Polymorphic α-Glucans as Structural Scaffolds in Cryptococcus Cell Walls for Chitin, Capsule, and Melanin: Insights from13C and1H Solid-State NMR

Polymorphic α-Glucans as Structural Scaffolds in Cryptococcus Cell Walls for Chitin, Capsule, and Melanin: Insights from 13C and 1H Solid-State NMR | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Polymorphic α-Glucans as Structural Scaffolds in Cryptococcus Cell Walls for Chitin, Capsule, and Melanin: Insights from 13 C and 1 H Solid-State NMR Ankur Ankur , View ORCID Profile Jayasubba Reddy Yarava , View ORCID Profile Isha Gautam , View ORCID Profile Faith J. Scott , View ORCID Profile Frederic Mentink-Vigier , Christine Chrissian , Li Xie , Dibakar Roy , View ORCID Profile Ruth E. Stark , Tamara L. Doering , Ping Wang , View ORCID Profile Tuo Wang doi: https://doi.org/10.1101/2025.04.18.649559 Ankur Ankur 1 Department of Chemistry, Michigan State University , East Lansing, MI, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Jayasubba Reddy Yarava 1 Department of Chemistry, Michigan State University , East Lansing, MI, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Jayasubba Reddy Yarava Isha Gautam 1 Department of Chemistry, Michigan State University , East Lansing, MI, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Isha Gautam Faith J. Scott 2 National High Magnetic Field Laboratory, Florida State University , Tallahassee, FL, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Faith J. Scott Frederic Mentink-Vigier 2 National High Magnetic Field Laboratory, Florida State University , Tallahassee, FL, USA 3 Department of Chemistry and Biochemistry, Florida State University , Tallahassee, FL, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Frederic Mentink-Vigier Christine Chrissian 4 Department of Chemistry and Biochemistry, City College of New York , New York, NY, USA 5 CUNY Institute for Macromolecular Assemblies, The City University of New York , New York, NY, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Li Xie 1 Department of Chemistry, Michigan State University , East Lansing, MI, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Dibakar Roy 1 Department of Chemistry, Michigan State University , East Lansing, MI, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Ruth E. Stark 4 Department of Chemistry and Biochemistry, City College of New York , New York, NY, USA 5 CUNY Institute for Macromolecular Assemblies, The City University of New York , New York, NY, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Ruth E. Stark Tamara L. Doering 6 Department of Molecular Microbiology, Washington University in St. Louis School of Medicine , St. Louis, MO, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Ping Wang 7 Departments of Microbiology, Immunology & Parasitology, Louisiana State University Health Sciences Center , New Orleans, LA, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Tuo Wang 1 Department of Chemistry, Michigan State University , East Lansing, MI, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Tuo Wang For correspondence: wangtuo1{at}msu.edu Abstract Full Text Info/History Metrics Supplementary material Preview PDF ABSTRACT Cryptococcus species are major fungal pathogens responsible for life-threatening infections in approximately a million individuals globally each year, with alarmingly high mortality rates. These fungi are distinguished by a distinctive cell wall architecture further reinforced by two virulence-associated layers, melanin and capsule, rendering them insensitive to antifungal agents targeting the cell wall, such as echinocandins. The molecular interplay between these three biomolecular layers remains poorly understood. Here we employ solid-state NMR spectroscopy to examine intact cells of both wild-type and capsule-deficient strains of C. neoformans , along with its melanized cells. High-resolution 13 C and 1 H data revealed five distinct structural forms of α-1,3-glucans that play versatile roles in forming the rigid cell wall scaffold by interacting with chitin microfibrils and chitosan, and in stabilizing the mobile matrix by associating with β-1,6-glucan and a small fraction of β-1,3-glucan. Two primary forms of α-1,3-glucans were distributed throughout the cell wall, hosting melanin deposition in the inner domain and capsule attachment on the cell surface. These findings offer a paradigm shift in understanding the cryptococcal cell wall and its interaction with two key virulence factors on opposite sides, raising critical biochemical questions that could inform the development of more effective antifungal treatments for cryptococcosis. INTRODUCTION Cryptococcus species are a group of encapsulated basidiomycetous fungi causing life-threatening infections in immunocompromised and immunocompetent individuals 1 . These fungi produce characteristic virulence factors, including the antiphagocytic polysaccharide capsule, antioxidant melanin pigment, and extracellular proteases, including ureases and phospholipases 2 , 3 . Each year, approximately 200,000 cases of cryptococcal meningitis are reported worldwide, with mortality rates of 20-70% that vary significantly depending on disease severity and patient health status 4 , 5 . Untreated diseases are uniformly fatal. Cryptococcus neoformans and Cryptococcus gattii are the primary species responsible for these infections, with C. neoformans accounting for 90% of clinical isolates 6 , 7 . Pulmonary infections are established upon inhalation of spores or desiccated yeast cells 8 . In immunocompromised individuals, particularly when associated with HIV/AIDS, the infection can disseminate, enabling the pathogen to traverse the blood-brain barrier and induce meningoencephalitis—an inflammatory condition affecting the brain and its surrounding tissues 9 , 10 . The standard treatment regimen for cryptococcosis involves amphotericin B, flucytosine, and fluconazole, typically administered for 6 to 12 months; however, this prolonged therapy is both financially burdensome and associated with significant adverse effects 11 , 12 . Recent antifungal research has focused on targeting fungal cell wall biosynthesis, leading to the successful development of echinocandins, which inhibit β-1,3-glucan synthesis—an essential structural component of most fungal cell walls 13 . However, echinocandins exhibit