Exploring the Ligninolytic Capabilities of the Oleaginous Yeast Cutaneotrichosporon oleaginosum

Exploring the Ligninolytic Capabilities of the Oleaginous Yeast Cutaneotrichosporon oleaginosum | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (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],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Article Exploring the Ligninolytic Capabilities of the Oleaginous Yeast Cutaneotrichosporon oleaginosum Austin Gluth, Yunqiao Pu, Dehong Hu, Xiaowen Chen, Zachary Johnson, and 5 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-6335743/v1 This work is licensed under a CC BY 4.0 License Status: Published Journal Publication published 18 Jan, 2026 Read the published version in Scientific Reports → Version 1 posted 10 You are reading this latest preprint version Abstract The current paradigm in synthetic biology for lignin bioconversion platforms includes primarily bacteria and filamentous fungi. Yeast are notoriously understudied for their role in lignin degradation and utilization, despite their ubiquity in saprophytic microbial communities. A few publications report ligninolytic yeasts, but investigations to date have relied on model aromatic compounds or lignin-containing substrates replete with other carbon sources. In this work, we use a suite of analytical tools to evaluate interactions between corn stover-extracted lignin and the oleaginous yeast Cutaneotrichosporon oleaginosum . Notably, 2D-NMR analysis showed a significant decrease in the H-lignin component as well as resinol (β-β) and phenylcoumaran (β-5) linkages. Using super-resolution fluorescence microscopy, we demonstrated that this yeast may uptake polymeric lignin. To explore mechanisms of lignin degradation, transport, and aromatics catabolism, extensive secretomics and proteomics analyses were conducted. Compared to carbon-limited glucose and “No Carbon” controls, several putative laccases, quinone reductases, superoxide dismutases, and glyoxal/oxalate oxidases were upregulated in the lignin condition. Excitingly, two ferric reductases and a oxalate exchanger were only observed on lignin. These results indicate that C. oleaginosum may perform extracellular quinone redox cycling to generate lignin-modifying reactive oxygen species. These findings enhance our understanding of lignin utilization by yeast and provide valuable insights for metabolic engineering. Biological sciences/Biotechnology/Proteomics Biological sciences/Microbiology/Fungi/Fungal biology Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Introduction Lignin constitutes a complex and diverse biopolymer, and, importantly, it is the largest renewable source of aromatics. As much as 40% of the energy density of lignocellulosic biomass is found in lignin, and it has a higher carbon content than carbohydrates: 68 wt% for the lignin derivative guaiacyl vs. 44% for cellulose based on glucose ( 1 ). Bacteria and filamentous fungi are prime candidates for leveraging this rich carbon source given their ligninolytic capabilities ( 2 – 5 ). In general these microorganisms degrade lignin using oxidoreductases such as laccases and peroxidases as well as other redox-active molecules ( 6 – 8 ). Compared to filamentous fungi, bacteria such as Pseudomonas putida have higher growth rates and are amenable to genetic manipulation; nevertheless, white rot fungi like Phanerchaete chrysosporium are superior lignin degraders ( 9 ). In contrast the potential of yeasts (particularly basidiomycetes) as biocatalysts for lignin valorization and their roles in lignin-containing microbial niches have rarely been investigated. As a result, a clarity regarding their ability to degrade (or at least modify) lignin and what enzymes may functionally confer these capabilities is lacking ( 10 ). Though scarce, there are reports of basidiomycete yeasts (and one ascomycete) with activities against lignin as the major carbon source. These include species of the genera Rhodotorula , Geotrichum , Tausonia , and Cutaneotrichosporon ( 11 – 14 ). Laccase, manganese peroxidase, and lignin peroxidase activities have been observed in yeasts—some of which display the oleaginous phenotype under nutrient-limiting conditions ( 14 – 17 ). Oleaginous yeast accumulate lipids and are promising candidates for sustainable synthesis of palm oil substitutes, lubricants, plastics, and fuels ( 18 – 21 ). Of these yeasts, Cutaneotrichosporon oleaginosum (formerly known as Cryptococcus curvatus and later Trichosporon oleaginosus) is poised for lignin valorization because it can accumulate lipids using the monoaromatic resorcinol ( 22 – 24 ). However, it is unclear if this yeast can degrade lignin and what enzymatic mechanisms are involved both extra- and intracellularly. A recent functional genomics analysis of C. oleaginosum cultivated on alkali-pretreated corn stover has expanded our knowledge of yeast aromatics catabolism but again raises questions about its ligninolytic abilities due to ambiguous results from K-lignin analysis, NMR spectroscopy, and enzymatic activity assays ( 14 , 25 ). For instance, a decrease in acetyl groups was observed whereas no changes to the (hemi)cellulose fractions were reported raising the possibility that the lignin in the black liquor was blocked ( 25 ). The hydrolysate derived from dilute-alkali corn stover pretreatment contains numerous accessible carbon sources apart from lignin: monomeric sugars, oligosaccharides, organic acids, and monoaromatics ( 26 , 27 ). Other studies have used modified (Kraft) lignin in the presence of glucose, which obfuscates secretomics results ( 28 ). We cultivated C. oleaginosum on lignin purified from alkali-pretreated corn stover to directly evaluate lignin degradation/modification ( 29 ). We hypothesize that the proposed yeast secretes ligninolytic enzymes during carbon limitation, and that liberated aromatics and polymeric lignin are proximally transported into the cell. Our 2D-NMR results demonstrate decreases in the H- and G-lignin motifs and relative changes to bonding in the lignin structure. Our fluorescence microscopy results suggest cellular uptake of polymeric lignin. Passive and active transport of lignin-derived aromatics has been studied, but there is a dearth of information regarding lignin fragments ( 30 – 33 ). Saccharomyces cerevisiae can uptake nanoparticles of polyaromatic polystyrene, which indirectly supports yeast lignin uptake ( 34 ). Our fluorescence microscopy approach which harnesses the native fluorescent properties of lignin corroborates this ( 35 , 36 ). Finally, we developed a comprehensive secretomics and proteomics workflow to study the expressed metabolism of C. oleaginosum , thereby pinpointing putative enzymes for lignin modification as well as transport and utilization of lignin-derived aromatics. This comprehensive work provides targets for functional evaluation and additional impetus for developing basidiomycete yeasts as chassis for lignin valorization. Results and Discussion Cultivation on Alkali Lignin is Reminiscent of the “No Carbon” Condition A multifaceted approach was employed to study the cultivation of C. oleaginosum on alkali lignin and is summarized in Fig. 1 . Lignin from dilute alkali-pretreated corn stover was extracted with successive acid-base titrations and analyzed via the K-Lignin assay. The overall lignin component was 87.06% with a low ash content of 0.30% ( SI Appendix , Table S1 ). Though the isolated lignin contained structural saccharides, the high lignin and low ash contents are ideal for this work. Four conditions were evaluated in this study: 1 g/L lignin, 1 g/L glucose, 1 g/L benzoate, and a “No Carbon” control in which no additional carbon source was added to the base medium. The “No Carbon” control was included to uncover differences specifically attributed to interactions with lignin. An additional control of uninoculated medium with 1 g/L lignin was included to check for contamination and as a baseline comparison for absorbance assays. Additional analyzes of growth, lignin modification, and lignin uptake were conducted to lay a foundation for studying the extracellular and cellular proteomes of C. oleaginosum . After 144 hours of cultivation, a decrease in cell dry weight (CDW) was observed for the lignin condition (Fig. 2 A). Despite this, 8.5% and 11.3% decreases in lignin content were observed at 96 and 144 hours, respectively, according to UV absorbance at 320 nm (Fig. 2 B). These results were confirmed using the Prussian Blue assay (Fig. 2 C). The 96-hour time point was chosen to compare the CDWs of different carbon-limited conditions. Unexpectedly, a slight albeit significant difference between the “No Carbon” and lignin conditions was recorded and was supported by an increase in viable cell counts with minimal differences in cell viability (Fig. 2 D–F). It’s possible that the observed difference in growth is due to utilization of structural saccharides and/or acetyl groups, which constitute ~ 13% of the alkali lignin ( SI Appendix , Table S1 ). At 96 hours, lipid titers and fatty acid compositions for the “No Carbon” and lignin conditions were similar ( SI Appendix , Fig. S1 ). Stearic acid (C18:0) was not observed in these two conditions, and a general increase in fatty acid desaturation was apparent, indicating that lipids were mobilized in the “No Carbon” and lignin conditions for carbon and energy generation ( 37 , 38 ). Overall, increased growth in the lignin condition vs. the “No Carbon” condition was unexpected because of the sheer recalcitrance of lignin. NMR Results Demonstrate Selective Modification of Alkali Lignin Two-dimensional (2D) heteronuclear single-quantum correlation (HSQC) NMR spectroscopy was used to study the effects of C. oleaginosum on lignin’s chemical structures (Fig. 3 A–B). HSQC provides a semi-quantitative analysis of lignin’s structural features such as monolignol composition and interunit linkages. Lignin recovered from the lyophilized yeast culture supernatant was compared to the control alkali lignin. NMR analysis revealed changes to lignin structures after yeast cultivation. The control lignin had a S/G/H ratio of 57:31:12, whereas after yeast fermentation, the lignin in the supernatant had a ratio of 66:29:5. Compared to the control, this is approximately a 59.8% decrease in the H-lignin unit, which is comprised of the p -coumaryl alcohol monomer. This suggests that the yeast preferentially utilizes un-methylated aromatics, or that p -coumarate, p -hydroxybenzoate, and p -hydroxyphenol pendents are more susceptible to liberation. Predominant release of these H-lignin derived aromatics was reported recently and lends some support to the latter ( 39 ). It follows that we observed a 16.5% decrease in p -coumarate esters and a 13.4% decrease is ferulate esters as previously reported ( 25 ). We did not observe a significant decrease in the S/G (syringyl/guaiacyl) ratio, whereas a minor potentially insignificant decrease was observed previously using alkali-pretreated corn stover ( 25 ). Given the decrease in total aromatics (Fig. 2 B–C), it’s possible that a simulatenous decrease in S/G motifs occurred. The aliphatic region showed a 75.8% and 60.5% depletion of resinol (β-β) and phenylcoumaran β-5 linkages, respectively (Fig. 3 B–C). Interestingly, there was a slight increase of the overall β-O-4 linkage signal. Among the various types of β-O-4 linkages, a small decrease in G/H-type was observed, but the S-type β-O-4 linkage dominated. The decrease of G/H-type β-O-4 could be attributed in part to the decrease of H-type β-O-4 linkage, consistent with the decrease of H unit as observed from aromatic regions of the HSQC spectra. Previous 2D-NMR results of this yeast grown on alkali-pretreated corn stover demonstrated no decrease in β-O-4; however, the high presence of contaminating polysaccharides complicates interpretation ( 25 ). The two new cross peaks at δ C /δ H 68.2/3.91 and δ C /δ H 72.2/3.48 ppm in the aliphatic region correspond to Ar-CH(OH)-O-CH 2 -COOH. In studies of another basidiomycete yeast as well as a white rot fungus cultivated on lignin, 1D-NMR results demonstrated that these newly formed peaks arise from the cleavage of bonds between lignin subunits ( 13 , 40 ). Compared to those and other studies, a 96-hour time point caused apparent changes of lignin structures, but longer cultivation time may be needed to observe more lignin degradation ( 11 , 12 , 14 , 41 ). However, a decrease in ligninolytic activity during prolonged cultivation has been reported, and our results directly demonstrate the ligninolytic capacity of C. oleaginosum ( 42 ). Secretomics and Proteomics Reveal Stark Contrasts between Carbon-limited Conditions Teasing out differences in protein expression due to the presence of lignin is crucial for discovering ligninolytic oxidoreductases and related enzymes. The single-pot, solid-phase-enhanced sample preparation (SP3) approach facilitated high protein digestion efficiencies ( 43% total coverage of the modeled genome) compared to recent studies of C. oleaginosum ( SI Appendix , Supporting Text and Table S4 ) ( 43 – 45 ). In the secretomics results, 245 unique proteins passed the SignalP signal peptide prediction score of 0.85, and 14 of these were only observed in the lignin condition (Fig. 5 A and SI Appendix , Dataset S1) ( 46 ). 3,579 unique proteins were observed in the proteomics results with 56 of those only observed in the lignin condition (Fig. 5 A and SI Appendix , Dataset S2). The Pearson correlation coefficients for lignin vs. glucose replicates were ~ 0.60 and ~ 0.90 for the secretomics and proteomics results, respectively (Fig. 5 B–C). Proteins unique to the lignin condition were also observed for the lignin vs. “No Carbon”, along with a lower Pearson correlation coefficient (~ 0.80) for the secretomics data ( SI Appendix , Fig. S2 A–B). For cellular proteomics results, correlation between lignin vs. “No Carbon” samples was high (~ 0.97), which is expected due to carbon starvation ( SI Appendix , Fig. S2 C). Nevertheless, principle component analyses show a clear separation for each condition ( SI Appendix , Fig. S3 ). Carbohydrate active enzyme (CAZymes) and differential expression results are summarized in Fig. 5 D–E. CAZy annotations from the Conserved Unique Peptide Patterns (CUPP) platform were used because of the protein-grouping capabilities of the CUPP algorithm, which can help distinguish certain functional annotations beyond CAZy subfamilies ( 47 ). 