Figure 4), used in combination with the J-coupling-based 2D hCH, provided an efficient spectroscopic tool to track the spin

topologies of mobile biopolymers present at the *C. neoformans* cell surface.

We started the assignment process by tracking the five spin systems based on their ¹³C → (¹³C) → ¹H connectivity. We first considered the signals observed in the region 95–106 ppm, a range that typically encodes anomeric C₁ carbons. In this spectral region, we observed five major C₁ contributions and linked the ¹³C chemical shifts to their chemically bonded ¹H resonances (Figure 3D). The polarization pathway encoded in the hCcH experiment allowed us to connect C₁ (acting as the polarization source) to nearby carbon positions by utilizing ¹³C-¹³C connectivity, and finally to detect them via their bonded protons. As illustrated in Figure 4A, the TOCSY scheme allows a polarization propagation to all carbon positions, i.e., C₂, C₃, C₄, C₅, and C₆ (except for Xylp, which has only five carbon), and then directly detect their bonded proton, i.e., H₂, H₃, H₄, H₅, and H₆ (except for Xylp). The combined use of 2D hCH and hCcH spectra allowed us to identify most ¹³C resonances of the four main spin systems in *C. neoformans*, and to link them to their bonded ¹H resonances. For one spin system (later assigned to GlcpA), its close H₁ chemical shift to water impeded further inspection due to the spectral overlap with strong water peaks. First, we identified a spin system topology encoding a 5-membered pyranosyl carbon ring that we assigned to xylose (Xylp), characterized by a C₅-H₅ cross-peak with typical chemical shifts at ~67.5 and ~3.3 ppm,⁴⁹,⁵⁰,⁵³ and importantly by an anomeric C₁ located at the farthest downfield (~105.8 ppm). The nature of the Xylp β-glycosidic linkage, which could be either (1 → 2)-linked or (1 → 4)-linked to Manp, directly impacts the value of the Xylp C₁ chemical shift.⁴⁹,⁵³ We detected an Xylp C₁ chemical shift at ~105.8 ppm, which suggests its substitution to Manp at position O-2, while O-4 substitution would have led to a ¹³C chemical shift of ~106.7 ppm.⁴⁹ Figure 4B shows
polarization transfer starting from C₃–H₃ and C₄–H₄ to their
adjacent carbons.

The farthest upfield H₁ resonance was assigned to Manp, based on previous NMR studies⁴⁹,⁶¹ and confirmed by the presence of methylene signals (CH₂O) at ~64 ppm (no methylene in Xylp and GlcpA), which we assigned to Manp C₆ (Figure 4C). Based on previous studies,⁵³,⁶² it has been shown for the C. neoformans serotype A that Manp has virtually two distinct local environment, one substituted at position 2 with GlcpA and one substituted with Xylp, respectively, named ManG and ManX following the notation used by Cherniak and co-workers in ref 63. We detected two signals of low intensity encoding for C₂–H₁ correlations but linked to a unique H₁ chemical shift, suggesting two putative C₂ local environments, at ~72.5 and ~80 ppm (Figure 4C). This suggests that the NMR chemical shift at the C₂ position is sensitive to O-2 substitution. The molar ratio for serotype A is 2:3:1 for Xyl/ Man/GlcA, and accordingly, we assigned the highest intensity signal at ~72.5 ppm to ManX and the signal at ~80 ppm to C₂ linked to the O-2 substitution in ManG. The two remaining spin systems were putatively assigned to Gal units, the main carbohydrate component of GXMGal,¹⁶,⁵⁵⁻⁵⁷ accounting for ~50% of the monosaccharide composition.⁵⁶ The detected C₁ chemical shifts at ~99 and ~95.5 ppm (Figure 4D) suggest an α form (α-Gal), while a β form (β-Gal) would have led to a signal ~105–110 ppm. Although the solution NMR chemical shifts are available for various Gal units in GXMGal,¹⁶,⁵⁵⁻⁵⁷ the extent of heterogeneity is important with the concomitant presence of α- and β-forms for galactan and galactofuranose associated with various substitutions, and the presence of a neutral galactoxylomannan (NGalXM) that led to the challenging solution NMR analysis of purified GXMGal and NGalXM.¹⁹,⁵⁵⁻⁵⁷ Consequently, an unambiguous distinction between galactopyranose and galactofuranose was not possible solely based on their spin system connectivity. Therefore, dynamically filtered ¹H signals provided a tool to investigate capsular polysaccharides and discriminate them from their counterparts in the cell wall.