limited efficacy against Cryptococcus species, with the underlying mechanisms for their reduced activity remaining poorly understood but likely attributed to the unique cell wall architecture of Cryptococcus species 14 , 15 . The cryptococcal cell wall is a dynamic, multilayered structure essential for virulence, immune evasion, and stress resistance 8 , 16 . Decades of chemical and imaging analyses have led to the proposition of a two-layered model of cryptococcal cell walls, in which the inner layer comprises an alkali-insoluble meshwork of β-glucans and chitin/chitosan, and the outer layer consists of an alkali-soluble fraction containing α- and β-glucans 16 - 18 . Unlike in other yeasts, where β-1,3-glucan is predominant, β-glucans in the Cryptococcus cell wall are primarily β-1,6-linked, forming covalent linkages with β-1,3-glucan, chitin, and cell wall proteins, while β-1,3-glucan is less abundant 15 , 19 . The cryptococcal cell wall is further associated with two additional layers of biomacromolecules that both serve as key virulence factors: a capsular layer coating the surface and melanin deposited between the cell wall polysaccharides and the plasma membrane 20 - 23 . The cryptococcal capsule, which provides structural integrity and mediates host-pathogen interactions, is primarily composed of glucuronoxylomannan (GXM), with minor fractions of glucuronoxylomannogalactan (GXMGal) and mannoproteins 24 - 26 . These capsular polysaccharides are found both attached to the cell wall and released as exopolysaccharides 27 . Under nutrient-deficient conditions, Cryptococcus polymerizes external polyphenolic compounds, leading to melanin deposition in its cell wall 28 , 29 . These melanin granules form layered structures that protect fungi against oxidative stress, contribute to virulence, and allow small molecules, such as sugars and amino acids, to pass while restricting access by larger antifungal compounds 30 , 31 . Current knowledge of capsular carbohydrates is based mainly on exopolysaccharides isolated from culture supernatants, rather than those associated with the cell wall, while studies on melanin have focused on extracts called melanin ghosts, making it challenging to understand the interaction patterns of these complex and heterogeneous polymers within the cell wall 27 , 32 . Recently, solid-state NMR spectroscopy has been introduced to enable high-resolution structural analysis of polysaccharides within intact fungal cells, thereby eliminating the need for cell disruption or fractionation and allowing the investigation of native physiological architectures and interactions without perturbation. Here, we use 13 C and 1 H solid-state NMR, enhanced by the sensitivity-boosting Dynamic Nuclear Polarization (DNP) technique 33 - 36 , to explore the structural polymorphism and assembly of carbohydrate polymers in intact cells of wild-type and acapsular C. neoformans strains. Our detailed examination of the cell wall reveals that β-1,6-glucan is the predominant component of the mobile matrix, accompanied by smaller amounts of β-1,3-glucans and mannoproteins, and is also present—albeit less abundantly—in the rigid phase alongside chitin, chitosan, and α-1,3-glucans. α-1,3-glucans dominate the rigid core and exhibit five distinct structural forms, each contributing uniquely to mechanical reinforcement, mobile matrix integration, capsular polysaccharide anchoring, and melanin deposition. These novel structural insights elucidate the organization of C. neoformans cell walls, provide a molecular perspective on their interface with melanin and the capsule, and underscore the critical and diverse structural roles of α-glucans, highlighting their potential as novel therapeutic targets. RESULTS Predominance of α-1,3-glucans in the rigid core of cell wall in Cryptococcus Cryptococcus species were initially classified into subtypes based on capsular polysaccharide antigenicity, but are now distinguished by DNA sequencing, ecological characteristics, and pathobiological evidence 37 - 40 . In this study, we selected several representative strains of the dominant pathogen C. neoformans for examination. To enable studies of the capsule, we compared a clinical wild-type strain H99 with an acapsular pka1 mutant, both of serotype A and in the same background, as well as an environmental strain JEC20 with a hypocapsular Cap70 mutant, both of serotype D (also termed C. deneoformans ) and in the same background 41 , 42 . These mutants either completely lack or significantly suppress capsule formation on the cell surface, as confirmed by SEM imaging ( Fig. 1a ). Correspondingly, the mean of total cell diameter decreased from 5.7 µm in H99 cells to 4.6 µm in pka1 cells and from 5.9 µm in JEC20 cells to 4.2 µm in Cap70 cells ( Fig. 1b and Supplementary Table 1 ). Download figure Open in new tab Figure 1. Polysaccharide composition varies in wildtype strains and capsule mutants of C. neoformans . ( a ) SEM images of C. neoformans serotype A strains (wildtype H99 and acapsular pka1 ) and serotype D (wildtype JEC20 and hypocapsular Cap70). ( b ) Total cell diameter measured from SEM images. Boxes represent the interquartile range (IQR), with whiskers extending to 1.5 times the IQR. Open circles: average; horizontal lines: median. Sample sizes: H99 and pka1 (n = 24), JEC20 (n = 19), and Cap70 (n = 32). ( c ) 2D 13 C- 13 C CORD correlation spectra of four C. neoformans samples. The absence of α-1,2,3-mannose and xylose signals in the acapsular pka1 mutant is highlighted with dashed brown and magenta squares, respectively, with intensity reductions observed in the hypocapsular Cap70 mutant. Observed carbohydrates include α-1,3-glucan (A), chitin (Ch), chitosan (Cs), β-1,6-glucan (H), α-1,2,3-mannose (Mn), and xylose (X). Three subtypes of α-1,3-glucan (A a , A b , and A c ) are indicated by superscripts. ( d ) Simplified structural representations of key polysaccharides, with NMR abbreviations and key carbon sites labeled. ( e ) Molar composition of rigid polysaccharides estimated from resolved cross-peak volumes in 2D 13 C CORD spectra. ( f, g ) Overlay of refocused J-INADEQUATE spectra obtained via direct polarization (DP) for mobile molecules (cyan) and cross polarization (CP) for rigid molecules (purple) reveals ( f ) β-1,6-glucan dominance in the mobile fraction and ( g ) preferential localization of A a and A b in the rigid fraction, while A c is distributed across both rigid and mobile phases. ( h ) Molar composition of mobile polysaccharides estimated from well-resolved peaks in DP refocused J-INADEQUATE spectra. Rigid polysaccharides were analyzed using a 2D 13 C- 13 C CORD correlation experiment 43 , revealing strong signals for α-1,3-glucans, along with weaker signals for chitin, chitosan, and β-1,6-glucan ( Fig. 1c, d ; Supplementary Fig. 1 and Table 2 ). Intensity analysis indicated that α-1,3-glucans constitute 66-84% of the rigid polysaccharides across all four strains, while chitin and chitosan together account for 6-13% ( Fig. 1e and Supplementary Table 3 ). These findings suggest that α-1,3-glucan plays a critical and unanticipated role in maintaining the mechanical scaffold of the cell wall, likely by associating with the microfibrils formed by chitin and chitosan. Unexpectedly, signals corresponding to capsular components, including the α-1,2,3-linked mannose residues along the glucuronoxylomannan backbone and its xylose branches, were also detected within the rigid polysaccharides of H99 and JEC20 cells ( Fig. 1c and Supplementary Fig. 2 ). These signals were entirely absent in the acapsular pka1 mutant. In the hypocapsular Cap70 mutant, xylose signals were depleted, while α-1,2,3-linked mannose residues were retained but at reduced intensity. Previous studies have identified capsule molecules in the mobile phase of C. neoformans cells cultured in capsule-inducing media 44 , and our observations show that capsular polysaccharides could be structurally rigidified through interactions with cell wall polysaccharides. Identification of three structurally and dynamically distinct forms of α-1,3-glucans Three magnetically distinct subtypes of α-1,3-glucans—designated as types a, b, and c—were unequivocally identified within the rigid fraction, exhibiting a sequentially decreasing prevalence within each sample ( Fig. 1c and Supplementary Fig. 3 ). Type-a represents the most prevalent allomorphic form, comprising 34-41% of the rigid carbohydrate fraction across all samples ( Fig. 1e ). Type-b accounts for 20-36% of the rigid fraction, while type-c extends from the rigid phase into the mobile phase, constituting 6-11% of the rigid fraction ( Fig. 1e ) and 3-7% of the mobile fraction, as demonstrated later. In Cap70 cells, type-b α-1,3-glucan was remarkably enriched, reaching levels comparable to type-a in this mutant, suggesting a possible compensatory adaptation to the hypocapsular state. The three spectroscopically distinguishable forms of α-1,3-glucans arise from local structural perturbations, including conformational variations and differential molecular interactions with neighboring components. Type-a and type-b exhibited identical 13 C chemical shifts at C1 and C3 (101.4-101.6 ppm and 85.0-85.5 ppm, respectively), the carbon sites involved in glycosidic linkages between adjacent sugar units along the polymer chain, indicating that they share the same helical screw conformation. However, these two forms are best distinguished by their C4, C5, and C6 chemical shifts, which reflect differences in hydroxymethyl conformation related to the exocyclic -CH 2 OH group. In contrast, type-c exhibited significantly lower 13 C chemical shifts at C1 (100.4 ppm) and C3 (81.9 ppm), indicating an entirely restructured helical screw conformation compared to the other forms. These α-1,3-glucan allomorphic forms also have distinct dynamic properties: while all three were identified within the rigid fraction, a small amount of type-c was also detected in the mobile fraction ( Fig. 1f and Supplementary Figs. 3, 4 ). This specific form of α-glucan may serve as a transitional component between the rigid α-glucan and chitin domains and the dynamic matrix primarily composed of β-1,6-glucan (56-77% of the mobile fraction; Fig. 1g, h ), β-1,3-glucan (15-21%), and some α-1,2-linked mannose residues, likely derived from mannoproteins ( Fig. 1h and Supplementary Table 4 ). This finding provides evidence for the critical role of α-1,3-glucan in mediating physical interactions that bridge rigid and dynamic domains. In the mobile phase, we observed abundant β-1,6-glucan, which is expected as it is unusually rich in Cryptococcus compared to other yeasts such as Saccharomyces cerevisiae 15 , 19 . The structure of cryptococcal β-1,6-glucan consists of short chains, often branched with β-1,3-glucan, and our data further identified it as the dominant component of the mobile matrix. The mobile phase also contains proteins and lipids; however, their widespread distribution throughout the cell precluded a focused analysis of their specific contributions to cell wall structure ( Supplementary Fig. 5 ). Rigid α-glucan forms interact with capsules to form dehydrated domains on cell surface Hydration profiles of rigid biopolymers were analyzed using water 1 H polarization transfer to carbohydrates via water-editing experiments, where the S/S 0 intensity ratios between water-edited (S) and control (S 0 ) spectra reflect the extent of water retention at specific carbon sites ( Fig. 2a and Supplementary Fig. 6 ) 45 - 47 . Two α-1,3-glucan subtypes, a and b, were the least hydrated molecules in Cryptococcus cell walls, with average S/S 0 values of 0.26-0.43 across all four strains ( Fig. 2a and Supplementary Table 5 ). Meanwhile, in the examination of the molecular motions of rigid polysaccharides through NMR relaxation measurements, the resolvable carbon sites of type-a α-1,3-glucan exhibited the longest 13 C-T 1 and 1 H-T 1ρ relaxation time constants across the four strains, with average values of 2.7-3.9 s ( Fig. 2b ; Supplementary Fig. 7 and Table 6 ) and 9.7-12.6 ms ( Fig. 2c and Supplementary Fig. 8 ), respectively. These values are significantly higher than those of the chitin and β-1,6-glucans in the rigid phase, indicating that α-1,3-glucans experience highly restricted molecular motion on both nanosecond and microsecond timescales. The combination of restricted dynamics and limited hydration suggests that types-a and b α-1,3-glucans aggregate into large complexes that effectively exclude bulk water. In contrast, type-c α-1,3-glucan was not only more hydrated but also more heterogeneously hydrated compared to other subforms ( Fig. 2a ). This further