42 CAZymes were observed in the secretome, while 51 were observed intracellularly (Fig. 5 D). For both the secretomics and cellular proteomics datasets, relative quantification was performed using LFQ intensities (“DE-filtered”, Fig. 5 D). In the lignin vs. glucose secretomics data, 45 proteins had log 2 fold changes (“FC”) > 1 and adjusted p-values < 0.05, while 44 were significantly downregulated (Fig. 5 E). In the cellular proteomics data, 207 proteins were upregulated, and 130 were downregulated. To provide an overview, pathway enrichment was conducted (Fig. 5 F). It’s evident that an expression regime was upregulated to access diverse carbon sources (e.g., galactose, sucrose, flavonoids/aromatics, glycans, etc.) during carbon limitation. Upregulation of N-glycan synthesis suggests restructuring of the cell wall perhaps in response to stress. Lastly, the prevalence of amino acid and other carbon recycling pathways (e.g., fatty acid degradation) was expected; however, it is intriguing to see downregulation of glutathione and cysteine/methionine metabolic pathways. This may highlight the importance of antioxidants during carbon limitation ( 48 ) and raises questions about the toxicity of lignin/aromatics and byproducts of lignin breakdown. With deep coverage proteomics results, we projected an extracellular mechanism for yeast lignin modification (SI Appendix, Fig. S4 ) and routes for funneling aromatics to central metabolism (Fig. 6 ) in the following subsections. C. oleaginosum Expresses Oxidoreductases for Extracellular Quinone Redox Cycling Lignin degradation is an oxidative extracellular process involving the concerted effort of reactive oxygen species (ROS), Fenton chemistry, and various enzymes ( 49 ). This complex process requires extracellular production of ROS and aromatic radicals including hydrogen peroxide (H 2 O 2 ), hydroxyl radical ( −• OH), superoxide (O 2 −• ), cation radicals, and phenoxy radicals ( 50 ). These highly reactive species can be harnessed by electron-transferring auxiliary activity (AA) family oxidoreductases to attack lignocellulose ( 51 ). In addition to a repertoire of laccases (AA1) and peroxidases (AA2), fungi also employ non-enzymatic mechanisms to modify lignin ( 52 – 56 ). At the heart of this mechanism is the hydroquinone-quinone redox cycle that reduces Fe 3+ to ultimately produce H 2 O 2 for initiating enzymatic catalysis, and −• OH to react directly with lignin units ( 49 , 50 ). Interestingly, whether or not basidiomycete yeasts have the functional capacity for quinone redox cycling has yet to be addressed according to Web of Science and PubMed searches using variations of the following wildcard search queries: ((yeast) AND (hydroquinone redox cycl* OR quinone redox cycl* OR Fenton chemistry)) AND (lignin OR aromatic*). There are several requirements for the hydroquinone redox cycle: oxidoreductases, reductants, and an acidic pH ( 50 , 57 ). A combination of BLASTp searches and CAZy CUPP annotations were used to find candidates in our proteomics data ( SI Appendix , Fig. S4 and Dataset S3) ( 47 , 51 , 58 ). Ferric reductases are important for iron uptake (generally as Fe 2+ ) and maintaining an active quinone redox cycle ( 59 , 60 ). Unfortunately, none of the ferric reductases annotated in the Joint Genome Institute (JGI) database were observed intracellularly; however, they were observed in the secretome despite being localized to the cell membrane ( 59 , 61 ). These include Co_379781, Co_368607, and Co_366398 (the latter only observed in lignin replicates), which share 35–40% identity with a ferric reductase (Pospl1_130030) from the model brown rot fungus Rhodonia placenta MAD 698R ( SI Appendix , Dataset S3) ( 62 , 63 ). Co_414234 was also only observed in the lignin condition and has a ferric reductase transmembrane component-like domain (IPR013130) ( 64 ). In the absence of ferric reductases, Fe 3+ can still evolve as a result of Fe 2+ oxidation and participate in oxidation of hydroquinone and semiquinone species ( 49 ). Fe 2+ can react with oxygen to generate O 2 −• and with H 2 O 2 to produce −• OH + − OH—the latter may partially explain the observed pH increase in the lignin-containing cultures ( SI Appendix , Fig. S4 ). H 2 O 2 is generated from other sources including O 2 −• ( 50 ). Superoxide dismutase (SOD) catalyzes the conversion of O 2 −• to H 2 O 2 and is essential for combatting oxidative stress ( 50 ). Though localization predictions from Tremellomycetes loosely related to C. oleaginosum suggest intracellular localization, there are several fungi that secrete SOD variants ( 50 , 65 ). Two copper/zinc-dependent SODs (Co_394365 and Co_419771) with predicted signal peptides and secretion predictions were observed ( 66 ). Co_419771 was significantly upregulated in the lignin condition (FC 6.51), which suggests severe oxidative stress. Glucose–methanol–choline (GMC) oxidoreductases (AA3), glyoxal oxidases (AA5_1), and oxalate oxidases can also produce H 2 O 2 using a variety of reductants ( 51 ). GMC oxidoreductases (AA3_2) can utilize aryl-alcohols and sugars as electron donors, whereas glyoxal oxidases convert simple aldehydes to carboxylic acids ( 67 , 68 ). Oxalate is an important iron chelator during wood decay, and is detoxified to carbon dioxide and H 2 O 2 via oxalate oxidase ( 69 , 70 ). Two GMC oxidoreductases were observed extracellularly (Co_352536 and Co_ 372521) and one intracellularly (Co_342568; DEP FC 4.94). The first two GMC oxidoreductases belong to CUPP branch AA3:6.5, while the other may have a different function as it is categorized in CUPP branch AA3:10.1. Two glyoxal oxidases were observed in the secretomics dataset (Co_344356 and Co_26860). The latter had a signal peptide prediction score of 0.69 but was significantly upregulated in the lignin condition (FC 2.26). An enzyme of the cupin superfamily (Co_373698) was also significantly upregulated (FC 2.73) and is classified as an oxalate oxidase (or superoxide dismutase) by InterPro (IPR001929) ( 64 ). Two putative sulfate/bicarbonate/oxalate exchangers were observed in the proteomics results: Co_335524 and Co_352469, which was only observed in the lignin condition. Laccases and quinone oxidoreductases play significant roles in quinone redox cycles. Laccases are multicopper oxidases that use O 2 to oxidize hydroquinones to semiquinones ( 52 , 71 ). Semiquinones are unstable products that undergo autooxidation to form quinones ( 49 ). Two laccases with predicted signal peptides were upregulated in the secretome of the lignin condition: Co_368910 (FC 5.57) and Co_ 369862 (FC 5.68) ( SI Appendix , Fig. S4 ). Another putative laccase Co_416150 that shares > 37% identity with a laccase from the white-rot fungus Trametes versicolor ( SI Appendix , Dataset S3) was observed but downregulated in the secretome (FC -0.96) ( 72 ). Interestingly, it was only observed in the cellular proteome of the lignin condition. Co_368910, Co_372508, and Co_416150 are secreted AA1 CAZymes according to MULocDeep prediction, and the first two are members of CUPP branch AA1:73.1. ( 66 ). In contrast, Co_369862 may locate to the cell membrane and was downregulated (FC -3.29) ( 66 ). It is a member of the subfamily AA1_2, which includes ferroxidases that oxidize Fe 2+ to modulate Fenton reactions and decrease −• OH ( 73 , 74 ). Indeed, the eggNOG annotation for this enzyme was FET3, a ferroxidase involved in iron uptake. Nonetheless, this enzyme shares 30–45% identity with phenol-oxidizing laccases from T. versicolor ( SI Appendix , Dataset S3) ( 72 , 75 , 76 ). Studies of FET3-like proteins in basidiomycetes show that these multicopper oxidases ostensibly have dual ferroxidase/laccase activities ( 72 , 75 , 77 , 78 ). More research into putative yeast laccases is clearly needed. Quinone oxidoreductases including 1,4-benzoquinone reductase (AA6) are involved in aromatics degradation and protection against reactive quinones ( 51 , 79 – 81 ). A putative quinone oxidoreductase (Co_369953; FC 0.69) and two annotated 1,4-benzoquinone reductases (Co_380727, FC 1.30; Co_164578, FC 0.66) were upregulated in the lignin condition of the proteomics results ( SI Appendix , Fig. S4 ). According to BLASTp results, these enzymes share > 45% identity with quinone oxidoreductases from Rhodonia placenta ( SI Appendix , Dataset S3) ( 63 ). Moreover, Co_380727 is a predicted cell membrane protein, which has > 45% identity with functionally characterized quinone oxidoreductases from the white rot fungus Phanerochaete chrysosporium and brown rot fungus Gloeophyllum trabeum ( 66 , 80 , 82 ). A cyptoplasmic protein Co_342726 only observed in the lignin replicates was annotated as QOR1, a quinone oxidoreductase, by eggNOG; nevertheless, a conserved domain search on NCBI returned a prostaglandin dehydrogenase (cd05288) ( 83 ). In addition to the quinone redox cycle, peroxidases can also oxidize hydroquinone to yield 1,4-benzoquinone ( 79 ). Though many peroxidases were observed in the secretome (Co_396749, Co_237718, Co_382630, and Co_372514), none of them were upregulated, and only one (Co_396749) had a secretion prediction ( 66 ). Given that both the glucose and lignin culture conditions were carbon-limited, this may be explained by general upregulation of genes under carbon catabolite derepression ( 84 ). Peroxidase expression may also be a function of culture conditions like nitrogen source as Martinez et al. observed with Pleurotus eryngii ( 49 , 85 ). Nevertheless, peroxidase activity has been observed with C. oleaginosum as well as another Trichosporonaceae yeast ( 14 , 86 ). BLASTp results of AA2 peroxidases from T. versicolor , P. chrysosporium , and the recently isolated relative Cutaneotrichosporon cavernicola yield Co_382630 and Co_372514 with > 55% identity ( SI Appendix , Dataset S3) ( 50 , 87 ). Interestingly, these enzymes were not annotated by CUPP nor were AA2 CAZy annotations included for these entries on JGI ( 43 , 47 , 51 , 88 ). Based on the aforementioned and because these enzymes do not have predicted signal peptides, it’s likely that these enzymes are AA2 class I peroxidases. This class of peroxidases has lower redox potentials than class II lignin and manganese peroxidases, and they are generally involved in oxidative stress response ( 89 ). These proteomics results suggest a capacity for quinone redox cycling by C. oleaginosum , even though the reactivity of Fenton chemistry was likely suboptimal under the near neutral culture conditions studied here (starting pH of 6.5; see SI Appendix , Fig. S4 ) ( 57 ). To bolster this conclusion, the mechanisms by which hydroquinones are generated de novo and secreted to help initiate redox cycling will need to be studied in yeast. Hydroquinone secretion has been observed with filamentous fungi; however, it is unclear what transporters are involved because few have been functionally characterized ( 54 , 90 , 91 ). Of course, this is also the case for the other candidate enzymes discussed hitherto—direct functional genomics analyses will be essential for resolving ambiguities regarding this yeast’s laccase and peroxidase activities ( 14 , 25 ). Overall, these findings highlight the need for additional research to validate the proposed candidates and fully understand the quinone redox cycling potential of C. oleaginosum and other yeasts. C. oleaginosum May Uptake Poymeric Lignin but the Mechanism Remains Unclear Transport mechanisms for lignin-derived aromatics in yeast are underexplored ( 92 , 93 ). In bacteria and filamentous fungi, active transport of monoaromatics involves ATP-binding cassette (ABC) transporters and major facilitator superfamily (MFS) transporters ( 33 ). Passive transport has also been suggested, though most research has been conducted in bacteria which have drastically different cellular envelopes ( 30 , 94 ). Proteomics can help elucidate potential aromatics transporters by highlighting those that are upregulated or only observed in the lignin condition. We observed 380 of the 790 JGI-annotated transporters ( 43 , 95 ). The relative abundances of 246 were quantified for lignin vs. glucose: 48 were upregulated (FC > 0.8) while 18 were downregulated (FC < − 0.8). Two enzymes annotated as p -coumarate-CoA synthases were upregulated (Co_383485, FC 2.57; and Co_386597, FC 1.26; SI Appendix , Dataset S2). These are annotated members of a Fatty Acid Group Translocation Family that use ATP to attach CoA to p -coumarate and other aromatics in a process that may be coupled to transport ( 43 , 66 , 95 ). Upregulation of these enzymes is in line with a decrease in H-lignin (Fig. 3 A and C). Enzymes with cell membrane localization predictions include an arabinose, xylose, and/or aromatics transporter (Co_378211; FC 0.83), a MFS transporter (Co_419501; DEP FC 3.06), and an amino acid transporter (Co_391320; FC 1.37) with a phenol hydroxylase domain (IPR012941) ( 64 ). A MFS high affinity nicotinate permease (Co_96030) and an ABC transporter (Co_345222) were observed exclusively in the lignin condition. The latter is annotated as a plant cuticular wax and/or lipid exporter. Plant cuticles have complex compositions that include aromatics ( 96 ). Lastly, a MFS quinate:H + symporter (Co_96856) was only observed in the lignin condition. Fewer transporters were upregulated in our results compared to a recent study of the yeast Rhodosporidium fluviale cultivated on black liquor, but this may be due to substantially higher aromatics concentrations in their medium ( 28 ). We also conducted searches using hydroxybenzoate transporters recently characterized in the pathogenic yeast Candida parapsilosis , but no high-scoring alignments were found ( SI Appendix , Dataset S3) ( 97 ). A recent publication reported active transport and subsequent degradation of an aromatic dimer by P. chrysosporium ( 93 ). However, there is limited research explicitly addressing lignin uptake in microbial systems. We speculate that polymeric lignin can be internalized by C. oleaginosum based on uptake of polyaromatic polystyrene in yeast ( 34 , 98 , 99 ). As a preliminary investigation, we used confocal fluorescence microscopy to harness the autofluorescent signatures of lignin and study its uptake (Fig. 4 A–B and SI Appendix , Fig. S5 ) ( 35 , 36 ). We employed the cell segmentation software Cellpose to analyze yeast autofluorescence using various controls ( 100 – 102 ). Cellpose identified cells with high accuracy even in the presence of lignin particles, thereby facilitating cell population measurements of fluorescence ( SI Appendix , Fig. S6A). Though unapparent to the naked eye, cells in the lignin condition have significantly higher fluorescence intensities for all comparisons except lignin vs. benzoate at 48 hours (Fig. 4 B and SI Appendix , Fig. S6B–C). It’s possible that the increase in yeast autofluorescence confounds any differences, perhaps due to carbon-limited stress response and/or aromatics toxicity, given that 1 g/L benzoate is close to the minimum inhibitory concentration of C. oleaginosum ( 22 , 100 , 103 , 104 ). We also performed super resolution fluorescence imaging to illustrate structural changes due to lignin interactions (Fig. 4 C). For conditions without lignin, the weak autofluorescence is consistent with the structure of mitochondria. With lignin the fluorescence is more punctate and widespread throughout the cell, indicating significant changes to subcellular morphology. Moreover, there is a clear delineation of the cellular envelope. Using a dye to label mitochondria, the observed autofluorescence in the absence of lignin is partially due to mitochondria, whereas in the presence of lignin there is no apparent dye localization (Fig. 4 D). This suggests drastic changes to subcellular processes and possibility of lignin-induced membrane destabilization, which has been observed using other nanoparticles ( SI Appendix , Supporting Text) ( 105 ). Overall, these results suggest polymeric lignin uptake by C. oleaginosum , but investigations employing labeled lignin nanoparticles and electron microscopy will be needed for validation and to explore mechanisms. Pathways for Aromatics Catabolism Are Upregulated in the Lignin Condition Yeasts funnel aromatic monomers to intermediates such as protocatechuate, catechol, and hydroxyquinol via intricate upper pathways ( 106 ). Dioxygenases cleave the aromatic rings yielding intermediates that are metabolized through lower pathways (e.g., branches of the β-ketoadipate pathway), ultimately leading to central metabolites ( 107 , 108 ). Aromatics catabolic enzymes and the corresponding pathways are largely uncharacterized in C. oleaginosum . In an earlier report, cleavage positions for catechol, protocatechuate, and hydroxyquinol were assayed using clarified lysates ( 25 ). Catechol and hydroxyquinol underwent ortho -cleavage, whereas cleavage of proteocatechuate was negligible suggesting that it is decarboxylated to hydroxyquinol or catabolized via some other route. They also functionally annotated a catechol dioxygenase (homolog of Co_374756) that has intradiol and extradiol ring-cleaving activities. Beyond these analyses, specific catabolic pathways were not proposed; therefore, we scoured publications in which aromatics catabolic enzymes were functionally characterized in fungi (and some cases bacteria). We then conducted extensive BLASTp searches against proteins observed in our proteomics results to project putative aromatics catabolic pathways in C. oleaginosum ( SI Appendix , Datasets S1–3) ( 58 ). Rationalized by our 2D-NMR results and by the significance of ferulate G-lignin monomers in corn stover ( 109 ), we focused our efforts on projecting catabolic pathways for H-lignin derived aromatics (refer to SI Appendix , Supporting Text for more information about predicted pathways for G-lignin, S-lignin, and phenol derivatives). C. oleaginosum utilizes H-lignin model aromatics p -coumarate and p -hydroxybenzoate, but pathways have yet to be projected ( 110 – 112 ). In filamentous