**In Situ ¹H Detection and Identification of C. neoformans Triglycerides.** Several signals encoded in the 2D INEPT-based hCH and hCcH experiments could not be linked to the polysaccharide structures. In our recent MAS NMR study of *A. fumigatus* conidia, in their swollen and germinating morphotype,³⁷ we uncovered that the ¹³C-detected DP INADEQUATE experiment constitutes a powerful tool to access mobile lipids from the fungal cell envelope. For *A. fumigatus*, lipid assignment relied on identifying SQ-DQ ¹³C–¹³C connectivity patterns, especially those linking C–C unsaturation to the aliphatic acyl chain. The ¹³C lipid aliphatic chemical shift range is narrow (~20–38 ppm) and could be the source of spectral misinterpretation. To improve the *in situ* spectral identification of cellular lipids in *C. neoformans*, we inspected in more detail the *J*-coupling-based 2D hCH and hCcH (Figure 5) spectra and found several signals with chemical shifts and correlation patterns that suggest the presence of lipidic spin system topology. First, we identified a characteristic connection between carbons at 64.4 ppm, which is connected to two diastereopic protons at 4.3 and 4.1 ppm (highlighted in pink in Figure 5B). We established a connection to a ¹³C signal at 71.8 and 5.2 ppm in the ¹H dimension (shown in Figure 5C). This pattern was assigned to a (–CH₂–O–) group connected to a (O–CH₂–CH–O–) group, corroborated by similar chemical shifts that we found

for two glycerol moieties in fungal lipids.³⁷ Then, a characteristic pattern was identified based on the polarization transfer connectivity between adjacent methylene groups, leading to the identification of two –CH₂–CH₂– connectivities. We found that the addition of ¹H–¹³C correlations presented here favors the rapid identification of – (¹³C–¹³C–¹H)– patterns to connect and distinguish adjacent CC pairs, in comparison to the 2D ¹³C-detected CC DP INADEQUATE experiment. We then identified a ¹³C–¹³C double bond between carbons C₇ and C₆ (the carbon position numbering was set to start with C₁ at the carbonyl position) based on their characteristic chemical shift values at ~130–132 ppm and their corresponding vinylic protons at ~5.5 ppm (Figure 5E). Taken together, a clear connection pattern for an acyl chain with an unsaturation was established from C₂H₂ to C₁₂; the latter connected to H₁₂ with a chemical shift characteristic of methyl protons (~0.9 ppm).

The only missing connection in the *J*-coupling-based hCcH experiment is between C₄ and C₅, and we speculate lower sensitivity arising from the specific dynamical regime. Here, the use of ¹H-detected experiments allowed us to establish a –CH₂–CH₂–(CH₂)ₓ–CH₂–CH=CH–CH₂–CH₂–CH₂–CH₂–CH₃ spin topology, starting from C₂ to C₁₂. We then linked the first C₂ methylene to carbon at 174.4 ppm, which we assigned to an ester carbonyl. Altogether, this network of interconnected ¹H and ¹³C spin systems led to the identification of a lipid structure that reassembled the characteristics of triacylglycerol (TAG) (Figure 5F). Thus, analogous to our previous report on *A. fumigatus*, the main mobile lipid component assigned to encapsulated *C. neoformans* is TAG. The observation of TAG as a major lipid in *C. neoformans* is in line with previous NMR studies from Stark and Casadevall on nonmelanized, melanized, and so-called melanin ghost samples.⁶⁴,⁶⁵ Note that sterol esters, previously detected in *C. neoformans* melanin ghost, were not detected in our study using whole *C. neoformans* yeasts. TAG is commonly known as an energy source in eukaryotic cells and it has been proposed that its content is unusually high in *C. neoformans*.⁶⁶ Its precise and putative dual location is still a matter of debate, with TAG located in the endoplasmic reticulum and putatively in specific organelles, namely lipid droplets.⁶⁷,⁶⁸ Because we did not detect any phospholipids or sterol ester in our 2D hCcH spectrum but rather TAG, we hypothesize that the dynamic filter created in this experiment to promote the NMR detection of highly mobile molecules has efficiently filtered out lipids in immobilized compartments (i.e., the lipid membrane) to favor lipids in highly mobile subcompartments (i.e., droplets). Interestingly TAG molecules could not be seen in CP-based experiments. This suggests that these triglycerides maintain an important level of mobility without having a smaller population in a more static state. The same observation was made for TAG found in *A. fumigatus*.³⁷