supports the notion that type-c α-1,3-glucan plays a crucial role in bridging the rigid and mobile phases of the α-1,3-glucan matrix ( Fig. 1e, h ). Download figure Open in new tab Figure 2. Hydration, dynamics, and DNP spectra reveal α-glucan interactions with capsule and chitin. (a) Intensity ratios (S/S 0 ) between water-edited peak spectra (S) and control spectra (S 0 ) showing the extent of water association for α-1,3-glucan (A), chitin (Ch), 1,2,3-linked mannan (Mn), and β-1,6-glucan (H), with the superscripts indicating the respective subtypes. Open circles: average; horizontal lines: median. Data size: A a (n=16) in all four samples; A b , A c , Ch, H (n=9 each) in all; Mn (n=9) in H99 and JEC20 only. (b) 13 C-T 1 relaxation time constants of rigid cell wall polysaccharides. ( c ) 1 H-T 1ρ relaxation time constants of rigid polysaccharides. For both panels b and c, error bars are s.d. of the fit parameters and horizontal lines represent the average. ( d ) DNP-enhanced 2D 13 C- 13 C DARR spectrum of H99 cells. A new, minor form of α-1,3-glucan (A d ) is resolved. Intermolecular cross peaks were observed between chitin and type-a and type-d α-1,3-glucans, with dashed lines to mark the intramolecular and intermolecular cross peaks of these two α-1,3-glucan subtypes. ( e ) DNP-enhanced 2D 13 C- 13 C PAR spectrum with 15 ms recoupling time. Top: observation of a cross peak between chitin and A d . Bottom: the dashed line indicates the diagonal of the C3 sites, with off-diagonal intensities showing intermolecular cross peaks. β-1,6-glucans are the best-hydrated rigid polysaccharide in C. neoformans , with high S/S 0 averages of 0.53-0.77 ( Fig. 2a ). β-1,6-glucan also exhibited short 13 C-T 1 (0.9-1.5 s) and 1 H-T 1ρ (4.4-6.8 ms) relaxation times— about half to two-thirds those of type-a α-1,3-glucans ( Fig. 2b, c ). β-1,6-glucan is the most dynamic molecule, maintaining substantial hydration even within the rigid core. Chitin displays intermediate rigidity and hydration profiles, typically between β-1,6-glucan and type-a α-1,3-glucan. Chitin forms a microfibrillar structure primarily through antiparallel chain packing but is not the most rigid molecule in the cell wall. This may be due to a high level of deacetylation to chitosan, estimated to be 30-40% based on molar composition ( Fig. 1e ), which confers dynamics on these microfibrils. Unexpectedly, 1,3-linked mannose residues, which compose the backbone of GXM, exhibited low S/S 0 values, averaging 0.38 and 0.41 in H99 and JEC20 cells, respectively. This indicates significant dehydration of capsular polysaccharides in both wild-type serotypes A and D strains ( Fig. 2a ), which is counterintuitive because capsular GXM is surface-exposed and would therefore be expected to be well-hydrated. A plausible explanation is that GXM associates with α-1,3-glucan subtypes a and b, which not only rigidifies GXM but also forms a dehydrated layer on the cell surface. This structural organization contrasts with cryptococcal capsules produced in capsule-inducing media, where GXM is loosely associated with cell wall molecules, thus remaining mobile and solubilized 44 . Interestingly, the two serotypes responded differently to the absence or reduction of capsular polysaccharides. In serotype A, water association decreased substantially in the pka1 mutant compared to wild-type H99 cells ( Fig. 2a ), with average S/S 0 ratios dropping from 0.42 to 0.27 for type-a α-glucan, 0.58 to 0.33 for type-c α-glucan, 0.77 to 0.56 for β-1,6-glucan, and 0.75 to 0.43 for chitin. In contrast, serotype D exhibited increased hydration for most cell wall polysaccharides, with S/S 0 values rising from 0.26 in JEC20 to 0.37 in Cap70 cells for type-a α-glucan, 0.45 to 0.59 for type-c α-glucan, and 0.49 to 0.67 for chitin, except for β-1,6-glucan, whose hydration level decreased. Therefore, different serotypes may exhibit variations in cell wall organization and adopt distinct compensatory mechanisms to remodel their polysaccharides in response to capsule depletion. However, a consistent observation is that chitin dynamics became more confined in both acapsular and hypocapsular mutants ( Fig. 2b, c ), suggesting strengthened interactions with α-glucans, likely due to increased availability of interaction sites on types-a and b α-glucans after GXM removal. Two specific α-glucans interact with chitin to form partially ordered domains in the cell wall We used DNP to enhance the NMR sensitivity for polysaccharides in H99 cells by 11-fold through polarization transfer from electrons in the biradical AsymPol-POK to the 1 H and then 13 C nuclei in these cellular biomolecules 33 , 48 . The observed signals arose from partially ordered molecules, including chitin, chitosan, types-a and b α-glucans, and a newly identified minor form, type-d α-1,3-glucan (labeled as A d in Fig. 2d ). This new form of α-1,3-glucan exhibited unique C3 and C4 chemical shifts at 87.6 ppm and 67.8 ppm, with weak signals detectable only through DNP enhancement. Meanwhile, signals from mobile and semi-mobile molecules, such as β-1,6-glucan and type-c α-1,3-glucan, were broadened out due to a diverse ensemble of conformations trapped at the cryogenic temperature. With the enhanced sensitivity, strong intermolecular interactions were detected between chitin and types-a and type-d α-1,3-glucans, as shown by the unambiguous Ch2-A a 3, Ch4-A a 3, A a 3-Ch4, Ch4-A d 3, and A d 3-Ch4 cross-peaks observed in the 100-ms dipolar-assisted rotational resonance (DARR) spectrum ( Fig. 2d ). We also observed three cross peaks in a 15-ms proton-assisted recoupling (PAR) spectrum, including Ch2-A d 1, Ch4-A a 3, and A a 3-Ch4, further confirming the interactions between chitin and these two specific forms of α-1,3-glucans ( Fig. 2e ) 49 , 50 . In addition, a cross peak between the carbon-3 sites of type-a and type-d α-1,3-glucans (A a 3-A d 3) was detected ( Fig. 2e ). Together, these observations confirm that type-d and type-a α-1,3-glucans are associated with each other and further packed with chitin in the rigid domain of the cell wall, while type-b and type-c α-1,3-glucans are not colocalized with chitin microfibrils. Two structural forms of α-1,3-glucans host melanin deposition In addition to the capsule, another crucial virulence factor of Cryptococcus species is their ability to