fungi, conversion of p -coumarate to p -hydroxybenzoate has been extensively studied ( 113 – 116 ). Lubbers et al. demonstrated that Aspergillus niger uses a CoA-dependent β-oxidative route akin to fatty acid degradation ( 117 ). Briefly, p -coumarate is converted to p -coumaroyl-SCoA by a hydroxycinnamate-CoA synthase (hcsA). The product is then hydrated and dehydrogenated by a multifunctional β-oxidation hydratase/dehydrogenase (foxA) to yield 4-hydroxyphenyl-β-ketopropionic acid-SCoA (HPOP-CoA). Afterwards, p -benzoyl-CoA is generated by 3-ketoacyl-CoA thiolase (katA). Finally, thioesterases remove CoA to produce p -hydroxybenzoate. We searched our proteomics data for homologs of characterized proteins. The BLASTp scores for Aspergilli hcsA were fairly low, so we also searched bacterial enzymes involved in the CoA-dependent non-oxidative pathway (Fig. 6 ; SI Appendix , Dataset S3) ( 117 – 121 ). Several putative genes were upregulated in the cytoplasm and peroxisome: Co_383485 (FC 2.57; cytoplasm), Co_386597 (FC 1.26; cytoplasm), Co_386202 (FC, 2.09; peroxisome), Co_376641 (FC 1.32; peroxisome), and Co_370579 (FC 1.05; peroxisome). Only one peroxisomal foxA homolog was upregulated (Co_233545; FC 0.98) and contained a C-terminal peroxisomal targeting sequence ( 66 , 122 , 123 ). BLASTp results of bacterial hydroxycinnamoyl-CoA hydratase-lyases returned Co_381918 (FC 0.93; peroxisome). Though many katA and thioesterase homologs were observed, few had peroxisomal predictions. According to MULocDeep, Co_368669 and Co_371904 are localized in the cytoplasm; however, both have C-terminal peroxisomal targeting sequences ( 66 , 122 , 123 ). β-oxidation of fatty acids was upregulated in the lignin condition (Fig. 5 F): those proteins may be candidates for CoA-dependent p -coumarate catabolism in C. oleaginosum . Prior to ring cleavage, p -hydroxybenzoate is converted to hydroxyquinol (Fig. 6 ). Phenol hydroxylase is active on phenol derivatives and has been characterized in the close relative Cutaneotrichosporon cutaneum ( 124 , 125 ). Three homologs were upregulated in our results: Co_391320 (FC 1.37), Co_374925 (FC 1.64), and Co_222522 (FC 2.31). Phenol hydroxylase from C. cutaneum is structurally related to fungal 4-hydroxybenzoate 3-hydroxylase, so these homologs may be involved in the conversion of p -hydroxybenzoate to protocatechuate ( 126 ). Oxidative decarboxylation of p -hydroxybenzoate and protocatechuate to hydroquinone and hydroxyquinol, respectively, has been reported for the yeast C. parapsilosis and white-rot fungi ( 127 , 128 ). Co_382250 was the highest scoring hit for 4-hydroxybenzoate 1-hydroxylase (i.e., MNX1) from C. parapsilosis but was not observed in our proteomics results, but the putative homolog Co_419806 was observed and upregulated 1.56-fold. It has a low alignment score with MNX1 but higher scores against putative basidiomycete decarboxylases ( 127 ). This is also the case for Co_417242 which was drastically upregulated (FC 4.25) and shows higher homology with a reported T. versicolor decarboxylase (TV_32834) that preferentially decarboxylates protocatechuate. The putative Gelatoporia subvermispora homolog (GS_120062) of Co_ 419806 displays activity against p -hydroxybenzoate and protocatechuate ( 127 ). Ring cleavage detoxifies aromatics and yields assimilable carbon sources ( 129 ). Enzymes that catabolize hydroxyquinol and protocatechuate have been investigated using C. parapsilosis and A. niger ( 128 , 130 ). Based on BLASTp results, Co_374756 is the top candidate for hydroxyquinol 1,2-dioxygenase activity ( SI Appendix , Dataset S3). Co_85172 is an additional candidate (FC 2.44), but it had a higher E-value (5.50E-39) against a protocatechuate 3,4-dioxygenase (NRRL3_1405) from A. niger . Another enzyme Co_340256 was annotated as the catalytic LigB subunit of aromatic extradiol ring-opening dioxygenase (PF02900) ( 64 , 76 ). LigB is a subunit of the heterodimer LigAB that exhibits protocatechuate 3,4-dioxygenase activity in the lignin-degrading bacteria Sphingobium ( 131 ). Intradiol hydroxyquinol cleavage yields maleylacetate, which is converted to β-ketoadipate by maleylacetate reductase. Negligable upregulation of Co_373930 (FC 0.25), a homolog of the A. niger candidate, was observed ( 132 ). A homolog of another candidate observed in C. parapsilosis was observed but not differentially expressed (Co_374156) ( 133 , 134 ). Intradiol cleavage of protocatechuate leads to formation of β-carboxymuconate, which undergoes cyclization via muconate cycloisomerase ( 135 ). The product γ-carboxymuconolactone can be decarboxylated to β-ketoadipate enol-lactone as observed in A. nidulans ( 136 ). A homolog of the decarboxylase studied by Martins et al. ( 136 ) was not observed in C. oleaginosum . Thankfully, the eggNOG description of Co_371232 (FC 2.94) was “Carboxymuconolactone decarboxylase family”, and the orthologous group annotation was “Sphingomonadales”, which raises interesting questions about horizontal gene transfer among soil-dwelling microorganisms ( 76 ). Two additional candidates include Co_350958 (FC 2.52; aminocarboxymuconate-semialdehyde decarboxylase) and Co_372335. As a whole, we speculate that mechanisms for lignin modification are present in C. oleaginosum and demonstrate upregulation of predicted catabolic pathways for H-lignin and G-lignin derivatives ( SI Appendix , Datasets S1–2). It follows that entry point enzymes for the TCA cycle and glyoxalate shunt were also upregulated, whereas glycolytic enzymes were downregulated (Fig. 6 ). Antioxidants are important for mitigating excess ROS such as H 2 O 2 , which can be generated extracellularly and re-enter the cell via direct permeation or aquaporins ( 50 ). We observed upregulation of several antioxidants including catalases (Co_370626, FC 2.07; Co_371234, FC 1.60) that neutralize H 2 O 2 . Nevertheless, many of these enzymes were only partially upregulated in the lignin vs. No Carbon comparison, which speaks to their general role in nutrient-limited stress response ( 48 ). For specific enzymes like glutathione-dependent formaldehyde-activating enzyme (Co_398886, FC 1.42), detoxification of formaldehyde from demethylation of aromatics may rationalize these differences ( 137 ). These findings hint at similarities in aromatics catabolism between yeasts and filamentous fungi of the Agaricomycotina subphylum, yet they also underscore gaps in our understanding. The lack of metabolomics results herein presents a significant opportunity for future studies but also poses challenges. These include limitations in current techniques, the scarcity of certain aromatics released even by white-rot fungi, and the intricate nature of lignin degradation intermediates ( 127 , 138 ). Assays that directly measure ROS generation will also further our understanding of redox mechanisms involved in aromatics detoxification and lignin degradation ( 139 ). Redox proteomics, which assesses protein oxidation, offers an innovative approach to explore how fungi manage redox homeostasis during lignin degradation ( 140 , 141 ). Conclusion In this exploratory study, we cultivated C. oleaginosum on alkali lignin and, according to 2D-NMR, observed modification of the H-lignin structure, p -coumarate and ferulate pendents, and interunit bonding. A comprehensive proteomics analysis revealed the potential for yeast quinone redox cycling as well as putative pathways for aromatics catabolism, which were upregulated in the lignin condition. Based on these results, we conclude that this yeast and likely other basidiomycete yeasts have a ligninolytic capacity, thereby playing salient roles in lignocellulose degradation and aromatics detoxification in native microbial communities. Nonetheless, challenges in analyzing secretomics data, especially under stressful carbon-limited conditions that may cause cell lysis, highlight the complexity of interpreting these results. Additionally, the influence of culture conditions, sampling time points, and localization of putative ligninolytic enzymes (e.g., Cryptococcus neoformans expresses cell well-bound laccase) further complicates interpretation of results using this yeast ( 142 , 143 ). These insights call for continued exploration and refinement of methodologies to enhance our understanding of the ligninolytic potential of yeasts. This will advance efforts to metabolically engineer yeasts to express complementary ligninolytic enzymes from white-rot fungi and optimize the utilization of lignin-derived aromatics. Methods A brief summary of relevant materials and methods is presented below. Additional details are included in SI Appendix , Supplementary Methods. Lignin Extraction from Corn Stover The National Renewable Energy Laboratory provided the black liquor fraction from dilute alkali-pretreated corn stover, which was used for lignin extraction ( 144 , 145 ). The extraction process involved serial dilute acid-base titrations, centrifugation, and solids washing ( 29 ). Compositional analysis details for the resulting extract are provided in SI Appendix , Table S1 ( 146 ). Yeast Strain, Medium Summary, and Cultivation Cutaneotrichosporon oleaginosum ATCC 20509 (formerly classified as Cryptococcus curvatus ) was used in this study. For long-term storage, glycerol (15% v/v) stocks stored at − 80°C were prepared from cells streaked on standard Yeast extract-Peptone-Dextrose (YPD) plates. YPD was used for short term plate storage at 4°C as well as generating seed cultures. The yeast was cultivated in a YNB-based experimental medium containing 1 g/L carbon source (lignin extract, glucose, or benzoate) or no additional carbon source (“No Carbon” condition). Cultivation conditions were 28°C shaking at 180 rpm. 2D-HSQC-NMR Two-dimensional (2D) heteronuclear single-quantum correlation (HSQC) NMR spectroscopy was used to compare alkali lignin (control) to lignin in the lyophilized yeast culture supernatants ( 147 , 148 ). Fluorescence Microscopy Live yeast cultures were imaged by an inverted fluorescence confocal microscope with oil immersion. The fluorescence from lignin was excited by a 488 nm wavelength laser according to a preliminary analysis of lignin excitation and emission wavelengths ( SI Appendix , Fig. S5 ). The Cellpose software was employed for cellular segmentation to analyze yeast cell fluorescence for the aforementioned carbon source conditions. Secretomics and Cellular Proteomics Supernatants and washed pellets were collected from yeast cultivations to generate extracellular (secretomics) and intracellular (cellular proteomics) samples, respectively. All samples were processed using the SP3 method but with modifications depending on the sample type (Fig. 1 ) ( 149 , 150 ). All samples were digested with an enzyme cocktail containing trypsin (Promega) and Lys-C (Wako Chemicals). Samples were then processed with Zip-Tip C18 SPE and analyzed using LC-MS/MS. Data Analysis and Processing Two-tailed t-tests were used for determining statistically significant differences in growth, total aromatics, and fluorescence intensities among conditions. For these analyses, p-values were adjusted for multiple comparisons using the Holm’s method. Significance levels and error bars are defined in each figure caption. Protein sequences and annotations for C. oleaginosum were retrieved from the respective Joint Genome Institute (JGI) database ( 43 , 151 ). Bottom-up proteomics data were processed with MaxQuant for label-free quantification ( 152 ). All log 2 foldchange (“FC”) data correspond to differential expression analyses using the DEqMS package with MaxQuant LFQ intensities, unless otherwise specified as using the DEP package with normalized raw intensity values ( 153 , 154 ). For proteomics differential expression analysis, p-values were adjusted using the the Benjamini–Hochberg procedure. Refer to SI Appendix , Supplementary Methods for details regarding RStudio Bioconductor packages and annotation prediction tools used for bioinformatics analyses ( 155 ). Declarations Competing Interest Statement The authors declare no competing interests. Author Contribution A.G. and B.Y. designed research; A.G., Y.P., D.H., and X.L. performed research; Y.P., D.H., X.C., A.J.R., T.Z., and W-J.Q. contributed new reagents/analytical tools; A.G., Y.P., D.H., and Z.J. analyzed data; A.G. wrote the original draft; B,Y, and W-J.Q assisted with conceptualization, supervision, and acquisition of experimental resources and project funding. all authors contributed to the writing and final edition of the final manuscript. Acknowledgement We acknowledge the U.S. Department of Energy (DOE) Office of Energy Efficiency & Renewable Energy (DE-EE0008250, and DE-EE0009763), the U.S. Department of Agriculture (USDA) National Institute of Food and Agriculture (Hatch/Multi State project 1017904), and the Bioproducts, Science and Engineering Laboratory, Department of Biological Systems Engineering at Washington State University (WSU). Portions of this research were also supported by the Predictive Phenomics Initiative (TZ) conducted under the Laboratory Directed Research and Development Program at Pacific Northwest National Laboratory (PNNL) in addition to a project award from the Environmental Molecular Sciences Laboratory, a DOE Office of Science User Facility sponsored by the Biological and Environmental Research program under Contract No. DE-AC05-76RL01830. YP and AJR acknowledge partial support from the Center for Bioenergy Innovation (CBI) at Oak Ridge National Laboratory, a U.S. DOE Bioenergy Research Center supported by the Office of Biological and Environmental Research in the DOE Office of Science. AG is grateful for support from the PNNL-WSU Distinguished Graduate Research Program Fellowship. We are also thankful for the FAMEs protocol from PNNL scientists Mark G. Butcher and Angela M. Melville. Data Availability The data supporting this study’s findings are available from the corresponding author (Drs. 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Supplementary Files 3.16.25SIAppendixNatureCommunicationsUsingAlkaliLignintoInvestigateLignin...docx SIAppendixDatasetS1SecretomicsResults.xlsx SIAppendixDatasetS2CellularProteomicsResults.xlsx SIAppendixDatasetS3BLASTpResults.xlsx Cite Share Download PDF Status: Published Journal Publication published 18 Jan, 2026 Read the published version in Scientific Reports → Version 1 posted Editorial decision: Revision requested 28 May, 2025 Reviews received at journal 27 May, 2025 Reviewers agreed at journal 13 May, 2025 Reviews received at journal 28 Apr, 2025 Reviewers agreed at journal 15 Apr, 2025 Reviewers invited by journal 15 Apr, 2025 Editor assigned by journal 09 Apr, 2025 Editor invited by journal 09 Apr, 2025 Submission checks completed at journal 07 Apr, 2025 First submitted to journal 29 Mar, 2025 You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. 