**In situ Dynamics Measurements of C. neoformans Polysaccharides.** MAS NMR spectroscopy constitutes a powerful experimental method to measure and quantify the dynamics of biomolecules in complex samples.⁶⁹⁻⁷¹ Two approaches are commonly employed: one quantifies the dipolar-coupling strength. The motion of the vector connecting the two spins (often a chemical bond) reduces the coupling strength, and this reduction is directly proportional to the amplitude of motions faster than a few tens of microseconds. We quantified the amplitude of motion for the cell wall part (α-1,3-glucan, β-1,3-glucan, and chitin), using a ¹H–¹³C
dipolar-coupling measurement with the REDOR scheme<sup>72,73</sup>
(Figure S4). The determined dipolar-coupling strength is close
to the rigid-limit value, which shows that these glycans have
only very limited mobility on time scales of up to tens of
microseconds.

Another class of experiments quantifies the spin relaxation rate constants, which are sensitive to the amplitudes and time scales of mobility. The rotating-frame spin relaxation (R₁ρ) is particularly sensitive to motion in the range of ca. 0.1–100 µs, i.e., high R₁ρ rate constants are expected when the site undergoes motion on this time scale.⁷⁴ In systems with high proton density, the rate constant may contain contributions not due to dynamics (dipolar dephasing). While this effect clearly persists for ¹H even at MAS frequencies of 100 kHz, and ¹H R₁ρ rate constants only provide a semiquantitative measure of dynamics.⁷⁵

The R₁ρ relaxation rates for α-1,3-glucan, β-1,3-glucan, and
chitin were found to be, respectively, 11.0 ± 1.3, 8.0 ± 1.5, and
17 ± 7 s⁻¹ for ¹³C (13 kHz spin-lock field strength) and 55 ±
11, 93 ± 6, and 102 ± 1 s⁻¹ for ¹H (Figure 6). The ¹³C R₁ρ

<figure><img src="image_15.png" /><figcaption>**Figure 6.** *In situ* polysaccharide dynamics measurement in the C. neoformans cell wall and capsule. ¹³C CP R₁ρ (A) and ¹H CP R₁ρ (B) rates of cell wall polysaccharides are determined at a MAS frequency of 105 kHz. (C) ¹³C INEPT R1 of capsular polysaccharides.</figcaption></figure>

values on *in situ* polysaccharides are similar to those obtained
for Cα carbons in rigid protein elements, such as in secondary
structures of fibrillar protein assemblies<sup>76</sup> and microcrystalline
proteins.<sup>77</sup> The differences suggest an increasing amount of μs
motion from α-1,3-glucan to β-1,3-glucan to chitin. Chitin,
which shows more μs motion, is often presented as the main
carbohydrate component in the interaction with the
cytoplasmic membrane,<sup>78</sup> i.e., the most buried in the cell
wall. However, analogous to what others<sup>36,48</sup> and we<sup>37</sup> found
for A. fumigatus, chitin molecules can form a large number of

allomorphs within the cell wall,²⁴ which may explain the
slightly increased µs mobility. We also measured the ¹³C R₁ρ
rate constants in a J-coupling-based ¹H–¹³C experiment, where
the correlation peaks for Manp, Glc*pA*, Xylp, and the two Gal
spin systems were resolved. The ¹³C R₁ρ rate constants for
these GXM/GXMGal polysaccharides are an order of
magnitude lower than those for the cell-wall sites (~1 to 1.6
s⁻¹, Figure 6C), showing that the glycans of the capsule
undergo significant dynamics.