synthesize melanin, which is deposited in the cell wall, and forms a protective barrier against environmental and host stressors 31 . Melanin polymers are proposed to be composed of stacked aromatic and indolic rings, formed via the oxidative polymerization of L-3,4-dihydroxyphenylalanine (L-DOPA) catalyzed by the laccase enzyme ( Fig. 3a, b ) 51 , 52 . Since Cryptococcus requires an exogenous substrate for melanin biosynthesis, we supplemented the minimal medium with 1 mM ring- 13 C 6 -labeled L-DOPA to cultivate melanin-rich C. neoformans H99 cells 53 - 55 . Consequently, the melanin-labeled cells exhibited enhanced aromatic 13 C signals in the 110-160 ppm range, particularly in the 140-150 ppm region, corresponding to non-protonated melanin carbon sites ( Fig. 3c , see expanded spectral region). These distinct melanin aromatic signals enabled us to establish correlations with carbohydrate proton resonances to map out melanin-carbohydrate interactions within intact cells. Download figure Open in new tab Figure 3. 1 H-detected solid-state NMR shows interactions of α-1,3-glucan with melanin. ( a ) Structure of melanin precursor ring- 13 C 6 -labeled L-DOPA. Asterisks indicate the 13 C-labeling sites. ( b ) Representative structure of DHI melanin formed by covalently linked subunits. ( c ) 1D 13 C CP spectra of non-melanized (orange) and melanized (black) C. neoformans H99 cells. The spectra are normalized by the α-1,3-glucan carbon 1 peak at 101 ppm (asterisk). The zoom-in region shows the aromatic signals from melanin, with intensities magnified by 8-fold. ( d ) Polymorphic forms of α-1,3-glucans (A a , A b , and A e ) resolved using 1 H-detected 3D hCCH TOCSY (WALTZ-16) spectrum. 1 H chemical shifts (ppm) are in grey. Key strips are extracted for ω 2 -ω 3 ( 13 C- 1 H) and ω 1 -ω 3 ( 13 C- 1 H) planes. ( e ) 2D hChH with 0.8 ms RFDR mixing showing intermolecular interaction between melanin non-protonated carbons and α-1,3-glucan protons. ( f ) Structural summary of observed interactions between cell wall α-1,3-glucan and melanin units. Carbon and nitrogen atom numbering in melanin fragments follows the standard convention for indoles. Blue solid lines: cross peaks observed between non-protonated carbons and protons in protonated sites within melanin. Dashed lines in purple and orange indicate the intermolecular interactions between melanin carbons and protons in type-a and type-b α-1,3-glucans, respectively. The experiments were performed on 600 MHz NMR at 60 kHz MAS. We further identified three polymorphic forms of α-1,3-glucan in melanized C. neoformans cells, designated A a , A b , and A e , from the 13 C- 1 H strips extracted from the 3D hCCH TOCSY (WALTZ-16) spectrum ( Fig. 3d and Supplementary Table 7 ). These polymorphs exhibited distinct 1 H chemical shifts: the anomeric carbon (A1) correlated with three distinct 1 H resonances at 5.5 ppm (A a ), 4.8 ppm (A b ), and 6.1 ppm (A e ) in the ω 2 -ω 3 13 C- 1 H strip, while the C3 position correlated with protons at 3.5 ppm (A a ), 3.8 ppm (A b ), and 4.4 ppm (A e ) ( Fig. 3d ). The ω 1 -ω 3 1 H- 13 C strip extracted from 70.4 ppm further revealed the through-bond carbon connectivity of these three polymorphic forms and established correlations with distinct carbon sites, such as the resolvable carbons at positions 3 and 1 ( Fig. 3d ). In this analysis, the A c and A d forms detected in 13 C-based experiments were not observed in this analysis due to their semi-dynamic nature or low abundance. Next, we measured a 2D hChH spectrum, which employed an extended, 0.8 ms radio frequency driven recoupling (RFDR) mixing period to facilitate long-range 1 H- 1 H polarization transfer ( Fig. 3e ). Notably, this experiment facilitated the observation of extensive correlations between non-protonated carbons and protons at protonated sites within the melanin structure ( Fig. 3e ; blue solid lines in Fig. 3f ). The 13 C and 1 H chemical shifts identified in melanin are summarized in Supplementary Fig. 9 and Tables 8 and 9 . In melanin subunit D, key cross-peaks include those between carbon-9 and protons-2/3 (D9-D2/3 H ), as well as D5-D7 H and D8-D7 H ( Fig. 3e, f ). Similar cross-peaks were observed within other melanin subunits, such as A7/8-A7 H , B9-B3/4 H , B5-B4 H , C4-C3 H , and C6-C7 H . The C6-N H cross-peak further revealed spatial proximity between C-fragment carbons and indole amide protons from the same or neighboring subunits. Importantly, the hChH experiment simultaneously revealed extensive intermolecular interactions between the carbons in the indole rings of melanin and the protons in carbohydrates. For example, the carbon 4 of melanin subunit-C and carbon 8 of subunit-D, two non-protonated carbons resonating at 130 ppm, exhibited correlations with the proton 1 of type-a α-1,3-glucan at 5.5 ppm, the proton 1 of type-b α-1,3-glucan at 4.8 ppm, and the proton 3 of type-b α-1,3-glucan at 3.8 ppm, resulting in the C4/D8-A a 1 H , C4/D8-A b 1 H , and C4/D8-A b 3 H cross peaks ( Fig. 3e ). Type-a and type-b α-1,3-glucans showed 13 and 10 strong cross-peaks with melanin, respectively, including A8/B5-A a 1 H , A5-A b 3 H , A5-A b 1 H , A6-A b 1 H , C5-A a 1 H , C8-A a 1 H , C6-A b 1 H , and C6-A b 3 H , in addition to those previously described ( Fig. 3e, f ). These intermolecular interactions provide the first molecular-level experimental evidence that α-1,3-glucans act as a previously unrecognized partner with melanin, with their type-a and type-b structural variants serving as the primary interactors. DISCUSSION Recent advances in solid-state NMR spectroscopy have significantly enhanced our ability to investigate fungal cell walls, offering detailed insights into the structures of wall polymers, as well as their hydration, dynamics, and intermolecular interactions 56 - 60 . Such analysis describes molecular behavior in the context of intact cells, in a way that prior methods, which relied on physical, chemical, and/or enzymatic perturbation, could not 61 , 62 . The application of these techniques to the major fungal pathogen C. neoformans in this study has uncovered unexpected features of the cell wall and revealed its interactions with two critical virulence factors: the extracellular capsule and cell-associated melanin. Previous studies of cryptococcal cell walls, based on structural and imaging