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Also discoverable on Platform About Our Team In Review Editorial Policies Advisory Board Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {"props":{"pageProps":{"initialData":{"identity":"rs-6335743","acceptedTermsAndConditions":true,"allowDirectSubmit":false,"archivedVersions":[],"articleType":"Article","associatedPublications":[],"authors":[{"id":446786095,"identity":"b597d516-546b-4688-b18c-c1c35e8563ac","order_by":0,"name":"Austin Gluth","email":"","orcid":"","institution":"Washington State Univerity","correspondingAuthor":false,"prefix":"","firstName":"Austin","middleName":"","lastName":"Gluth","suffix":""},{"id":446786096,"identity":"7b582dc9-3df8-4589-b0e8-4db2731a08f7","order_by":1,"name":"Yunqiao Pu","email":"","orcid":"","institution":"ORNL","correspondingAuthor":false,"prefix":"","firstName":"Yunqiao","middleName":"","lastName":"Pu","suffix":""},{"id":446786097,"identity":"fe78c92d-577b-43c2-ad66-f23f9b7bfb2d","order_by":2,"name":"Dehong Hu","email":"","orcid":"","institution":"Pacific Northwest National Laboratory","correspondingAuthor":false,"prefix":"","firstName":"Dehong","middleName":"","lastName":"Hu","suffix":""},{"id":446786098,"identity":"859cecd3-c845-416d-aaa9-91255c3bdde7","order_by":3,"name":"Xiaowen Chen","email":"","orcid":"","institution":"National Renewable Energy Laboratory","correspondingAuthor":false,"prefix":"","firstName":"Xiaowen","middleName":"","lastName":"Chen","suffix":""},{"id":446786099,"identity":"c40fd823-0354-4943-a45a-b574eed79091","order_by":4,"name":"Zachary Johnson","email":"","orcid":"","institution":"Washington State University","correspondingAuthor":false,"prefix":"","firstName":"Zachary","middleName":"","lastName":"Johnson","suffix":""},{"id":446786100,"identity":"af5b07ab-2442-4cdb-98f0-0a63e56d1c6a","order_by":5,"name":"Xiaolu Li","email":"","orcid":"","institution":"Pacific Northwest National Laboratory","correspondingAuthor":false,"prefix":"","firstName":"Xiaolu","middleName":"","lastName":"Li","suffix":""},{"id":446786101,"identity":"fcfc8b38-ea68-49ec-9b67-2280c3c2c656","order_by":6,"name":"Arthur Ragauskas","email":"","orcid":"","institution":"University of Tennessee","correspondingAuthor":false,"prefix":"","firstName":"Arthur","middleName":"","lastName":"Ragauskas","suffix":""},{"id":446786102,"identity":"d462f603-1ced-4392-a0db-412a4a6f4ba8","order_by":7,"name":"Wei-Jun Qian","email":"","orcid":"","institution":"Pacific Northwest National Lab","correspondingAuthor":false,"prefix":"","firstName":"Wei-Jun","middleName":"","lastName":"Qian","suffix":""},{"id":446786103,"identity":"a403dc4d-4be5-4403-b636-35867ed0a0ff","order_by":8,"name":"Tong Zhang","email":"","orcid":"","institution":"Pacific Northwest National Laboratory","correspondingAuthor":false,"prefix":"","firstName":"Tong","middleName":"","lastName":"Zhang","suffix":""},{"id":446786094,"identity":"bd0b26fd-4552-4918-b57b-d718f8783a7f","order_by":9,"name":"Bin Yang","email":"data:image/png;base64,iVBORw0KGgoAAAANSUhEUgAAAZAAAAAyAQMAAABI0h/eAAAABlBMVEX///8AAABVwtN+AAAACXBIWXMAAA7EAAAOxAGVKw4bAAAAw0lEQVRIiWNgGAWjYBACA2Yogx9CMeNUialFsoFoLXDGAWK1mLPzHmDmqbljt/lG+jMJhgrrxAZCWiyb+RKYeY49S952I8dMguFMOmEtBod5DJh52A4nm93OYZNgbDtMrJZ/h5ONZwMdxviPWC28bYftDKQTzCQYG4jQYtnMY3Bwbt/hBIn7b4wtEo6lGxPUYs5/xvDBm2+H7fl7jj+88aHGWpagFhA4xMPAAHFPAjHKQYDxBwODPbGKR8EoGAWjYAQCAJRgO57ICm81AAAAAElFTkSuQmCC","orcid":"","institution":"Washington State University","correspondingAuthor":true,"prefix":"","firstName":"Bin","middleName":"","lastName":"Yang","suffix":""}],"badges":[],"createdAt":"2025-03-29 19:53:14","currentVersionCode":1,"declarations":"","doi":"10.21203/rs.3.rs-6335743/v1","doiUrl":"https://doi.org/10.21203/rs.3.rs-6335743/v1","draftVersion":[],"editorialEvents":[{"content":"https://doi.org/10.1038/s41598-026-36483-5","type":"published","date":"2026-01-18T16:30:04+00:00"}],"editorialNote":"","failedWorkflow":false,"files":[{"id":82137130,"identity":"278ab1c0-7112-41be-b904-082f3bc08179","added_by":"auto","created_at":"2025-05-07 06:16:45","extension":"png","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":866275,"visible":true,"origin":"","legend":"\u003cp\u003eSummary of the approach used in this study. First, alkali lignin was extracted from dilute alkali pretreated corn stover black liquor. Then, \u003cem\u003eC. oleaginosum\u003c/em\u003e was cultivated in a YNB-based medium containing either 1 g/L of the extracted lignin, glucose, benzoate, or no additional carbon source. A variety of assays and analytical tools were used to collect extracellular and (intra)cellular metrics for growth, lipid content, lignin modification, and lignin/aromatics transport. Finally, a label-free SP3 proteomics workflow was employed to discover putative reactive oxygen species (ROS)-mediated mechanisms and pathways for lignin and aromatics degradation.\u003c/p\u003e","description":"","filename":"image1.png","url":"https://assets-eu.researchsquare.com/files/rs-6335743/v1/543e67b2e64929602bda7437.png"},{"id":82135425,"identity":"16861b74-3f3d-4797-aa78-3b5f9040f4e4","added_by":"auto","created_at":"2025-05-07 06:08:43","extension":"png","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":1117243,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cem\u003eC. oleaginosum \u003c/em\u003ecan degrade lignin but cannot efficiently utilize it for growth. (A) Bar chart of cell dry weights (CDW) from cultivation on alkali-purified lignin. (B) Box plot showing the % decrease in UV absorbance (320 nm) of culture supernatants from the lignin condition. The 0 hour time point designates the absorbance median and range (variation) for the lignin medium blank. (C) Bar chart presenting the % decrease in absorbance at 96 hours according to the Prussian blue assay. (D) CDWs after 96 hours of cultivation on other carbon sources. (E) Line plot of viable cell counts over time for the lignin and “No Carbon” conditions. (F) Line plot of % cell viability over time according to trypan blue staining. Colored regression lines are included with gray shading that delineates 95% confidence intervals. Results shown in panels A–D are derived from biological triplicates, and the error bars represent one standard deviation. Significance levels from two-tailed t-tests are included as asterisks in the bar charts and box plots. One asterisk is defined as a p-value of 0.05, whereas two is 0.005 and so on. No significant difference is “ns”.\u003c/p\u003e","description":"","filename":"image2.png","url":"https://assets-eu.researchsquare.com/files/rs-6335743/v1/1434b185be6892d207e7bc80.png"},{"id":82135422,"identity":"60c1c2f6-27fa-4cb1-b5fc-e3f7f1c1c86b","added_by":"auto","created_at":"2025-05-07 06:08:42","extension":"png","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":664134,"visible":true,"origin":"","legend":"\u003cp\u003eNMR results pinpoint lignin structural modifications due to yeast cultivation. (A) Aromatics region of 2D-HSQC-NMR spectra for control lignin and lyophilized supernatant from yeast cultivation with lignin. Abbreviations and the corresponding features are defined below the spectra. (B) Aliphatic region of 2D-HSQC-NMR spectra. Note that the five grey dots adjacent to Bγ designate xylan. Dashed black box inlay designates newly formed peaks likely due to cleavage of bonds between lignin subunits. (C) Summary of semi-quantitative changes to interunit lignin bonds. Note that the increase in β-O-4 can be explained by a relative decrease in other bonds as well as lignin motifs shown in panels A and B.\u003c/p\u003e","description":"","filename":"image3.png","url":"https://assets-eu.researchsquare.com/files/rs-6335743/v1/5b7edd15c53d0ecb82ffdb53.png"},{"id":82137125,"identity":"8b362e2b-2c87-49c3-9b36-332095225304","added_by":"auto","created_at":"2025-05-07 06:16:43","extension":"png","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":1912401,"visible":true,"origin":"","legend":"\u003cp\u003eHigh-resolution microscopy reveals lignin-dependent yeast cell fluorescence and morphological differences. (A) Fluorescence microscopy results showing lignin autofluorescence and subtle differences between lignin and benzoate cultures. (B) Fluorescence intensity distributions for cell populations recorded over time. (C) Airyscan images show difference in fluorescence of the cellular envelope and putative mitochondrial clusters. (D) Airyscan images of yeast labeled with MitoTracker Red CMXRos Dye. 5 µm scale bars are shown in the bottom right corners for Panel D. Differences in yeast fluorescence intensities may be attributed to mitochondrial autofluorescence, lignin interactions, and/or carbon-limited oxidative stress (48). For panel B, cell population fluorescence intensities for three images were averaged then these three mean values were used for t-tests. Significance levels from two-tailed t-tests are included as asterisks in the box plots. One asterisk is a p-value of 0.05. No significant difference is “ns”.\u003c/p\u003e","description":"","filename":"image4.png","url":"https://assets-eu.researchsquare.com/files/rs-6335743/v1/3ebc7bdb42a59f604bf49f58.png"},{"id":82135435,"identity":"3c7a5785-92a1-4686-8f2c-9ea54b4e6b1a","added_by":"auto","created_at":"2025-05-07 06:08:44","extension":"png","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":861464,"visible":true,"origin":"","legend":"\u003cp\u003eDistinct protein expression patterns in lignin and glucose conditions uncovered by secretomics and proteomics. (A) Venn diagrams indicating overlap of unique, quantifiable protein IDs. Only one missing LFQ intensity value was permitted among the replicates of each condition. Secretomics results were filtered using a cutoff of 0.85 according to SignalP 6.0 (46). (B) Correlogram of Pearson correlations for overlapping LFQ results from secretomics data. (C) Correlogram of Pearson correlations for proteomics data. (D) Donut charts of CUPP-annotated CAZymes found in our secretomics and proteomics data (47). “Observed” refers to all unique identifications. “DE-filtered” refers to IDs for which two LFQ values were present to conduct differential expression analysis. (E) Volcano plots summarizing FCs from separate differential expression analyses of secretomics and proteomics data. The vertical purple lines designate FCs of −1 and +1, whereas the dashed black line designates an adjusted p-value of 0.05. (F) Summarized dot plot results from KEGG pathway enrichment analysis using protein IDs with significant differences (adjusted p-value ≤ 0.05) in expression: “Down” refers to those with FC ≤ −1, and “Up” refers to FC ≥ 1.\u003c/p\u003e","description":"","filename":"image5.png","url":"https://assets-eu.researchsquare.com/files/rs-6335743/v1/73cc4ea9efeb1ecb128d2365.png"},{"id":82135429,"identity":"ec88723f-1207-4e58-bead-d7ce0e10967c","added_by":"auto","created_at":"2025-05-07 06:08:43","extension":"png","order_by":6,"title":"Figure 6","display":"","copyAsset":false,"role":"figure","size":1601125,"visible":true,"origin":"","legend":"\u003cp\u003eProposed pathways for lignin modification and catabolism of lignin-derived aromatics. According to our NMR results, extensive homology searches, and literature review, pathways are only shown for H- and G-lignin-derived aromatics. Additional details regarding ROS-mediated mechanisms for lignin modification are provided in the \u003cem\u003eSI Appendix\u003c/em\u003e, Fig. S4. The predicted pathways for G-lignin derivatives are discussed in \u003cem\u003eSI Appendix\u003c/em\u003e, Supporting Text. JGI protein IDs are included within boxes that are colored according to the average FC for lignin vs. glucose. Note that boldface green lettering corresponds to proteins only observed in the lignin condition. Boldface blue lettering corresponds to proteins that were observed, but for which no differential expression analysis could be conducted due to missing intensity values. Boldface purple lettering corresponds to IDs for which raw intensity values were used in DEP differential expression analysis due to missing LFQ intensity values. Question marks and dotted arrows represent uncertain annotations. The numbers in circles designate the following putative enzyme annotations: 1. Laccases, 2. Peroxidases, 3. Phenol 2-monooxygenase, 4. Catechol 1,2-dioxygenase, 5. Muconate cycloisomerase, 6. Muconolactone isomerase, 7. β-ketoadipate enol-lactone hydrolase, 8. β-ketoadipate-succinyl-CoA transferase, 9. β-ketoadipyl-CoA thiolase, 10. Benzoate 4-monooxygenase (Cytochrome P450), 11. \u003cem\u003ep\u003c/em\u003e-Coumarate-CoA ligase, 12. Cinnamoyl-CoA hydratase/dehydrogenase (first arrow = IDs on the left) and 3-ketoacyl CoA thiolase (second arrow = IDs on the right), 13. Thioesterase, 14. Hydroxylase, 15. Protocatechuate 3,4-dioxygenase, 16. β-carboxymuconate lactonizing enzyme, 17. γ-carboxymuconolactone decarboxylase, 18. Monooxygenase/decarboxylase, 19. Hydroxyquinol 1,2-dioxygenase, 20. Maleylacetate reductase, 21. Cinnamoyl-CoA reductase, 22. Aldehyde dehydrogenase/cytochrome P450 (see IDs below coniferyl aldehyde structure), 23. Feruloyl-CoA synthetase, 24. Cinnamoyl-CoA hydratase/dehydrogenase (first arrow = IDs on the left) and 3-ketoacyl CoA thiolase (second arrow = IDs on the right), 25. Thioesterase, 26. Vanillin dehydrogenase, 27. Vanillyl alcohol oxidase, 28. Vanillate hydroxylase, 29. O-demethylase, 30. (Di)methoxyhydroquinone 1,2-dioxygenase, 31. \u003cem\u003ep\u003c/em\u003e-Hydroxybenzoate decarboxylase, 32. GMC oxidoreductase, 33. Gluconate kinase.\u003c/p\u003e","description":"","filename":"image6.png","url":"https://assets-eu.researchsquare.com/files/rs-6335743/v1/6e125537a8d307540857aef5.png"},{"id":100614648,"identity":"5e6ad5c8-c429-421a-9c04-14880268486e","added_by":"auto","created_at":"2026-01-19 17:22:37","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":7968513,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-6335743/v1/3de0a2d7-75f7-41f8-8d66-491af8ce7eb3.pdf"},{"id":82135423,"identity":"f94d71fa-7785-450d-a658-0d477b271734","added_by":"auto","created_at":"2025-05-07 06:08:42","extension":"docx","order_by":1,"title":"","display":"","copyAsset":false,"role":"supplement","size":15348268,"visible":true,"origin":"","legend":"","description":"","filename":"3.16.25SIAppendixNatureCommunicationsUsingAlkaliLignintoInvestigateLignin...docx","url":"https://assets-eu.researchsquare.com/files/rs-6335743/v1/eef42544a7ffc317eeaf6c50.docx"},{"id":82135426,"identity":"8deecdb4-f087-4e29-83f3-f24167a44b42","added_by":"auto","created_at":"2025-05-07 06:08:43","extension":"xlsx","order_by":2,"title":"","display":"","copyAsset":false,"role":"supplement","size":4076535,"visible":true,"origin":"","legend":"","description":"","filename":"SIAppendixDatasetS1SecretomicsResults.xlsx","url":"https://assets-eu.researchsquare.com/files/rs-6335743/v1/23c88eca8b87a9de0783b7f0.xlsx"},{"id":82135421,"identity":"7a40a39b-389a-46eb-bafb-1e51bf8e9d5c","added_by":"auto","created_at":"2025-05-07 06:08:42","extension":"xlsx","order_by":3,"title":"","display":"","copyAsset":false,"role":"supplement","size":3994413,"visible":true,"origin":"","legend":"","description":"","filename":"SIAppendixDatasetS2CellularProteomicsResults.xlsx","url":"https://assets-eu.researchsquare.com/files/rs-6335743/v1/933893b2203f929c7f1ec45d.xlsx"},{"id":82135476,"identity":"0fe5a4fb-0dc4-4e0a-a033-d2b010e0354d","added_by":"auto","created_at":"2025-05-07 06:08:45","extension":"xlsx","order_by":4,"title":"","display":"","copyAsset":false,"role":"supplement","size":49701,"visible":true,"origin":"","legend":"","description":"","filename":"SIAppendixDatasetS3BLASTpResults.xlsx","url":"https://assets-eu.researchsquare.com/files/rs-6335743/v1/025b2a6cfeb53daa7cd96a5f.xlsx"}],"financialInterests":"No competing interests reported.","formattedTitle":"Exploring the Ligninolytic Capabilities of the Oleaginous Yeast Cutaneotrichosporon oleaginosum","fulltext":[{"header":"Introduction","content":"\u003cp\u003eLignin constitutes a complex and diverse biopolymer, and, importantly, it is the largest renewable source of aromatics. As much as 40% of the energy density of lignocellulosic biomass is found in lignin, and it has a higher carbon content than carbohydrates: 68 wt% for the lignin derivative guaiacyl vs. 44% for cellulose based on glucose (\u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e). Bacteria and filamentous fungi are prime candidates for leveraging this rich carbon source given their ligninolytic capabilities (\u003cspan additionalcitationids=\"CR3 CR4\" citationid=\"CR2\" class=\"CitationRef\"\u003e2\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e). In general these microorganisms degrade lignin using oxidoreductases such as laccases and peroxidases as well as other redox-active molecules (\u003cspan additionalcitationids=\"CR7\" citationid=\"CR6\" class=\"CitationRef\"\u003e6\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR8\" class=\"CitationRef\"\u003e8\u003c/span\u003e). Compared to filamentous fungi, bacteria such as \u003cem\u003ePseudomonas putida\u003c/em\u003e have higher growth rates and are amenable to genetic manipulation; nevertheless, white rot fungi like \u003cem\u003ePhanerchaete chrysosporium\u003c/em\u003e are superior lignin degraders (\u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e9\u003c/span\u003e). In contrast the potential of yeasts (particularly basidiomycetes) as biocatalysts for lignin valorization and their roles in lignin-containing microbial niches have rarely been investigated. As a result, a clarity regarding their ability to degrade (or at least modify) lignin and what enzymes may functionally confer these capabilities is lacking (\u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eThough scarce, there are reports of basidiomycete yeasts (and one ascomycete) with activities against lignin as the major carbon source. These include species of the genera \u003cem\u003eRhodotorula\u003c/em\u003e, \u003cem\u003eGeotrichum\u003c/em\u003e, \u003cem\u003eTausonia\u003c/em\u003e, and \u003cem\u003eCutaneotrichosporon\u003c/em\u003e (\u003cspan additionalcitationids=\"CR12 CR13\" citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e). Laccase, manganese peroxidase, and lignin peroxidase activities have been observed in yeasts\u0026mdash;some of which display the oleaginous phenotype under nutrient-limiting conditions (\u003cspan additionalcitationids=\"CR15 CR16\" citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR17\" class=\"CitationRef\"\u003e17\u003c/span\u003e). Oleaginous yeast accumulate lipids and are promising candidates for sustainable synthesis of palm oil substitutes, lubricants, plastics, and fuels (\u003cspan additionalcitationids=\"CR19 CR20\" citationid=\"CR18\" class=\"CitationRef\"\u003e18\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e). Of these yeasts, \u003cem\u003eCutaneotrichosporon oleaginosum\u003c/em\u003e (formerly known as \u003cem\u003eCryptococcus curvatus\u003c/em\u003e and later \u003cem\u003eTrichosporon\u003c/em\u003e oleaginosus) is poised for lignin valorization because it can accumulate lipids using the monoaromatic resorcinol (\u003cspan additionalcitationids=\"CR23\" citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e). However, it is unclear if this yeast can degrade lignin and what enzymatic mechanisms are involved both extra- and intracellularly. A recent functional genomics analysis of \u003cem\u003eC. oleaginosum\u003c/em\u003e cultivated on alkali-pretreated corn stover has expanded our knowledge of yeast aromatics catabolism but again raises questions about its ligninolytic abilities due to ambiguous results from K-lignin analysis, NMR spectroscopy, and enzymatic activity assays (\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e, \u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e). For instance, a decrease in acetyl groups was observed whereas no changes to the (hemi)cellulose fractions were reported raising the possibility that the lignin in the black liquor was blocked (\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eThe hydrolysate derived from dilute-alkali corn stover pretreatment contains numerous accessible carbon sources apart from lignin: monomeric sugars, oligosaccharides, organic acids, and monoaromatics (\u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e, \u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e). Other studies have used modified (Kraft) lignin in the presence of glucose, which obfuscates secretomics results (\u003cspan citationid=\"CR28\" class=\"CitationRef\"\u003e28\u003c/span\u003e). We cultivated \u003cem\u003eC. oleaginosum\u003c/em\u003e on lignin purified from alkali-pretreated corn stover to directly evaluate lignin degradation/modification (\u003cspan citationid=\"CR29\" class=\"CitationRef\"\u003e29\u003c/span\u003e). We hypothesize that the proposed yeast secretes ligninolytic enzymes during carbon limitation, and that liberated aromatics and polymeric lignin are proximally transported into the cell. Our 2D-NMR results demonstrate decreases in the H- and G-lignin motifs and relative changes to bonding in the lignin structure. Our fluorescence microscopy results suggest cellular uptake of polymeric lignin. Passive and active transport of lignin-derived aromatics has been studied, but there is a dearth of information regarding lignin fragments (\u003cspan additionalcitationids=\"CR31 CR32\" citationid=\"CR30\" class=\"CitationRef\"\u003e30\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR33\" class=\"CitationRef\"\u003e33\u003c/span\u003e). \u003cem\u003eSaccharomyces cerevisiae\u003c/em\u003e can uptake nanoparticles of polyaromatic polystyrene, which indirectly supports yeast lignin uptake (\u003cspan citationid=\"CR34\" class=\"CitationRef\"\u003e34\u003c/span\u003e). Our fluorescence microscopy approach which harnesses the native fluorescent properties of lignin corroborates this (\u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e35\u003c/span\u003e, \u003cspan citationid=\"CR36\" class=\"CitationRef\"\u003e36\u003c/span\u003e). Finally, we developed a comprehensive secretomics and proteomics workflow to study the expressed metabolism of \u003cem\u003eC. oleaginosum\u003c/em\u003e, thereby pinpointing putative enzymes for lignin modification as well as transport and utilization of lignin-derived aromatics. This comprehensive work provides targets for functional evaluation and additional impetus for developing basidiomycete yeasts as chassis for lignin valorization.\u003c/p\u003e"},{"header":"Results and Discussion","content":"\u003cdiv id=\"Sec3\" class=\"Section2\"\u003e \u003ch2\u003eCultivation on Alkali Lignin is Reminiscent of the \u0026ldquo;No Carbon\u0026rdquo; Condition\u003c/h2\u003e \u003cp\u003eA multifaceted approach was employed to study the cultivation of \u003cem\u003eC. oleaginosum\u003c/em\u003e on alkali lignin and is summarized in Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003e. Lignin from dilute alkali-pretreated corn stover was extracted with successive acid-base titrations and analyzed via the K-Lignin assay. The overall lignin component was 87.06% with a low ash content of 0.30% (\u003cem\u003eSI Appendix\u003c/em\u003e, Table \u003cspan refid=\"MOESM1\" class=\"InternalRef\"\u003eS1\u003c/span\u003e). Though the isolated lignin contained structural saccharides, the high lignin and low ash contents are ideal for this work. Four conditions were evaluated in this study: 1 g/L lignin, 1 g/L glucose, 1 g/L benzoate, and a \u0026ldquo;No Carbon\u0026rdquo; control in which no additional carbon source was added to the base medium. The \u0026ldquo;No Carbon\u0026rdquo; control was included to uncover differences specifically attributed to interactions with lignin. An additional control of uninoculated medium with 1 g/L lignin was included to check for contamination and as a baseline comparison for absorbance assays. Additional analyzes of growth, lignin modification, and lignin uptake were conducted to lay a foundation for studying the extracellular and cellular proteomes of \u003cem\u003eC. oleaginosum\u003c/em\u003e.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eAfter 144 hours of cultivation, a decrease in cell dry weight (CDW) was observed for the lignin condition (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eA). Despite this, 8.5% and 11.3% decreases in lignin content were observed at 96 and 144 hours, respectively, according to UV absorbance at 320 nm (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eB). These results were confirmed using the Prussian Blue assay (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eC). The 96-hour time point was chosen to compare the CDWs of different carbon-limited conditions. Unexpectedly, a slight albeit significant difference between the \u0026ldquo;No Carbon\u0026rdquo; and lignin conditions was recorded and was supported by an increase in viable cell counts with minimal differences in cell viability (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eD\u0026ndash;F). It\u0026rsquo;s possible that the observed difference in growth is due to utilization of structural saccharides and/or acetyl groups, which constitute\u0026thinsp;~\u0026thinsp;13% of the alkali lignin (\u003cem\u003eSI Appendix\u003c/em\u003e, Table \u003cspan refid=\"MOESM1\" class=\"InternalRef\"\u003eS1\u003c/span\u003e). At 96 hours, lipid titers and fatty acid compositions for the \u0026ldquo;No Carbon\u0026rdquo; and lignin conditions were similar (\u003cem\u003eSI Appendix\u003c/em\u003e, Fig. \u003cspan refid=\"MOESM1\" class=\"InternalRef\"\u003eS1\u003c/span\u003e). Stearic acid (C18:0) was not observed in these two conditions, and a general increase in fatty acid desaturation was apparent, indicating that lipids were mobilized in the \u0026ldquo;No Carbon\u0026rdquo; and lignin conditions for carbon and energy generation (\u003cspan citationid=\"CR37\" class=\"CitationRef\"\u003e37\u003c/span\u003e, \u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e). Overall, increased growth in the lignin condition vs. the \u0026ldquo;No Carbon\u0026rdquo; condition was unexpected because of the sheer recalcitrance of lignin.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eNMR Results Demonstrate Selective Modification of Alkali Lignin\u003c/h3\u003e\n\u003cp\u003eTwo-dimensional (2D) heteronuclear single-quantum correlation (HSQC) NMR spectroscopy was used to study the effects of \u003cem\u003eC. oleaginosum\u003c/em\u003e on lignin\u0026rsquo;s chemical structures (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eA\u0026ndash;B). HSQC provides a semi-quantitative analysis of lignin\u0026rsquo;s structural features such as monolignol composition and interunit linkages. Lignin recovered from the lyophilized yeast culture supernatant was compared to the control alkali lignin. NMR analysis revealed changes to lignin structures after yeast cultivation. The control lignin had a S/G/H ratio of 57:31:12, whereas after yeast fermentation, the lignin in the supernatant had a ratio of 66:29:5. Compared to the control, this is approximately a 59.8% decrease in the H-lignin unit, which is comprised of the \u003cem\u003ep\u003c/em\u003e-coumaryl alcohol monomer. This suggests that the yeast preferentially utilizes un-methylated aromatics, or that \u003cem\u003ep\u003c/em\u003e-coumarate, \u003cem\u003ep\u003c/em\u003e-hydroxybenzoate, and \u003cem\u003ep\u003c/em\u003e-hydroxyphenol pendents are more susceptible to liberation. Predominant release of these H-lignin derived aromatics was reported recently and lends some support to the latter (\u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e). It follows that we observed a 16.5% decrease in \u003cem\u003ep\u003c/em\u003e-coumarate esters and a 13.4% decrease is ferulate esters as previously reported (\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e). We did not observe a significant decrease in the S/G (syringyl/guaiacyl) ratio, whereas a minor potentially insignificant decrease was observed previously using alkali-pretreated corn stover (\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e). Given the decrease in total aromatics (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eB\u0026ndash;C), it\u0026rsquo;s possible that a simulatenous decrease in S/G motifs occurred.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eThe aliphatic region showed a 75.8% and 60.5% depletion of resinol (β-β) and phenylcoumaran β-5 linkages, respectively (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eB\u0026ndash;C). Interestingly, there was a slight increase of the overall β-O-4 linkage signal. Among the various types of β-O-4 linkages, a small decrease in G/H-type was observed, but the S-type β-O-4 linkage dominated. The decrease of G/H-type β-O-4 could be attributed in part to the decrease of H-type β-O-4 linkage, consistent with the decrease of H unit as observed from aromatic regions of the HSQC spectra. Previous 2D-NMR results of this yeast grown on alkali-pretreated corn stover demonstrated no decrease in β-O-4; however, the high presence of contaminating polysaccharides complicates interpretation (\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e). The two new cross peaks at δ\u003csub\u003eC\u003c/sub\u003e/δ\u003csub\u003eH\u003c/sub\u003e 68.2/3.91 and δ\u003csub\u003eC\u003c/sub\u003e/δ\u003csub\u003eH\u003c/sub\u003e 72.2/3.48 ppm in the aliphatic region correspond to Ar-CH(OH)-O-CH\u003csub\u003e2\u003c/sub\u003e-COOH. In studies of another basidiomycete yeast as well as a white rot fungus cultivated on lignin, 1D-NMR results demonstrated that these newly formed peaks arise from the cleavage of bonds between lignin subunits (\u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e, \u003cspan citationid=\"CR40\" class=\"CitationRef\"\u003e40\u003c/span\u003e). Compared to those and other studies, a 96-hour time point caused apparent changes of lignin structures, but longer cultivation time may be needed to observe more lignin degradation (\u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e, \u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e, \u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e, \u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e41\u003c/span\u003e). However, a decrease in ligninolytic activity during prolonged cultivation has been reported, and our results directly demonstrate the ligninolytic capacity of \u003cem\u003eC. oleaginosum\u003c/em\u003e (\u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e42\u003c/span\u003e).\u003c/p\u003e\n\u003ch3\u003eSecretomics and Proteomics Reveal Stark Contrasts between Carbon-limited Conditions\u003c/h3\u003e\n\u003cp\u003eTeasing out differences in protein expression due to the presence of lignin is crucial for discovering ligninolytic oxidoreductases and related enzymes. The single-pot, solid-phase-enhanced sample preparation (SP3) approach facilitated high protein digestion efficiencies (\u0026lt;\u0026thinsp;10% missed cleavage) and a higher proteome coverage per sample (\u0026gt;\u0026thinsp;43% total coverage of the modeled genome) compared to recent studies of \u003cem\u003eC. oleaginosum\u003c/em\u003e (\u003cem\u003eSI Appendix\u003c/em\u003e, Supporting Text and Table \u003cspan refid=\"MOESM4\" class=\"InternalRef\"\u003eS4\u003c/span\u003e) (\u003cspan additionalcitationids=\"CR44\" citationid=\"CR43\" class=\"CitationRef\"\u003e43\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR45\" class=\"CitationRef\"\u003e45\u003c/span\u003e). In the secretomics results, 245 unique proteins passed the SignalP signal peptide prediction score of 0.85, and 14 of these were only observed in the lignin condition (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e5\u003c/span\u003eA and \u003cem\u003eSI Appendix\u003c/em\u003e, Dataset S1) (\u003cspan citationid=\"CR46\" class=\"CitationRef\"\u003e46\u003c/span\u003e). 3,579 unique proteins were observed in the proteomics results with 56 of those only observed in the lignin condition (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e5\u003c/span\u003eA and \u003cem\u003eSI Appendix\u003c/em\u003e, Dataset S2). The Pearson correlation coefficients for lignin vs. glucose replicates were ~\u0026thinsp;0.60 and ~\u0026thinsp;0.90 for the secretomics and proteomics results, respectively (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e5\u003c/span\u003eB\u0026ndash;C). Proteins unique to the lignin condition were also observed for the lignin vs. \u0026ldquo;No Carbon\u0026rdquo;, along with a lower Pearson correlation coefficient (~\u0026thinsp;0.80) for the secretomics data (\u003cem\u003eSI Appendix\u003c/em\u003e, Fig. \u003cspan refid=\"MOESM2\" class=\"InternalRef\"\u003eS2\u003c/span\u003eA\u0026ndash;B). For cellular proteomics results, correlation between lignin vs. \u0026ldquo;No Carbon\u0026rdquo; samples was high (~\u0026thinsp;0.97), which is expected due to carbon starvation (\u003cem\u003eSI Appendix\u003c/em\u003e, Fig. \u003cspan refid=\"MOESM2\" class=\"InternalRef\"\u003eS2\u003c/span\u003eC). Nevertheless, principle component analyses show a clear separation for each condition (\u003cem\u003eSI Appendix\u003c/em\u003e, Fig. \u003cspan refid=\"MOESM3\" class=\"InternalRef\"\u003eS3\u003c/span\u003e).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eCarbohydrate active enzyme (CAZymes) and differential expression results are summarized in Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e5\u003c/span\u003eD\u0026ndash;E. CAZy annotations from the Conserved Unique Peptide Patterns (CUPP) platform were used because of the protein-grouping capabilities of the CUPP algorithm, which can help distinguish certain functional annotations beyond CAZy subfamilies (\u003cspan citationid=\"CR47\" class=\"CitationRef\"\u003e47\u003c/span\u003e). 