■ CONCLUSIONS

Increasing evidence suggests that cell wall polysaccharides play
a predominant role in life-threatening infections caused by
fungal pathogens. In this context, C. neoformans is unique, as it
is the only encapsulated fungal pathogen. The capsule poses
complications since there are no efficient protocols to reliably
extract constituent components in a specific manner. As a
consequence, most analytical studies have focused on synthetic
polysaccharides<sup>79,80</sup> and on capsular polysaccharides purified
from cell cultures,<sup>20</sup> based on the hypothesis that they reflect
the native capsules, although this hypothesis has been
conversely discussed.<sup>15,81,82</sup>

Here, we demonstrate that dipolar-coupling-based experi-
ments specifically and uniquely reveal polysaccharides
immobilized on the cell wall of C. neoformans, including α-
1,3-glucan, β-1,3-glucan, and chitin/chitosan. It is interesting
to note that CP-based experiments, sensitive to rigid and
immobilized molecules, fail to detect any Manp, Glc*pA*, and
Xylp signals, suggesting that there is no detectable fraction of
immobilized GXM/GXMGal in the capsule, despite their high
molecular weights. Because C. neoformans possesses the ability
to modulate its capsule thickness upon external stimuli,<sup>20,83</sup> the
most surface-exposed component (i.e., the capsule) should
exhibit elastic properties to rapidly evolve and respond to the
external environment.<sup>82,84</sup> Frases and co-workers reported a
"micro-gel" ultrastructure for the capsule by microscopy
analysis,<sup>85</sup> characterized by highly branched polysaccharide
fibers, heterogeneous in length and width. Here, we
hypothesize that the heterogeneous and porous nature of the
capsule layer is directly a result of the high mobility of its
polysaccharide constituents because the observation of GXM
and GXMGal in the J-coupling-based MAS experiments
indicates a µs mobility regime. Putative GXM-cell wall
anchoring should have led to a fraction of rigid GXM
molecules, being well anchored to the cell wall. Because no
rigid and immobilized GXM could be detected in our
experiments, we hypothesize that GXM-cell wall anchoring, if
it exists, might only concern a minor fraction (<5%) of GXM.

We demonstrated that NMR pulse sequences utilizing J-
coupling-based transfer detect capsular polysaccharides to their
counterparts in the cell wall. The use of proton detection was
crucial to disambiguate polysaccharides with close ¹³C
chemical shifts, such as Glc*pA* and Xylp. The combined use
of ¹H and ¹³C signals enabled the identification of O-2
branching from Manp to position 1 to Xylp. Our data are in
line with the serotype used in this study (i.e., H99 isolate and
serotype A), which is composed of GXM to which Manp is
substituted at position O-2 by Xylp, while, e.g., serotype B
contains a mixture of Xylp-(1 → 2)-Manp and Xylp-(1 → 4)-
Manp. Furthermore, the different branching of Manp led to
several conformations; here, for serotype A to ManG/ManX
with O-2 branching, we found that Xylp-substituted ManX can
be distinguished from Glc*pA*-substituted ManG based on its
$C_2-H_1$ cross-peak correlation. The ability of MAS NMR to rapidly distinguish the molecular nature of branching between Manp and Xylp could be advantageous to further characterize clinical isolates, as they often exhibit strong GXM structural polymorphisms. In addition to polysaccharides from the GXM component, we detected two main pyranose-based structures from GXMGal. GXM and GXMGal represent the most structurally complex polysaccharides present in fungi.⁷⁸ A key aspect is their structural polymorphism, with the nature of the branching,²¹ 6-O-acetylation,¹⁶,⁵⁶ and the relative amounts of each of the units²¹,⁵³,⁶³ that depend on the *Cryptococcus* strain. Although most of the analytical studies have focused on GXM/GXMGal released in the culture medium, our MAS NMR study provides a relevant tool to access the structural features of near-native polysaccharides in GXM and GXMGal.