analyses, suggested a bilayer structure with an inner layer composed of interlinked chitin, chitosan, and β-glucans and an outer layer composed of α- and β-glucans 16 , 17 , 63 . In this model, melanin localizes to the inner aspect of the wall, near the plasma membrane, while the capsule polysaccharide is associated with α-1,3-glucan in the outer layer 23 , 25 . The integration of our NMR data with this existing model now supports major revisions ( Fig. 4 ). For example, our data demonstrate that α-1,3-glucan is a key structural component of cryptococcal cell walls, occurring both in the inner wall, where it interacts with melanin and chitin, and in the outer wall, where it promotes capsule attachment. We also found that β-1,6-glucan, an under-investigated component in the fungal cell wall 64 , is distributed across dynamically distinct domains. It is primarily localized in the mobile phase, where it forms the main component of the soft matrix, with smaller fractions extending into the rigid phase to contribute to structural integrity. This also explains the limited efficacy of echinocandins against Cryptococcus species, as β-1,3-glucan is present in low abundance, with β-1,6- and α-glucans dominating, and β-1,3-glucan being confined exclusively to the mobile phase. Download figure Open in new tab Figure 4. Organization of cryptococcal cell walls and association with capsule and melanin. The illustrative scheme is based on NMR data from H99 cells, integrated with previously reported biochemical and imaging analyses 16 , 31 , 63 . Five distinct forms of α-1,3-glucans constitute 70% of the rigid polysaccharides. Capsular polysaccharides, primarily GXM, are associated with type-a and type-b α-1,3-glucans in the cell wall. Melanin is stabilized by type-a and type-b α-1,3-glucans. Molecules associated with type-a and type-b α-1,3-glucans are rigidified and dehydrated. Chitin and chitosan, which together account for only 6% of rigid molecules, exhibit a high deacetylation level of approximately 30% and are stabilized through interactions with type-d and type-a α-1,3-glucans, both of which colocalize. The mobile phase (highlighted by dashed lines) is predominantly composed of β-1,6-glucan (70%), which regulates the water association of cryptococcal cell wall, with a minor fraction of β-1,3-glucan, present either as linear chains or as sidechains of β-1,6-glucan. Additionally, type-c α-1,3-glucan extends through rigid and mobile phases. Type-e α-1,3-glucan represents a minor structural variant that does not exhibit an association with melanin. Molecular fractions are considered in this model, though the representation is not strictly to scale. The background gradient illustrates the water distribution within the cell wall. Notably, the cell wall α-1,3-glucan of C. neoformans occurs in five distinct morphotypes, which we were able to resolve using high-resolution 13 C solid-state NMR, combined with advanced 1 H detection and DNP sensitivity enhancement ( Fig. 4 ). Type-a and type-b α-1,3-glucans are the predominant structured forms, comprising 55-75% of all rigid polysaccharides ( Fig. 1e ). These forms aggregate into partially dehydrated bundles, facilitating capsule deposition on the cell surface and rigidifying the capsule polysaccharide GXM ( Fig. 2a-c ; upper portion of Fig. 4 ). This highlights the essential role of α-1,3-glucan in capsule-cell wall attachment, and provides a structural explanation for why disruption of the α-1,3-glucan synthase gene results in viable C. neoformans cells lacking a surface capsule, despite the successful production of capsule components 65 . Surprisingly, these major forms of α-1,3-glucan (types a and b) that interact with capsule polysaccharide also serve to host melanin deposition ( Fig. 3e ; bottom portion of Fig. 4 ). These unexpected findings lead to two novel conclusions. First, they solve the mystery of which wall component helps organize melanin deposition, which was not known despite the implication of the cell wall in this process 29 , 51 . Second, they indicate that these two forms of α-1,3-glucans exhibit a broad spatial distribution, localized both at the cell surface, where they mediate capsule association, and near the plasma membrane, where they colocalize with melanin ( Fig. 4 ). This bimodal distribution significantly shifts the structural paradigm of cryptococcal cell walls, revealing that the inner layer, previously believed to consist of chitin, chitosan and β-glucan, contains additional components 16 . This model is also consistent with the persistent melanin-cell-wall association observed even in samples with relatively low chitin and chitosan content. The remaining three forms of α-1,3-glucan, though less abundant, are also functionally significant. Type-c is present in both the rigid and mobile phases ( Fig. 1e, h ), and thus plays a critical role in integrating α-1,3-glucan aggregates into the mobile matrix. Type-d α-1,3-glucan is a minor but highly ordered form that colocalizes with type-a α-glucan, both of which are physically associated with chitin microfibrils ( Fig. 2d , e ). Type-e, another minor form, underlies the 13 C signals of type-a and type-b α-glucans and is distinguishable by 1 H chemical shifts, indicating local structural variations compared to the predominant forms ( Fig. 3d ). Our results provide an unprecedented understanding of the structural complexity of α-1,3-glucan in the construction of cryptococcal cell walls and highlight its association with two key virulence factors anchored at opposite aspects of the cell wall. The multiple roles of this versatile polymer may also explain its unusually high abundance in Cryptococcus species. Meanwhile, these findings raise new questions. It remains unclear whether the observed structural forms result from complexity in α-1,3-glucan biosynthesis and whether they are universally present across different Cryptococcus species, other serotypes, and other pathogenic fungal species. It also remains to be determined whether inhibitors of α-1,3-glucan synthases or other enzymes involved in biosynthesis and modification could be developed as therapeutic targets. Addressing these questions would advance our understanding of cryptococcal cell wall biosynthesis and structure, providing molecular insights for the development of effective antifungal therapies