42 CAZymes were observed in the secretome, while 51 were observed intracellularly (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e5\u003c/span\u003eD). For both the secretomics and cellular proteomics datasets, relative quantification was performed using LFQ intensities (\u0026ldquo;DE-filtered\u0026rdquo;, Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e5\u003c/span\u003eD). In the lignin vs. glucose secretomics data, 45 proteins had log\u003csub\u003e2\u003c/sub\u003e fold changes (\u0026ldquo;FC\u0026rdquo;)\u0026thinsp;\u0026gt;\u0026thinsp;1 and adjusted p-values\u0026thinsp;\u0026lt;\u0026thinsp;0.05, while 44 were significantly downregulated (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e5\u003c/span\u003eE). In the cellular proteomics data, 207 proteins were upregulated, and 130 were downregulated. To provide an overview, pathway enrichment was conducted (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e5\u003c/span\u003eF). It\u0026rsquo;s evident that an expression regime was upregulated to access diverse carbon sources (e.g., galactose, sucrose, flavonoids/aromatics, glycans, etc.) during carbon limitation. Upregulation of N-glycan synthesis suggests restructuring of the cell wall perhaps in response to stress. Lastly, the prevalence of amino acid and other carbon recycling pathways (e.g., fatty acid degradation) was expected; however, it is intriguing to see downregulation of glutathione and cysteine/methionine metabolic pathways. This may highlight the importance of antioxidants during carbon limitation (\u003cspan citationid=\"CR48\" class=\"CitationRef\"\u003e48\u003c/span\u003e) and raises questions about the toxicity of lignin/aromatics and byproducts of lignin breakdown. With deep coverage proteomics results, we projected an extracellular mechanism for yeast lignin modification (SI Appendix, Fig. \u003cspan refid=\"MOESM4\" class=\"InternalRef\"\u003eS4\u003c/span\u003e) and routes for funneling aromatics to central metabolism (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e6\u003c/span\u003e) in the following subsections.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003e \u003cb\u003eC. oleaginosum\u003c/b\u003e \u003cb\u003eExpresses Oxidoreductases for Extracellular Quinone Redox Cycling\u003c/b\u003e\u003c/p\u003e \u003cp\u003eLignin degradation is an oxidative extracellular process involving the concerted effort of reactive oxygen species (ROS), Fenton chemistry, and various enzymes (\u003cspan citationid=\"CR49\" class=\"CitationRef\"\u003e49\u003c/span\u003e). This complex process requires extracellular production of ROS and aromatic radicals including hydrogen peroxide (H\u003csub\u003e2\u003c/sub\u003eO\u003csub\u003e2\u003c/sub\u003e), hydroxyl radical (\u003csup\u003e\u0026minus;\u0026bull;\u003c/sup\u003eOH), superoxide (O\u003csub\u003e2\u003c/sub\u003e\u003csup\u003e\u0026minus;\u0026bull;\u003c/sup\u003e), cation radicals, and phenoxy radicals (\u003cspan citationid=\"CR50\" class=\"CitationRef\"\u003e50\u003c/span\u003e). These highly reactive species can be harnessed by electron-transferring auxiliary activity (AA) family oxidoreductases to attack lignocellulose (\u003cspan citationid=\"CR51\" class=\"CitationRef\"\u003e51\u003c/span\u003e). In addition to a repertoire of laccases (AA1) and peroxidases (AA2), fungi also employ non-enzymatic mechanisms to modify lignin (\u003cspan additionalcitationids=\"CR53 CR54 CR55\" citationid=\"CR52\" class=\"CitationRef\"\u003e52\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR56\" class=\"CitationRef\"\u003e56\u003c/span\u003e). At the heart of this mechanism is the hydroquinone-quinone redox cycle that reduces Fe\u003csup\u003e3+\u003c/sup\u003e to ultimately produce H\u003csub\u003e2\u003c/sub\u003eO\u003csub\u003e2\u003c/sub\u003e for initiating enzymatic catalysis, and \u003csup\u003e\u0026minus;\u0026bull;\u003c/sup\u003eOH to react directly with lignin units (\u003cspan citationid=\"CR49\" class=\"CitationRef\"\u003e49\u003c/span\u003e, \u003cspan citationid=\"CR50\" class=\"CitationRef\"\u003e50\u003c/span\u003e). Interestingly, whether or not basidiomycete yeasts have the functional capacity for quinone redox cycling has yet to be addressed according to Web of Science and PubMed searches using variations of the following wildcard search queries: ((yeast) AND (hydroquinone redox cycl* OR quinone redox cycl* OR Fenton chemistry)) AND (lignin OR aromatic*).\u003c/p\u003e \u003cp\u003eThere are several requirements for the hydroquinone redox cycle: oxidoreductases, reductants, and an acidic pH (\u003cspan citationid=\"CR50\" class=\"CitationRef\"\u003e50\u003c/span\u003e, \u003cspan citationid=\"CR57\" class=\"CitationRef\"\u003e57\u003c/span\u003e). A combination of BLASTp searches and CAZy CUPP annotations were used to find candidates in our proteomics data (\u003cem\u003eSI Appendix\u003c/em\u003e, Fig. \u003cspan refid=\"MOESM4\" class=\"InternalRef\"\u003eS4\u003c/span\u003e and Dataset S3) (\u003cspan citationid=\"CR47\" class=\"CitationRef\"\u003e47\u003c/span\u003e, \u003cspan citationid=\"CR51\" class=\"CitationRef\"\u003e51\u003c/span\u003e, \u003cspan citationid=\"CR58\" class=\"CitationRef\"\u003e58\u003c/span\u003e). Ferric reductases are important for iron uptake (generally as Fe\u003csup\u003e2+\u003c/sup\u003e) and maintaining an active quinone redox cycle (\u003cspan citationid=\"CR59\" class=\"CitationRef\"\u003e59\u003c/span\u003e, \u003cspan citationid=\"CR60\" class=\"CitationRef\"\u003e60\u003c/span\u003e). Unfortunately, none of the ferric reductases annotated in the Joint Genome Institute (JGI) database were observed intracellularly; however, they were observed in the secretome despite being localized to the cell membrane (\u003cspan citationid=\"CR59\" class=\"CitationRef\"\u003e59\u003c/span\u003e, \u003cspan citationid=\"CR61\" class=\"CitationRef\"\u003e61\u003c/span\u003e). These include Co_379781, Co_368607, and Co_366398 (the latter only observed in lignin replicates), which share 35\u0026ndash;40% identity with a ferric reductase (Pospl1_130030) from the model brown rot fungus \u003cem\u003eRhodonia placenta\u003c/em\u003e MAD 698R (\u003cem\u003eSI Appendix\u003c/em\u003e, Dataset S3) (\u003cspan citationid=\"CR62\" class=\"CitationRef\"\u003e62\u003c/span\u003e, \u003cspan citationid=\"CR63\" class=\"CitationRef\"\u003e63\u003c/span\u003e). Co_414234 was also only observed in the lignin condition and has a ferric reductase transmembrane component-like domain (IPR013130) (\u003cspan citationid=\"CR64\" class=\"CitationRef\"\u003e64\u003c/span\u003e). In the absence of ferric reductases, Fe\u003csup\u003e3+\u003c/sup\u003e can still evolve as a result of Fe\u003csup\u003e2+\u003c/sup\u003e oxidation and participate in oxidation of hydroquinone and semiquinone species (\u003cspan citationid=\"CR49\" class=\"CitationRef\"\u003e49\u003c/span\u003e). Fe\u003csup\u003e2+\u003c/sup\u003e can react with oxygen to generate O\u003csub\u003e2\u003c/sub\u003e\u003csup\u003e\u0026minus;\u0026bull;\u003c/sup\u003e and with H\u003csub\u003e2\u003c/sub\u003eO\u003csub\u003e2\u003c/sub\u003e to produce \u003csup\u003e\u0026minus;\u0026bull;\u003c/sup\u003eOH + \u003csup\u003e\u0026minus;\u003c/sup\u003eOH\u0026mdash;the latter may partially explain the observed pH increase in the lignin-containing cultures (\u003cem\u003eSI Appendix\u003c/em\u003e, Fig. \u003cspan refid=\"MOESM4\" class=\"InternalRef\"\u003eS4\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eH\u003csub\u003e2\u003c/sub\u003eO\u003csub\u003e2\u003c/sub\u003e is generated from other sources including O\u003csub\u003e2\u003c/sub\u003e\u003csup\u003e\u0026minus;\u0026bull;\u003c/sup\u003e (\u003cspan citationid=\"CR50\" class=\"CitationRef\"\u003e50\u003c/span\u003e). Superoxide dismutase (SOD) catalyzes the conversion of O\u003csub\u003e2\u003c/sub\u003e\u003csup\u003e\u0026minus;\u0026bull;\u003c/sup\u003e to H\u003csub\u003e2\u003c/sub\u003eO\u003csub\u003e2\u003c/sub\u003e and is essential for combatting oxidative stress (\u003cspan citationid=\"CR50\" class=\"CitationRef\"\u003e50\u003c/span\u003e). Though localization predictions from \u003cem\u003eTremellomycetes\u003c/em\u003e loosely related to \u003cem\u003eC. oleaginosum\u003c/em\u003e suggest intracellular localization, there are several fungi that secrete SOD variants (\u003cspan citationid=\"CR50\" class=\"CitationRef\"\u003e50\u003c/span\u003e, \u003cspan citationid=\"CR65\" class=\"CitationRef\"\u003e65\u003c/span\u003e). Two copper/zinc-dependent SODs (Co_394365 and Co_419771) with predicted signal peptides and secretion predictions were observed (\u003cspan citationid=\"CR66\" class=\"CitationRef\"\u003e66\u003c/span\u003e). Co_419771 was significantly upregulated in the lignin condition (FC 6.51), which suggests severe oxidative stress. Glucose\u0026ndash;methanol\u0026ndash;choline (GMC) oxidoreductases (AA3), glyoxal oxidases (AA5_1), and oxalate oxidases can also produce H\u003csub\u003e2\u003c/sub\u003eO\u003csub\u003e2\u003c/sub\u003e using a variety of reductants (\u003cspan citationid=\"CR51\" class=\"CitationRef\"\u003e51\u003c/span\u003e). GMC oxidoreductases (AA3_2) can utilize aryl-alcohols and sugars as electron donors, whereas glyoxal oxidases convert simple aldehydes to carboxylic acids (\u003cspan citationid=\"CR67\" class=\"CitationRef\"\u003e67\u003c/span\u003e, \u003cspan citationid=\"CR68\" class=\"CitationRef\"\u003e68\u003c/span\u003e). Oxalate is an important iron chelator during wood decay, and is detoxified to carbon dioxide and H\u003csub\u003e2\u003c/sub\u003eO\u003csub\u003e2\u003c/sub\u003e via oxalate oxidase (\u003cspan citationid=\"CR69\" class=\"CitationRef\"\u003e69\u003c/span\u003e, \u003cspan citationid=\"CR70\" class=\"CitationRef\"\u003e70\u003c/span\u003e). Two GMC oxidoreductases were observed extracellularly (Co_352536 and Co_ 372521) and one intracellularly (Co_342568; DEP FC 4.94). The first two GMC oxidoreductases belong to CUPP branch AA3:6.5, while the other may have a different function as it is categorized in CUPP branch AA3:10.1. Two glyoxal oxidases were observed in the secretomics dataset (Co_344356 and Co_26860). The latter had a signal peptide prediction score of 0.69 but was significantly upregulated in the lignin condition (FC 2.26). An enzyme of the cupin superfamily (Co_373698) was also significantly upregulated (FC 2.73) and is classified as an oxalate oxidase (or superoxide dismutase) by InterPro (IPR001929) (\u003cspan citationid=\"CR64\" class=\"CitationRef\"\u003e64\u003c/span\u003e). Two putative sulfate/bicarbonate/oxalate exchangers were observed in the proteomics results: Co_335524 and Co_352469, which was only observed in the lignin condition.\u003c/p\u003e \u003cp\u003eLaccases and quinone oxidoreductases play significant roles in quinone redox cycles. Laccases are multicopper oxidases that use O\u003csub\u003e2\u003c/sub\u003e to oxidize hydroquinones to semiquinones (\u003cspan citationid=\"CR52\" class=\"CitationRef\"\u003e52\u003c/span\u003e, \u003cspan citationid=\"CR71\" class=\"CitationRef\"\u003e71\u003c/span\u003e). Semiquinones are unstable products that undergo autooxidation to form quinones (\u003cspan citationid=\"CR49\" class=\"CitationRef\"\u003e49\u003c/span\u003e). Two laccases with predicted signal peptides were upregulated in the secretome of the lignin condition: Co_368910 (FC 5.57) and Co_ 369862 (FC 5.68) (\u003cem\u003eSI Appendix\u003c/em\u003e, Fig. \u003cspan refid=\"MOESM4\" class=\"InternalRef\"\u003eS4\u003c/span\u003e). Another putative laccase Co_416150 that shares\u0026thinsp;\u0026gt;\u0026thinsp;37% identity with a laccase from the white-rot fungus \u003cem\u003eTrametes versicolor\u003c/em\u003e (\u003cem\u003eSI Appendix\u003c/em\u003e, Dataset S3) was observed but downregulated in the secretome (FC -0.96) (\u003cspan citationid=\"CR72\" class=\"CitationRef\"\u003e72\u003c/span\u003e). Interestingly, it was only observed in the cellular proteome of the lignin condition. Co_368910, Co_372508, and Co_416150 are secreted AA1 CAZymes according to MULocDeep prediction, and the first two are members of CUPP branch AA1:73.1. (\u003cspan citationid=\"CR66\" class=\"CitationRef\"\u003e66\u003c/span\u003e). In contrast, Co_369862 may locate to the cell membrane and was downregulated (FC -3.29) (\u003cspan citationid=\"CR66\" class=\"CitationRef\"\u003e66\u003c/span\u003e). It is a member of the subfamily AA1_2, which includes ferroxidases that oxidize Fe\u003csup\u003e2+\u003c/sup\u003e to modulate Fenton reactions and decrease \u003csup\u003e\u0026minus;\u0026bull;\u003c/sup\u003eOH (\u003cspan citationid=\"CR73\" class=\"CitationRef\"\u003e73\u003c/span\u003e, \u003cspan citationid=\"CR74\" class=\"CitationRef\"\u003e74\u003c/span\u003e). Indeed, the eggNOG annotation for this enzyme was FET3, a ferroxidase involved in iron uptake. Nonetheless, this enzyme shares 30\u0026ndash;45% identity with phenol-oxidizing laccases from \u003cem\u003eT. versicolor\u003c/em\u003e (\u003cem\u003eSI Appendix\u003c/em\u003e, Dataset S3) (\u003cspan citationid=\"CR72\" class=\"CitationRef\"\u003e72\u003c/span\u003e, \u003cspan citationid=\"CR75\" class=\"CitationRef\"\u003e75\u003c/span\u003e, \u003cspan citationid=\"CR76\" class=\"CitationRef\"\u003e76\u003c/span\u003e). Studies of FET3-like proteins in basidiomycetes show that these multicopper oxidases ostensibly have dual ferroxidase/laccase activities (\u003cspan citationid=\"CR72\" class=\"CitationRef\"\u003e72\u003c/span\u003e, \u003cspan citationid=\"CR75\" class=\"CitationRef\"\u003e75\u003c/span\u003e, \u003cspan citationid=\"CR77\" class=\"CitationRef\"\u003e77\u003c/span\u003e, \u003cspan citationid=\"CR78\" class=\"CitationRef\"\u003e78\u003c/span\u003e). More research into putative yeast laccases is clearly needed.\u003c/p\u003e \u003cp\u003eQuinone oxidoreductases including 1,4-benzoquinone reductase (AA6) are involved in aromatics degradation and protection against reactive quinones (\u003cspan citationid=\"CR51\" class=\"CitationRef\"\u003e51\u003c/span\u003e, \u003cspan additionalcitationids=\"CR80\" citationid=\"CR79\" class=\"CitationRef\"\u003e79\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR81\" class=\"CitationRef\"\u003e81\u003c/span\u003e). A putative quinone oxidoreductase (Co_369953; FC 0.69) and two annotated 1,4-benzoquinone reductases (Co_380727, FC 1.30; Co_164578, FC 0.66) were upregulated in the lignin condition of the proteomics results (\u003cem\u003eSI Appendix\u003c/em\u003e, Fig. \u003cspan refid=\"MOESM4\" class=\"InternalRef\"\u003eS4\u003c/span\u003e). According to BLASTp results, these enzymes share\u0026thinsp;\u0026gt;\u0026thinsp;45% identity with quinone oxidoreductases from \u003cem\u003eRhodonia placenta\u003c/em\u003e (\u003cem\u003eSI Appendix\u003c/em\u003e, Dataset S3) (\u003cspan citationid=\"CR63\" class=\"CitationRef\"\u003e63\u003c/span\u003e). Moreover, Co_380727 is a predicted cell membrane protein, which has \u0026gt;\u0026thinsp;45% identity with functionally characterized quinone oxidoreductases from the white rot fungus \u003cem\u003ePhanerochaete chrysosporium\u003c/em\u003e and brown rot fungus \u003cem\u003eGloeophyllum trabeum\u003c/em\u003e (\u003cspan citationid=\"CR66\" class=\"CitationRef\"\u003e66\u003c/span\u003e, \u003cspan citationid=\"CR80\" class=\"CitationRef\"\u003e80\u003c/span\u003e, \u003cspan citationid=\"CR82\" class=\"CitationRef\"\u003e82\u003c/span\u003e). A cyptoplasmic protein Co_342726 only observed in the lignin replicates was annotated as QOR1, a quinone oxidoreductase, by eggNOG; nevertheless, a conserved domain search on NCBI returned a prostaglandin dehydrogenase (cd05288) (\u003cspan citationid=\"CR83\" class=\"CitationRef\"\u003e83\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eIn addition to the quinone redox cycle, peroxidases can also oxidize hydroquinone to yield 1,4-benzoquinone (\u003cspan citationid=\"CR79\" class=\"CitationRef\"\u003e79\u003c/span\u003e). Though many peroxidases were observed in the secretome (Co_396749, Co_237718, Co_382630, and Co_372514), none of them were upregulated, and only one (Co_396749) had a secretion prediction (\u003cspan citationid=\"CR66\" class=\"CitationRef\"\u003e66\u003c/span\u003e). Given that both the glucose and lignin culture conditions were carbon-limited, this