In conclusion, this approach does not require time-
consuming and cell wall/capsule-damaging chemical treat-
ments and offers a promising route to study the impact of
antifungal molecules on the cell wall and capsule rearrange-
ment of C. neoformans. MAS NMR, in combination with
immunological methods and chemical analysis, may strengthen
the arsenal of analytical methods to answer key questions
related to fungal pathogenicity, such as drug susceptibility and
therapeutic drug monitoring. Importantly, the technique and
strategy can be transferred to encapsulated pathogenic bacteria,
including *Haemophilus influenzae*, *Neisseria meningitidis*, and
*Streptococcus pneumoniae*, in evaluating the organization of their
cell envelopes, as MAS NMR has already demonstrated its
ability to tackle bacterial peptidoglycans.⁸⁶,⁸⁷

■ MATERIALS AND METHODS

**Preparation of <sup>13</sup>C/<sup>15</sup>N Uniformly Labeled C. neoformans Yeast Cells.** Yeast-extract-peptone-dextrose (YPD) agar plates were streaked with frozen stocks (−80 °C) of C. neoformans H99 (serotype A) and incubated at 30 °C until the growth of yeast colonies. Then, a single colony was transferred to <sup>13</sup>C/<sup>15</sup>N-labeled liquid minimal medium (containing 1% <sup>13</sup>C-glucose and 0.1 M <sup>15</sup>N-ammonium chloride, sterile) and incubated at 30 °C in a shaking incubator maintained at 150 rpm. The yeast culture was arrested at its exponential growth phase (20 h after inoculating a single colony of C. neoformans into minimal medium), centrifuged to harvest the yeasts (1600g, 10 min), washed twice with sterile water, fixed with 2.5% para-formaldehyde (PFA) in phosphate-buffered saline (pH = 7.2) at ambient temperature for 1 h and then overnight at 4 °C, quenched with 0.1 M ammonium chloride, and centrifuged (1600g, 10 min; 3×), followed by washing with water (2×) and suspending in water; these fixed C. neoformans yeasts were stored at 4 °C until further use.

**Fluorescence Microscopy.** PFA-fixed *C. neoformans* yeasts (10⁶) were incubated with Calcofluor white (CFW; 50 µL; 10 µg/mL, aqueous) at ambient temperature in the dark for 10 min and centrifuged (1600g, 10 min), and pelleted yeasts were washed with water (three times). CFW-stained yeasts were then sequentially incubated with mouse monoclonal antibody E1 (generated against *C. neoformans* capsular polysaccharide; 50 µL at 5 µg/mL in phosphate-buffered saline)⁸⁸ and antimouse IgG-FITC (Sigma) at ambient temperature for 1 h, with centrifugation and washing (water, three times) of the yeasts between each incubation step. Further, these CFW-stained and immunolabeled *C. neoformans* yeasts were subjected to fluorescence microscopy (EVOS FL epifluorescence microscope (Life Technology)) with a 1.3-megapixel charge-coupled device (CCD) camera (Sony ICX455 monochrome) to capture the images.

**Solid-State NMR.** The sample was packed into a 0.7 mm Bruker rotor using a 1.3 mm Giotto biotech filling tool by ultracentrifugation at 25,000 rpm in a Beckman ultracentrifuge with a SW 32 Ti Swinging-Bucket Rotor for 18 h at 4 °C. The same material was also packed into a 4 mm rotor. The detailed experimental NMR

parameters are shown in Tables S1 and S2. All spectra for the
chemical shift assignments were acquired using Bruker 14.1 or 18.8 T
magnets. For all ¹H-detected spectra, the MISSISIPPI water
suppression scheme was applied. The sample temperature was ~20
°C, as measured by referencing sodium trimethylsilylpropanesulfonate
(DSS). The data were processed using the Bruker TopSpin program,
version 3.6.1 or 4.0, and analyzed using CcpNmr 2.4.2 software. The
relative abundance of the cell wall polysaccharide was determined by
extracting the cross-peak intensities from the 2D ¹³C–¹³C PDSD
spectrum.

MAS NMR relaxation measurements were acquired on a 21.1 T magnet at 105 kHz MAS. The detailed NMR parameters for the relaxation experiments are provided in Table S3. The site-specific ¹³C R₁ rates were measured using the saturation recovery method on C₁ (α-1,3-glucan and β-1,3-glucan) and C₂ positions (chitin), and H₁ (α-1,3-glucan and β-1,3-glucan) and H₃ positions (chitin). ¹H and ¹³C R₁ρ rates were obtained with a 13 kHz spin-lock power on the ¹H or ¹³C channels. The data were analyzed using Bruker DynamicsCenter 2.6.3. For CP and INEPT-based ¹H–¹³C REDOR measurements, 83.3 kHz (90°) ¹H and 50 kHz (180°) ¹³C pulses were applied with sampling every rotor period. The curves were fitted using Python scripts written in-house.