against cryptococcosis. METHODS Preparation of 13 C, 15 N-labeled fungal material C. neoformans strains H99, JEC20, pka1 , and Cap70 were cultured in 13 C, 15 N-enriched growth media. The growth medium consisted of 1.7% Yeast Nitrogen Base (YNB), 2.1% 13 C-glucose (Cambridge Isotope Laboratories, CLM-1396-PK), and 1.5% 15 N-ammonium sulfate (Cambridge Isotope Laboratories, NLM-713-PK), with the pH adjusted to 5.5 using 1M HEPES buffer. Cells were cultured in 100 mL liquid medium in 250-mL Erlenmeyer flasks and incubated at 37°C with shaking at 150 rpm (1364 × g). Fungal biomass was harvested by centrifugation at 4,000 rpm (13,700 × g) for 5 min at 4°C and the pellet was washed four times using nano-purified water followed by the centrifugation procedures. For solid-state NMR characterization, 35-45 mg of natively hydrated whole-cell material was packed into a 3.2-mm magic-angle spinning (MAS) rotor (Cortecnet, HZ16916). To generate melanin-rich C. neoformans cells, H99 cells was incubated for 10-14 days at 30°C with shaking at 150 rpm in a minimal medium (29.4 mM KH 2 PO 4 , 10 mM MgSO 4 , 13 mM glycine, 3 µM thiamine, 15 mM 13 C-glucose, pH 5.5) supplemented with 1 mM 13 C 6 -L-DOPA (Cambridge Isotope Laboratories, CLM-1007-PK) 53 , 54 . Melanized fungal cells were collected and washed following the protocol described above and packed into 3.2 mm and 1.3 mm MAS rotors (Cortecnet, HZ14752) for solid-state NMR characterization. Scanning electron microscopy A small amount of the same C. neformans material used for NMR analysis were suspended in distilled water, mixed with 4% glutaraldehyde in 0.1 M sodium phosphate buffer (pH 7.4), and fixed for 30 min at 4°C. To prepare coverslip-mounted samples, 1% Poly-L-Lysine (Sigma Aldrich, P1399) was applied as a single drop onto a plastic Petri dish, over which a 12 mm round glass coverslip was placed and allowed to adhere for 10 min. The coverslip was then removed, gently rinsed with water, and drained while preventing complete drying. A drop of the fixed cell suspension was placed on the previously poly-lysine-exposed surface of the coverslip and allowed to settle for 10 minutes. The sample was subsequently rinsed with water and dehydrated through a graded ethanol series (25%, 50%, 75%, 95%) for 10 min at each step, followed by three 10-min changes in 100% ethanol and critical point drying using a Leica Microsystems EM CPD300 critical point dryer (Leica Microsystems, Vienna, Austria) with carbon dioxide as the transitional fluid. Dried samples were mounted on aluminum stubs, sputter-coated with osmium, and examined using a JEOL 7500F field emission scanning electron microscope operating at 15 kV. Total cell diameter was quantified using ImageJ software (version V1.8.0_172) and reported in Supplementary Table 1 . 13 C solid-state NMR experiments High-resolution 1D and 2D solid-state NMR experiments were conducted at 800 MHz (18.8 Tesla) at Michigan State University. All 13 C-detection experiments were performed using a 3.2 mm HCN probe at 15 kHz MAS, with ambient temperatures between 283 K and 298 K. 13 C chemical shifts were externally referenced by calibrating the adamantane CH 2 peak to 38.48 ppm, and the resulting spectral reference (sr) value was applied to fungal spectra. Unless otherwise specified, typical radiofrequency field strengths were 83-100 kHz for 1 H decoupling, 62.5 kHz for 1 H hard pulses, and 50-62.5 kHz for 13 C. Experimental parameters for all NMR spectra are documented in Supplementary Table 10 . 1D 13 C spectra were acquired using various polarization methods to probe molecular dynamics. Rigid components were detected via dipolar-based 13 C cross-polarization (CP) with a 1-ms contact time, adhering to Hartmann-Hahn match conditions of 62.5 kHz for 13 C and 1 H. For quantitative analysis, 1D 13 C direct polarization (DP) experiments were conducted with a long recycle delay of 35 s, ensuring full longitudinal relaxation before the next scan. Additionally, a shorter recycle delay of 2 s in the same 13 C DP experiment enabled selective detection of mobile molecules with fast 13 C-T 1 relaxation. These 1D experiments were performed on three independently prepared C. neoformans sample replicates, demonstrating high reproducibility ( Supplementary Fig. 10 ). To facilitate resonance assignment, 2D 13 C- 13 C correlation experiment was conducted using a 53-ms CORD (combined R2 n v -driven) mixing period, which revealed intramolecular cross-peaks between carbon sites within each molecule ( Supplementary Fig. 1 ) 43 . Additionally, 2D 13 C refocused J-INADEQUATE experiments were conducted using either CP or DP (1.5 s recycle delay) to probe molecular domains with differing mobility ( Supplementary Fig. 4 ) 66 . This experiment correlated double-quantum (DQ) chemical shifts with two corresponding single-quantum (SQ) shifts, generating asymmetric spectra that enabled efficient through-bond carbon-connectivity tracking. To optimize carbohydrate detection, the τ period (out of four) was set to 2.3 ms. All 1D experiments, as well as 2D CORD experiments, were conducted for all C. neoformans strains. Assigned chemical shifts for rigid and mobile polysaccharides are documented in Supplementary Table 2 . Data acquisition was conducted using Topspin 3.5, and spectral analysis was performed in Topspin 4.0.8. Figures were prepared using Adobe Illustrator CS6 (V16.0.0). Molecular composition analysis was conducted by selecting only well-resolved signals in 2D 13 C CORD spectra for rigid components and 13 C DP refocused J-INADEQUATE spectra for mobile molecules ( Supplementary Tables 3 and 4 ). Peak volumes were quantified using the integration function in Bruker Topspin, with quantification based on the mean of resolved signals. Relative polysaccharide abundance was determined by normalizing the sum of integrals with their respective counts 67 , 68 . The standard error for each polysaccharide was calculated by dividing the standard deviation of integrated peak volumes by the total cross-peak count. The overall standard error was computed as the square root of the sum of squared errors for each polysaccharide. The percentage error was determined by normalizing the standard error with the average integrated peak volume and adjusting for each polysaccharide’s relative abundance 67 , 68 . Solid-state NMR of polymer hydration and dynamics All polymer hydration and dynamics experiments were conducted on a Bruker Avance Neo 400 MHz (9.4 T) NMR spectrometer at Michigan State University using a 3.2 mm HCN MAS Bruker probe at 280 K. Water accessibility of polysaccharides was assessed using 1D and 2D water-edited 13 C- 13 C correlation spectra 45 , 47 . A 1 H-T 2 filter (0.6-1.2 ms, strain-dependent) was applied to suppress carbohydrate signals to less than 10% but preserve 82-88% of water magnetization. ( Supplementary Fig. 6 ), followed by 1 H- 1 H mixing to transfer water 1 H magnetization to hydrated carbohydrates. 