may be explained by general upregulation of genes under carbon catabolite derepression (\u003cspan citationid=\"CR84\" class=\"CitationRef\"\u003e84\u003c/span\u003e). Peroxidase expression may also be a function of culture conditions like nitrogen source as Martinez et al. observed with \u003cem\u003ePleurotus eryngii\u003c/em\u003e (\u003cspan citationid=\"CR49\" class=\"CitationRef\"\u003e49\u003c/span\u003e, \u003cspan citationid=\"CR85\" class=\"CitationRef\"\u003e85\u003c/span\u003e). Nevertheless, peroxidase activity has been observed with \u003cem\u003eC. oleaginosum\u003c/em\u003e as well as another \u003cem\u003eTrichosporonaceae\u003c/em\u003e yeast (\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e, \u003cspan citationid=\"CR86\" class=\"CitationRef\"\u003e86\u003c/span\u003e). BLASTp results of AA2 peroxidases from \u003cem\u003eT. versicolor\u003c/em\u003e, \u003cem\u003eP. chrysosporium\u003c/em\u003e, and the recently isolated relative \u003cem\u003eCutaneotrichosporon cavernicola\u003c/em\u003e yield Co_382630 and Co_372514 with \u0026gt;\u0026thinsp;55% identity (\u003cem\u003eSI Appendix\u003c/em\u003e, Dataset S3) (\u003cspan citationid=\"CR50\" class=\"CitationRef\"\u003e50\u003c/span\u003e, \u003cspan citationid=\"CR87\" class=\"CitationRef\"\u003e87\u003c/span\u003e). Interestingly, these enzymes were not annotated by CUPP nor were AA2 CAZy annotations included for these entries on JGI (\u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e43\u003c/span\u003e, \u003cspan citationid=\"CR47\" class=\"CitationRef\"\u003e47\u003c/span\u003e, \u003cspan citationid=\"CR51\" class=\"CitationRef\"\u003e51\u003c/span\u003e, \u003cspan citationid=\"CR88\" class=\"CitationRef\"\u003e88\u003c/span\u003e). Based on the aforementioned and because these enzymes do not have predicted signal peptides, it\u0026rsquo;s likely that these enzymes are AA2 class I peroxidases. This class of peroxidases has lower redox potentials than class II lignin and manganese peroxidases, and they are generally involved in oxidative stress response (\u003cspan citationid=\"CR89\" class=\"CitationRef\"\u003e89\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eThese proteomics results suggest a capacity for quinone redox cycling by \u003cem\u003eC. oleaginosum\u003c/em\u003e, even though the reactivity of Fenton chemistry was likely suboptimal under the near neutral culture conditions studied here (starting pH of 6.5; see \u003cem\u003eSI Appendix\u003c/em\u003e, Fig. \u003cspan refid=\"MOESM4\" class=\"InternalRef\"\u003eS4\u003c/span\u003e) (\u003cspan citationid=\"CR57\" class=\"CitationRef\"\u003e57\u003c/span\u003e). To bolster this conclusion, the mechanisms by which hydroquinones are generated \u003cem\u003ede novo\u003c/em\u003e and secreted to help initiate redox cycling will need to be studied in yeast. Hydroquinone secretion has been observed with filamentous fungi; however, it is unclear what transporters are involved because few have been functionally characterized (\u003cspan citationid=\"CR54\" class=\"CitationRef\"\u003e54\u003c/span\u003e, \u003cspan citationid=\"CR90\" class=\"CitationRef\"\u003e90\u003c/span\u003e, \u003cspan citationid=\"CR91\" class=\"CitationRef\"\u003e91\u003c/span\u003e). Of course, this is also the case for the other candidate enzymes discussed hitherto\u0026mdash;direct functional genomics analyses will be essential for resolving ambiguities regarding this yeast\u0026rsquo;s laccase and peroxidase activities (\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e, \u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e). Overall, these findings highlight the need for additional research to validate the proposed candidates and fully understand the quinone redox cycling potential of \u003cem\u003eC. oleaginosum\u003c/em\u003e and other yeasts.\u003c/p\u003e \u003cp\u003e \u003cb\u003eC. oleaginosum\u003c/b\u003e \u003cb\u003eMay Uptake Poymeric Lignin but the Mechanism Remains Unclear\u003c/b\u003e\u003c/p\u003e \u003cp\u003eTransport mechanisms for lignin-derived aromatics in yeast are underexplored (\u003cspan citationid=\"CR92\" class=\"CitationRef\"\u003e92\u003c/span\u003e, \u003cspan citationid=\"CR93\" class=\"CitationRef\"\u003e93\u003c/span\u003e). In bacteria and filamentous fungi, active transport of monoaromatics involves ATP-binding cassette (ABC) transporters and major facilitator superfamily (MFS) transporters (\u003cspan citationid=\"CR33\" class=\"CitationRef\"\u003e33\u003c/span\u003e). Passive transport has also been suggested, though most research has been conducted in bacteria which have drastically different cellular envelopes (\u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e30\u003c/span\u003e, \u003cspan citationid=\"CR94\" class=\"CitationRef\"\u003e94\u003c/span\u003e). Proteomics can help elucidate potential aromatics transporters by highlighting those that are upregulated or only observed in the lignin condition. We observed 380 of the 790 JGI-annotated transporters (\u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e43\u003c/span\u003e, \u003cspan citationid=\"CR95\" class=\"CitationRef\"\u003e95\u003c/span\u003e). The relative abundances of 246 were quantified for lignin vs. glucose: 48 were upregulated (FC\u0026thinsp;\u0026gt;\u0026thinsp;0.8) while 18 were downregulated (FC\u0026thinsp;\u0026lt;\u0026thinsp;\u0026minus;\u0026thinsp;0.8). Two enzymes annotated as \u003cem\u003ep\u003c/em\u003e-coumarate-CoA synthases were upregulated (Co_383485, FC 2.57; and Co_386597, FC 1.26; \u003cem\u003eSI Appendix\u003c/em\u003e, Dataset S2). These are annotated members of a Fatty Acid Group Translocation Family that use ATP to attach CoA to \u003cem\u003ep\u003c/em\u003e-coumarate and other aromatics in a process that may be coupled to transport (\u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e43\u003c/span\u003e, \u003cspan citationid=\"CR66\" class=\"CitationRef\"\u003e66\u003c/span\u003e, \u003cspan citationid=\"CR95\" class=\"CitationRef\"\u003e95\u003c/span\u003e). Upregulation of these enzymes is in line with a decrease in H-lignin (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eA and C). Enzymes with cell membrane localization predictions include an arabinose, xylose, and/or aromatics transporter (Co_378211; FC 0.83), a MFS transporter (Co_419501; DEP FC 3.06), and an amino acid transporter (Co_391320; FC 1.37) with a phenol hydroxylase domain (IPR012941) (\u003cspan citationid=\"CR64\" class=\"CitationRef\"\u003e64\u003c/span\u003e). A MFS high affinity nicotinate permease (Co_96030) and an ABC transporter (Co_345222) were observed exclusively in the lignin condition. The latter is annotated as a plant cuticular wax and/or lipid exporter. Plant cuticles have complex compositions that include aromatics (\u003cspan citationid=\"CR96\" class=\"CitationRef\"\u003e96\u003c/span\u003e). Lastly, a MFS quinate:H\u0026thinsp;+\u0026thinsp;symporter (Co_96856) was only observed in the lignin condition. Fewer transporters were upregulated in our results compared to a recent study of the yeast \u003cem\u003eRhodosporidium fluviale\u003c/em\u003e cultivated on black liquor, but this may be due to substantially higher aromatics concentrations in their medium (\u003cspan citationid=\"CR28\" class=\"CitationRef\"\u003e28\u003c/span\u003e). We also conducted searches using hydroxybenzoate transporters recently characterized in the pathogenic yeast \u003cem\u003eCandida parapsilosis\u003c/em\u003e, but no high-scoring alignments were found (\u003cem\u003eSI Appendix\u003c/em\u003e, Dataset S3) (\u003cspan citationid=\"CR97\" class=\"CitationRef\"\u003e97\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eA recent publication reported active transport and subsequent degradation of an aromatic dimer by \u003cem\u003eP. chrysosporium\u003c/em\u003e (\u003cspan citationid=\"CR93\" class=\"CitationRef\"\u003e93\u003c/span\u003e). However, there is limited research explicitly addressing lignin uptake in microbial systems. We speculate that polymeric lignin can be internalized by \u003cem\u003eC. oleaginosum\u003c/em\u003e based on uptake of polyaromatic polystyrene in yeast (\u003cspan citationid=\"CR34\" class=\"CitationRef\"\u003e34\u003c/span\u003e, \u003cspan citationid=\"CR98\" class=\"CitationRef\"\u003e98\u003c/span\u003e, \u003cspan citationid=\"CR99\" class=\"CitationRef\"\u003e99\u003c/span\u003e). As a preliminary investigation, we used confocal fluorescence microscopy to harness the autofluorescent signatures of lignin and study its uptake (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e4\u003c/span\u003eA\u0026ndash;B and \u003cem\u003eSI Appendix\u003c/em\u003e, Fig. \u003cspan refid=\"MOESM5\" class=\"InternalRef\"\u003eS5\u003c/span\u003e) (\u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e35\u003c/span\u003e, \u003cspan citationid=\"CR36\" class=\"CitationRef\"\u003e36\u003c/span\u003e). We employed the cell segmentation software Cellpose to analyze yeast autofluorescence using various controls (\u003cspan additionalcitationids=\"CR101\" citationid=\"CR100\" class=\"CitationRef\"\u003e100\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR102\" class=\"CitationRef\"\u003e102\u003c/span\u003e). Cellpose identified cells with high accuracy even in the presence of lignin particles, thereby facilitating cell population measurements of fluorescence (\u003cem\u003eSI Appendix\u003c/em\u003e, Fig. S6A). Though unapparent to the naked eye, cells in the lignin condition have significantly higher fluorescence intensities for all comparisons except lignin vs. benzoate at 48 hours (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e4\u003c/span\u003eB and \u003cem\u003eSI Appendix\u003c/em\u003e, Fig. S6B\u0026ndash;C). It\u0026rsquo;s possible that the increase in yeast autofluorescence confounds any differences, perhaps due to carbon-limited stress response and/or aromatics toxicity, given that 1 g/L benzoate is close to the minimum inhibitory concentration of \u003cem\u003eC. oleaginosum\u003c/em\u003e (\u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e, \u003cspan citationid=\"CR100\" class=\"CitationRef\"\u003e100\u003c/span\u003e, \u003cspan citationid=\"CR103\" class=\"CitationRef\"\u003e103\u003c/span\u003e, \u003cspan citationid=\"CR104\" class=\"CitationRef\"\u003e104\u003c/span\u003e). We also performed super resolution fluorescence imaging to illustrate structural changes due to lignin interactions (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e4\u003c/span\u003eC). For conditions without lignin, the weak autofluorescence is consistent with the structure of mitochondria. With lignin the fluorescence is more punctate and widespread throughout the cell, indicating significant changes to subcellular morphology. Moreover, there is a clear delineation of the cellular envelope. Using a dye to label mitochondria, the observed autofluorescence in the absence of lignin is partially due to mitochondria, whereas in the presence of lignin there is no apparent dye localization (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e4\u003c/span\u003eD). This suggests drastic changes to subcellular processes and possibility of lignin-induced membrane destabilization, which has been observed using other nanoparticles (\u003cem\u003eSI Appendix\u003c/em\u003e, Supporting Text) (\u003cspan citationid=\"CR105\" class=\"CitationRef\"\u003e105\u003c/span\u003e). Overall, these results suggest polymeric lignin uptake by \u003cem\u003eC. oleaginosum\u003c/em\u003e, but investigations employing labeled lignin nanoparticles and electron microscopy will be needed for validation and to explore mechanisms.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e\n\u003ch3\u003ePathways for Aromatics Catabolism Are Upregulated in the Lignin Condition\u003c/h3\u003e\n\u003cp\u003eYeasts funnel aromatic monomers to intermediates such as protocatechuate, catechol, and hydroxyquinol via intricate upper pathways (\u003cspan citationid=\"CR106\" class=\"CitationRef\"\u003e106\u003c/span\u003e). Dioxygenases cleave the aromatic rings yielding intermediates that are metabolized through lower pathways (e.g., branches of the β-ketoadipate pathway), ultimately leading to central metabolites (\u003cspan citationid=\"CR107\" class=\"CitationRef\"\u003e107\u003c/span\u003e, \u003cspan citationid=\"CR108\" class=\"CitationRef\"\u003e108\u003c/span\u003e). Aromatics catabolic enzymes and the corresponding pathways are largely uncharacterized in \u003cem\u003eC. oleaginosum\u003c/em\u003e. In an earlier report, cleavage positions for catechol, protocatechuate, and hydroxyquinol were assayed using clarified lysates (\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e). Catechol and hydroxyquinol underwent \u003cem\u003eortho\u003c/em\u003e-cleavage, whereas cleavage of proteocatechuate was negligible suggesting that it is decarboxylated to hydroxyquinol or catabolized via some other route. They also functionally annotated a catechol dioxygenase (homolog of Co_374756) that has intradiol and extradiol ring-cleaving activities. Beyond these analyses, specific catabolic pathways were not proposed; therefore, we scoured publications in which aromatics catabolic enzymes were functionally characterized in fungi (and some cases bacteria). We then conducted extensive BLASTp searches against proteins observed in our proteomics results to project putative aromatics catabolic pathways in \u003cem\u003eC. oleaginosum\u003c/em\u003e (\u003cem\u003eSI Appendix\u003c/em\u003e, Datasets S1\u0026ndash;3) (\u003cspan citationid=\"CR58\" class=\"CitationRef\"\u003e58\u003c/span\u003e). Rationalized by our 2D-NMR results and by the significance of ferulate G-lignin monomers in corn stover (\u003cspan citationid=\"CR109\" class=\"CitationRef\"\u003e109\u003c/span\u003e), we focused our efforts on projecting catabolic pathways for H-lignin derived aromatics (refer to \u003cem\u003eSI Appendix\u003c/em\u003e, Supporting Text for more information about predicted pathways for G-lignin, S-lignin, and phenol derivatives).\u003c/p\u003e \u003cp\u003e \u003cem\u003eC. oleaginosum\u003c/em\u003e utilizes H-lignin model aromatics \u003cem\u003ep\u003c/em\u003e-coumarate and \u003cem\u003ep\u003c/em\u003e-hydroxybenzoate, but pathways have yet to be projected (\u003cspan additionalcitationids=\"CR111\" citationid=\"CR110\" class=\"CitationRef\"\u003e110\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR112\" class=\"CitationRef\"\u003e112\u003c/span\u003e). In filamentous fungi, conversion of \u003cem\u003ep\u003c/em\u003e-coumarate to \u003cem\u003ep\u003c/em\u003e-hydroxybenzoate has been extensively studied (\u003cspan additionalcitationids=\"CR114 CR115\" citationid=\"CR113\" class=\"CitationRef\"\u003e113\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR116\" class=\"CitationRef\"\u003e116\u003c/span\u003e). Lubbers et al. demonstrated that \u003cem\u003eAspergillus niger\u003c/em\u003e uses a CoA-dependent β-oxidative route akin to fatty acid degradation (\u003cspan citationid=\"CR117\" class=\"CitationRef\"\u003e117\u003c/span\u003e). Briefly, \u003cem\u003ep\u003c/em\u003e-coumarate is converted to \u003cem\u003ep\u003c/em\u003e-coumaroyl-SCoA by a hydroxycinnamate-CoA synthase (hcsA). The product is then hydrated and dehydrogenated by a multifunctional β-oxidation hydratase/dehydrogenase (foxA) to yield 4-hydroxyphenyl-β-ketopropionic acid-SCoA (HPOP-CoA). Afterwards, \u003cem\u003ep\u003c/em\u003e-benzoyl-CoA is generated by 3-ketoacyl-CoA thiolase (katA). Finally, thioesterases remove CoA to produce \u003cem\u003ep\u003c/em\u003e-hydroxybenzoate. We searched our proteomics data for homologs of characterized proteins. The BLASTp scores for \u003cem\u003eAspergilli\u003c/em\u003e hcsA were fairly low, so we also searched bacterial enzymes involved in the CoA-dependent non-oxidative pathway (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e6\u003c/span\u003e; \u003cem\u003eSI Appendix\u003c/em\u003e, Dataset S3) (\u003cspan additionalcitationids=\"CR118 CR119 CR120\" citationid=\"CR117\" class=\"CitationRef\"\u003e117\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR121\" class=\"CitationRef\"\u003e121\u003c/span\u003e). Several putative genes were upregulated in the cytoplasm and peroxisome: Co_383485 (FC 2.57; cytoplasm), Co_386597 (FC 1.26; cytoplasm), Co_386202 (FC, 2.09; peroxisome), Co_376641 (FC 1.32; peroxisome), and Co_370579 (FC 1.05; peroxisome). Only one peroxisomal foxA homolog was upregulated (Co_233545; FC 0.98) and contained a C-terminal peroxisomal targeting sequence (\u003cspan citationid=\"CR66\" class=\"CitationRef\"\u003e66\u003c/span\u003e, \u003cspan citationid=\"CR122\" class=\"CitationRef\"\u003e122\u003c/span\u003e, \u003cspan citationid=\"CR123\" class=\"CitationRef\"\u003e123\u003c/span\u003e). BLASTp results of bacterial hydroxycinnamoyl-CoA hydratase-lyases returned Co_381918 (FC 0.93; peroxisome). Though many katA and thioesterase homologs were observed, few had peroxisomal predictions. According to MULocDeep, Co_368669 and Co_371904 are localized in the cytoplasm; however, both have C-terminal peroxisomal targeting sequences (\u003cspan citationid=\"CR66\" class=\"CitationRef\"\u003e66\u003c/span\u003e, \u003cspan citationid=\"CR122\" class=\"CitationRef\"\u003e122\u003c/span\u003e, \u003cspan citationid=\"CR123\" class=\"CitationRef\"\u003e123\u003c/span\u003e). β-oxidation of fatty acids was upregulated in the lignin condition (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e5\u003c/span\u003eF): those proteins may be candidates for CoA-dependent \u003cem\u003ep\u003c/em\u003e-coumarate catabolism in \u003cem\u003eC. oleaginosum\u003c/em\u003e.