■ ASSOCIATED CONTENT

⊕ Supporting Information

The Supporting Information is available free of charge at
https://pubs.acs.org/doi/10.1021/jacs.4c16975.

NMR pulse sequences; ¹D ¹³C traces; 2D hNH and
hnCH experiments; REDOR data; and NMR acquisition
parameters (PDF)

■ AUTHOR INFORMATION

Corresponding Authors

Vishukumar Amanianda – Institut Pasteur, Unité Mycologie Moléculaire, Université Paris Cité, F-75015 Paris, France;
Present Address: Department of Biochemistry, Kasturba Medical College Manipal, Manipal Academy of Higher Education (MAHE), Manipal 576104, Karnataka, India;
Email: vaimanianda@gmail.com

Antoine Loquet – Univ. Bordeaux, CNRS, Bordeaux INP, CBMN, UMR 5248, IECB, F-33607 Pessac, France; Inserm, IECB, UAR3033, US01, Université de Bordeaux, CNRS, F-33607 Pessac, France; orcid.org/0000-0001-7176-7813;
Email: a.loquet@iecb.u-bordeaux.fr

Authors

Alons Lends – Univ. Bordeaux, CNRS, Bordeaux INP, CBMN, UMR 5248, IECB, F-33607 Pessac, France; Present Address: Latvian Institute of Organic Synthesis, Aizkraukles 21, LV-1006 Riga, Latvia

Gaelle Lamon – Univ. Bordeaux, CNRS, Bordeaux INP, CBMN, UMR 5248, IECB, F-33607 Pessac, France

Loic Delcourte – Univ. Bordeaux, CNRS, Bordeaux INP, CBMN, UMR 5248, IECB, F-33607 Pessac, France

Aude Sturny-Leclere – Institut Pasteur, Unité Mycologie Moléculaire, Université Paris Cité, F-75015 Paris, France

Axelle Grélard – Univ. Bordeaux, CNRS, Bordeaux INP, CBMN, UMR 5248, IECB, F-33607 Pessac, France

Estelle Morvan – Inserm, IECB, UAR3033, US01, Université de Bordeaux, CNRS, F-33607 Pessac, France

Muhammed Bilal Abdul-Shukkoor – Univ. Bordeaux, CNRS, Bordeaux INP, CBMN, UMR 5248, IECB, F-33607 Pessac, France

Mélanie Berbon – Univ. Bordeaux, CNRS, Bordeaux INP, CBMN, UMR 5248, IECB, F-33607 Pessac, France
Alicia Vallet – *Institut de Biologie Structurale, CEA, Université Grenoble Alpes, CNRS, F-38000 Grenoble, France*

Birgit Habenstein – *Univ. Bordeaux, CNRS, Bordeaux INP, CBMN, UMR 5248, IECB, F-33607 Pessac, France*;
orcid.org/0000-0003-3723-9552

Erick J. Dufourc – *Univ. Bordeaux, CNRS, Bordeaux INP, CBMN, UMR 5248, IECB, F-33607 Pessac, France*;
orcid.org/0000-0002-8221-9326

Paul Schanda – *Institute of Science and Technology Austria, Klosterneuburg 3400, Austria*; orcid.org/0000-0002-9350-7606

Complete contact information is available at:
https://pubs.acs.org/10.1021/jacs.4c16975

## Notes

The authors declare no competing financial interest.

## ACKNOWLEDGMENTS

We thank the ANR (ANR-16-CE11-0020-02 to A. Loquet, and V.A. and ANR-21-CE17-0032-01 grant FUNPOLYVAC to V.A.) as well as the Swiss National Science Foundation for early postdoc mobility project P2EZP2_184258 to A. Lends. This work has benefited from the Biophysical and Structural Chemistry Platform at Institut Européen de Chimie et Biologie IECB, Centre National de la Recherche Scientifique CNRS Unité d'Appui et de Recherche UAR 3033, INSERM US001, and CNRS (IR-RMN FR3050 and Infr analytics FR2054).

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