13 C detection was achieved via CP with a 1-ms contact time. For 2D water-edited experiments, a 4 ms 1 H mixing period and 50 ms DARR mixing were used. Intensity ratios (S/S 0 ) between the water-edited (S) and control (S 0 ) spectra were obtained to quantify water retention around each carbon site ( Supplementary Table 5 ). 13 C-T 1 relaxation was measured using the Torchia-CP scheme 69 with z-filter durations from 0.1 μs to 8 s. For 13 C-detected 1 H-T 1ρ relaxation, a Lee-Goldburg (LG) spinlock sequence combined with LG-CP suppressed 1 H spin diffusion, enabling site-specific measurements via bonded 13 C detection 70 , 71 . For both measurements, peak intensity decay was fitted to a single exponential equation to determine the corresponding relaxation time constants ( Supplementary Figs. 7, 8 and Table 6 ). The analysis was conducted using OriginPro 9. Proton-detection solid-state NMR experiments 2D hCH, 2D hChH, and 3D hCCH TOCSY correlation experiments were conducted using a 600 MHz Bruker Avance Neo spectrometer at Michigan State University, equipped with a 1.3 mm triple-resonance MAS probe. Samples were spun at 60 kHz MAS. The 2D hChH experiment employed a 0.8 ms RFDR-XY16 (radiofrequency-driven recoupling) mixing 72 , 73 . A total of 192 time-domain (TD) points were acquired for the indirect dimension, with 512 transients co-added per TD and a recycle delay of 2.0 s. The total experimental duration was 58 hr. A 2D hCH spectrum was acquired on the same 600 MHz Bruker spectrometer under identical conditions, but without RFDR mixing, to enable direct comparison. The hCH experiment were measured using a short second-CP of 100 µs. The 2D data were collected using the States-TPPI method 74 . Through-bond 13 C- 13 C correlations were established using the 3D hCCH TOCSY (total correlation spectroscopy) experiment 75 , employing a 15 ms WALTZ-16 (wideband alternating-phase low-power technique for zero-residual splitting) mixing period at an rf field strength of 21.4 kHz 76 . A total of 128 × 128 TD points were acquired in the T 1 and T 2 evolution periods, with 8 transients co-added per TD point. The total experimental time was 78 h. The Heteronuclear dipolar decoupling sequence slpTPPM (swept low-power two-pulse phase modulation) 77 was applied during the T 1 evolution period for the 2D hCH and 2D hChH sequences, and during both T 1 and T 2 periods in the 3D hCCH TOCSY experiment, with a rf field strength of 21.0 kHz. For the direct 1 H detection period, WALTZ-16 decoupling was applied on the 13 C channel with a rf field strength of 12.5 kHz for 2D hCH and 2D hChH, and at 21.4 kHz for 3D hCCH TOCSY. Water suppression was achieved using the MISSISSIPPI (multiple intense solvent suppression intended for sensitive spectroscopic investigation of protonated proteins) sequence 78 on the 1 H channel, with a rf of 15.2 kHz applied for 100 ms. The actual sample temperature was measured to be 304 K, based on the 1 H chemical shift of water relative to the DSS signal at 0 ppm. 1 H and 13 C chemical shifts of C. neoformans polysaccharides, as well as 13 C chemical shifts of melanin fragments detected in 1 H-detected experiments, were documented in Supplementary Tables 7 and 8 . Experimental details are summarized in Supplementary Table 11 . MAS-DNP analysis of inter-polysaccharide interactions A stock solution containing 10 mM AsymPol-POK bi-radicals in a d 6 -DMSO/H 2 O (10/90 vol%) mixture was prepared 48. 13 C, 15 N-labeled C. neoformans H99 cells were mixed with 50 µL of the stock solution and gently ground using a set of mortar and pestle to ensure radical penetration and distribution into the porous cell walls. Approximately 30 mg of the processed sample was packed into a 3.2-mm sapphire rotor for DNP experiments. All experiments were conducted on a 600 MHz/395 GHz MAS-DNP spectrometer at the National High Magnetic Field Laboratory (Tallahassee, FL, USA), equipped with an 89 mm bore and a gyrotron microwave source. Data acquisition utilized a 3.2-mm HCN probe operating at 8 kHz MAS and 100 K. The gyrotron cathode current was maintained between 130-150 mA, with a voltage setting of 16.2 kV. The power of the microwave irradiation was 6.5 W. The NMR sensitivity enhancement (ε on/off ) was 11, and the DNP signal buildup time was ∼2.9 s for C. neoformans carbohydrate signals. The PAR spectrum was acquired with 15 ms recoupling duration, during which the 1 H and 13 C irradiation frequencies were set at 53 kHz and 50 kHz, respectively 49 , 50 . The 2D 13 C- 13 C DARR spectrum was recorded with a 100 ms mixing time. DATA AVAILABILITY All relevant data that support the findings of this study are provided in the article and supplementary Information. All the original ssNMR data files will be deposited in the Zenodo repository, and the access code and DOI will be provided. AUTHOR CONTRIBUTIONS A.A., I.G., and P.W. prepared labeled and unlabeled samples. A.A., I.G., D.R., and L.X. conducted 13 C solid-state NMR experiments. J.R.Y. performed 1 H-detection solid-state NMR experiments. F.J.S. and F. M.-V. performed DNP measurements. A.A., I.G., J.R.Y., C.C., R.E.S. and T.L.D. analyzed the data. P.W. and T.W. designed the experiments. All authors contributed to manuscript writing. COMPETING INTERESTS The authors declare no competing interests. ACKNOWLEDGMENT This work was primarily supported by the National Institutes of Health (NIH) grant R01AI173270 to T.W. A portion of this work was performed at the National High Magnetic Field Laboratory, which is supported by the National Science Foundation Cooperative Agreement No. DMR-2128556 and the State of Florida. The MAS-DNP system at NHMFL is funded in part by NIH RM1-GM148766. 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