\u003c/p\u003e \u003cp\u003ePrior to ring cleavage, \u003cem\u003ep\u003c/em\u003e-hydroxybenzoate is converted to hydroxyquinol (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e6\u003c/span\u003e). Phenol hydroxylase is active on phenol derivatives and has been characterized in the close relative \u003cem\u003eCutaneotrichosporon cutaneum\u003c/em\u003e (\u003cspan citationid=\"CR124\" class=\"CitationRef\"\u003e124\u003c/span\u003e, \u003cspan citationid=\"CR125\" class=\"CitationRef\"\u003e125\u003c/span\u003e). Three homologs were upregulated in our results: Co_391320 (FC 1.37), Co_374925 (FC 1.64), and Co_222522 (FC 2.31). Phenol hydroxylase from \u003cem\u003eC. cutaneum\u003c/em\u003e is structurally related to fungal 4-hydroxybenzoate 3-hydroxylase, so these homologs may be involved in the conversion of \u003cem\u003ep\u003c/em\u003e-hydroxybenzoate to protocatechuate (\u003cspan citationid=\"CR126\" class=\"CitationRef\"\u003e126\u003c/span\u003e). Oxidative decarboxylation of \u003cem\u003ep\u003c/em\u003e-hydroxybenzoate and protocatechuate to hydroquinone and hydroxyquinol, respectively, has been reported for the yeast \u003cem\u003eC. parapsilosis\u003c/em\u003e and white-rot fungi (\u003cspan citationid=\"CR127\" class=\"CitationRef\"\u003e127\u003c/span\u003e, \u003cspan citationid=\"CR128\" class=\"CitationRef\"\u003e128\u003c/span\u003e). Co_382250 was the highest scoring hit for 4-hydroxybenzoate 1-hydroxylase (i.e., MNX1) from \u003cem\u003eC. parapsilosis\u003c/em\u003e but was not observed in our proteomics results, but the putative homolog Co_419806 was observed and upregulated 1.56-fold. It has a low alignment score with MNX1 but higher scores against putative basidiomycete decarboxylases (\u003cspan citationid=\"CR127\" class=\"CitationRef\"\u003e127\u003c/span\u003e). This is also the case for Co_417242 which was drastically upregulated (FC 4.25) and shows higher homology with a reported \u003cem\u003eT. versicolor\u003c/em\u003e decarboxylase (TV_32834) that preferentially decarboxylates protocatechuate. The putative \u003cem\u003eGelatoporia subvermispora\u003c/em\u003e homolog (GS_120062) of Co_ 419806 displays activity against \u003cem\u003ep\u003c/em\u003e-hydroxybenzoate and protocatechuate (\u003cspan citationid=\"CR127\" class=\"CitationRef\"\u003e127\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eRing cleavage detoxifies aromatics and yields assimilable carbon sources (\u003cspan citationid=\"CR129\" class=\"CitationRef\"\u003e129\u003c/span\u003e). Enzymes that catabolize hydroxyquinol and protocatechuate have been investigated using \u003cem\u003eC. parapsilosis\u003c/em\u003e and \u003cem\u003eA. niger\u003c/em\u003e (\u003cspan citationid=\"CR128\" class=\"CitationRef\"\u003e128\u003c/span\u003e, \u003cspan citationid=\"CR130\" class=\"CitationRef\"\u003e130\u003c/span\u003e). Based on BLASTp results, Co_374756 is the top candidate for hydroxyquinol 1,2-dioxygenase activity (\u003cem\u003eSI Appendix\u003c/em\u003e, Dataset S3). Co_85172 is an additional candidate (FC 2.44), but it had a higher E-value (5.50E-39) against a protocatechuate 3,4-dioxygenase (NRRL3_1405) from \u003cem\u003eA. niger\u003c/em\u003e. Another enzyme Co_340256 was annotated as the catalytic LigB subunit of aromatic extradiol ring-opening dioxygenase (PF02900) (\u003cspan citationid=\"CR64\" class=\"CitationRef\"\u003e64\u003c/span\u003e, \u003cspan citationid=\"CR76\" class=\"CitationRef\"\u003e76\u003c/span\u003e). LigB is a subunit of the heterodimer LigAB that exhibits protocatechuate 3,4-dioxygenase activity in the lignin-degrading bacteria \u003cem\u003eSphingobium\u003c/em\u003e (\u003cspan citationid=\"CR131\" class=\"CitationRef\"\u003e131\u003c/span\u003e). Intradiol hydroxyquinol cleavage yields maleylacetate, which is converted to β-ketoadipate by maleylacetate reductase. Negligable upregulation of Co_373930 (FC 0.25), a homolog of the \u003cem\u003eA. niger\u003c/em\u003e candidate, was observed (\u003cspan citationid=\"CR132\" class=\"CitationRef\"\u003e132\u003c/span\u003e). A homolog of another candidate observed in \u003cem\u003eC. parapsilosis\u003c/em\u003e was observed but not differentially expressed (Co_374156) (\u003cspan citationid=\"CR133\" class=\"CitationRef\"\u003e133\u003c/span\u003e, \u003cspan citationid=\"CR134\" class=\"CitationRef\"\u003e134\u003c/span\u003e). Intradiol cleavage of protocatechuate leads to formation of β-carboxymuconate, which undergoes cyclization via muconate cycloisomerase (\u003cspan citationid=\"CR135\" class=\"CitationRef\"\u003e135\u003c/span\u003e). The product γ-carboxymuconolactone can be decarboxylated to β-ketoadipate enol-lactone as observed in \u003cem\u003eA. nidulans\u003c/em\u003e (\u003cspan citationid=\"CR136\" class=\"CitationRef\"\u003e136\u003c/span\u003e). A homolog of the decarboxylase studied by Martins et al. (\u003cspan citationid=\"CR136\" class=\"CitationRef\"\u003e136\u003c/span\u003e) was not observed in \u003cem\u003eC. oleaginosum\u003c/em\u003e. Thankfully, the eggNOG description of Co_371232 (FC 2.94) was \u0026ldquo;Carboxymuconolactone decarboxylase family\u0026rdquo;, and the orthologous group annotation was \u0026ldquo;Sphingomonadales\u0026rdquo;, which raises interesting questions about horizontal gene transfer among soil-dwelling microorganisms (\u003cspan citationid=\"CR76\" class=\"CitationRef\"\u003e76\u003c/span\u003e). Two additional candidates include Co_350958 (FC 2.52; aminocarboxymuconate-semialdehyde decarboxylase) and Co_372335.\u003c/p\u003e \u003cp\u003eAs a whole, we speculate that mechanisms for lignin modification are present in \u003cem\u003eC. oleaginosum\u003c/em\u003e and demonstrate upregulation of predicted catabolic pathways for H-lignin and G-lignin derivatives (\u003cem\u003eSI Appendix\u003c/em\u003e, Datasets S1\u0026ndash;2). It follows that entry point enzymes for the TCA cycle and glyoxalate shunt were also upregulated, whereas glycolytic enzymes were downregulated (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e6\u003c/span\u003e). Antioxidants are important for mitigating excess ROS such as H\u003csub\u003e2\u003c/sub\u003eO\u003csub\u003e2\u003c/sub\u003e, which can be generated extracellularly and re-enter the cell via direct permeation or aquaporins (\u003cspan citationid=\"CR50\" class=\"CitationRef\"\u003e50\u003c/span\u003e). We observed upregulation of several antioxidants including catalases (Co_370626, FC 2.07; Co_371234, FC 1.60) that neutralize H\u003csub\u003e2\u003c/sub\u003eO\u003csub\u003e2\u003c/sub\u003e. Nevertheless, many of these enzymes were only partially upregulated in the lignin vs. No Carbon comparison, which speaks to their general role in nutrient-limited stress response (\u003cspan citationid=\"CR48\" class=\"CitationRef\"\u003e48\u003c/span\u003e). For specific enzymes like glutathione-dependent formaldehyde-activating enzyme (Co_398886, FC 1.42), detoxification of formaldehyde from demethylation of aromatics may rationalize these differences (\u003cspan citationid=\"CR137\" class=\"CitationRef\"\u003e137\u003c/span\u003e). These findings hint at similarities in aromatics catabolism between yeasts and filamentous fungi of the Agaricomycotina subphylum, yet they also underscore gaps in our understanding. The lack of metabolomics results herein presents a significant opportunity for future studies but also poses challenges. These include limitations in current techniques, the scarcity of certain aromatics released even by white-rot fungi, and the intricate nature of lignin degradation intermediates (\u003cspan citationid=\"CR127\" class=\"CitationRef\"\u003e127\u003c/span\u003e, \u003cspan citationid=\"CR138\" class=\"CitationRef\"\u003e138\u003c/span\u003e). Assays that directly measure ROS generation will also further our understanding of redox mechanisms involved in aromatics detoxification and lignin degradation (\u003cspan citationid=\"CR139\" class=\"CitationRef\"\u003e139\u003c/span\u003e). Redox proteomics, which assesses protein oxidation, offers an innovative approach to explore how fungi manage redox homeostasis during lignin degradation (\u003cspan citationid=\"CR140\" class=\"CitationRef\"\u003e140\u003c/span\u003e, \u003cspan citationid=\"CR141\" class=\"CitationRef\"\u003e141\u003c/span\u003e).\u003c/p\u003e"},{"header":"Conclusion","content":"\u003cp\u003eIn this exploratory study, we cultivated \u003cem\u003eC. oleaginosum\u003c/em\u003e on alkali lignin and, according to 2D-NMR, observed modification of the H-lignin structure, \u003cem\u003ep\u003c/em\u003e-coumarate and ferulate pendents, and interunit bonding. A comprehensive proteomics analysis revealed the potential for yeast quinone redox cycling as well as putative pathways for aromatics catabolism, which were upregulated in the lignin condition. Based on these results, we conclude that this yeast and likely other basidiomycete yeasts have a ligninolytic capacity, thereby playing salient roles in lignocellulose degradation and aromatics detoxification in native microbial communities. Nonetheless, challenges in analyzing secretomics data, especially under stressful carbon-limited conditions that may cause cell lysis, highlight the complexity of interpreting these results. Additionally, the influence of culture conditions, sampling time points, and localization of putative ligninolytic enzymes (e.g., \u003cem\u003eCryptococcus neoformans\u003c/em\u003e expresses cell well-bound laccase) further complicates interpretation of results using this yeast (\u003cspan citationid=\"CR142\" class=\"CitationRef\"\u003e142\u003c/span\u003e, \u003cspan citationid=\"CR143\" class=\"CitationRef\"\u003e143\u003c/span\u003e). These insights call for continued exploration and refinement of methodologies to enhance our understanding of the ligninolytic potential of yeasts. This will advance efforts to metabolically engineer yeasts to express complementary ligninolytic enzymes from white-rot fungi and optimize the utilization of lignin-derived aromatics.\u003c/p\u003e "},{"header":"Methods","content":"\u003cdiv id=\"Sec8\" class=\"Section2\"\u003e \u003cp\u003eA brief summary of relevant materials and methods is presented below. Additional details are included in \u003cem\u003eSI Appendix\u003c/em\u003e, Supplementary Methods.\u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eLignin Extraction from Corn Stover\u003c/h3\u003e\n\u003cp\u003eThe National Renewable Energy Laboratory provided the black liquor fraction from dilute alkali-pretreated corn stover, which was used for lignin extraction (\u003cspan citationid=\"CR144\" class=\"CitationRef\"\u003e144\u003c/span\u003e, \u003cspan citationid=\"CR145\" class=\"CitationRef\"\u003e145\u003c/span\u003e). The extraction process involved serial dilute acid-base titrations, centrifugation, and solids washing (\u003cspan citationid=\"CR29\" class=\"CitationRef\"\u003e29\u003c/span\u003e). Compositional analysis details for the resulting extract are provided in \u003cem\u003eSI Appendix\u003c/em\u003e, Table \u003cspan refid=\"MOESM1\" class=\"InternalRef\"\u003eS1\u003c/span\u003e (\u003cspan citationid=\"CR146\" class=\"CitationRef\"\u003e146\u003c/span\u003e).\u003c/p\u003e\n\u003ch3\u003eYeast Strain, Medium Summary, and Cultivation\u003c/h3\u003e\n\u003cp\u003e \u003cem\u003eCutaneotrichosporon oleaginosum\u003c/em\u003e ATCC 20509 (formerly classified as \u003cem\u003eCryptococcus curvatus\u003c/em\u003e) was used in this study. For long-term storage, glycerol (15% v/v) stocks stored at \u0026minus;\u0026thinsp;80\u0026deg;C were prepared from cells streaked on standard Yeast extract-Peptone-Dextrose (YPD) plates. YPD was used for short term plate storage at 4\u0026deg;C as well as generating seed cultures. The yeast was cultivated in a YNB-based experimental medium containing 1 g/L carbon source (lignin extract, glucose, or benzoate) or no additional carbon source (\u0026ldquo;No Carbon\u0026rdquo; condition). Cultivation conditions were 28\u0026deg;C shaking at 180 rpm.\u003c/p\u003e \u003cdiv id=\"Sec11\" class=\"Section2\"\u003e \u003ch2\u003e2D-HSQC-NMR\u003c/h2\u003e \u003cp\u003eTwo-dimensional (2D) heteronuclear single-quantum correlation (HSQC) NMR spectroscopy was used to compare alkali lignin (control) to lignin in the lyophilized yeast culture supernatants (\u003cspan citationid=\"CR147\" class=\"CitationRef\"\u003e147\u003c/span\u003e, \u003cspan citationid=\"CR148\" class=\"CitationRef\"\u003e148\u003c/span\u003e).\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec12\" class=\"Section2\"\u003e \u003ch2\u003eFluorescence Microscopy\u003c/h2\u003e \u003cp\u003eLive yeast cultures were imaged by an inverted fluorescence confocal microscope with oil immersion. The fluorescence from lignin was excited by a 488 nm wavelength laser according to a preliminary analysis of lignin excitation and emission wavelengths (\u003cem\u003eSI Appendix\u003c/em\u003e, Fig. \u003cspan refid=\"MOESM5\" class=\"InternalRef\"\u003eS5\u003c/span\u003e). The Cellpose software was employed for cellular segmentation to analyze yeast cell fluorescence for the aforementioned carbon source conditions.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec13\" class=\"Section2\"\u003e \u003ch2\u003eSecretomics and Cellular Proteomics\u003c/h2\u003e \u003cp\u003eSupernatants and washed pellets were collected from yeast cultivations to generate extracellular (secretomics) and intracellular (cellular proteomics) samples, respectively. All samples were processed using the SP3 method but with modifications depending on the sample type (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003e) (\u003cspan citationid=\"CR149\" class=\"CitationRef\"\u003e149\u003c/span\u003e, \u003cspan citationid=\"CR150\" class=\"CitationRef\"\u003e150\u003c/span\u003e). All samples were digested with an enzyme cocktail containing trypsin (Promega) and Lys-C (Wako Chemicals). Samples were then processed with Zip-Tip C18 SPE and analyzed using LC-MS/MS.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec14\" class=\"Section2\"\u003e \u003ch2\u003eData Analysis and Processing\u003c/h2\u003e \u003cp\u003eTwo-tailed t-tests were used for determining statistically significant differences in growth, total aromatics, and fluorescence intensities among conditions. For these analyses, p-values were adjusted for multiple comparisons using the Holm\u0026rsquo;s method. Significance levels and error bars are defined in each figure caption. Protein sequences and annotations for \u003cem\u003eC. oleaginosum\u003c/em\u003e were retrieved from the respective Joint Genome Institute (JGI) database (\u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e43\u003c/span\u003e, \u003cspan citationid=\"CR151\" class=\"CitationRef\"\u003e151\u003c/span\u003e). Bottom-up proteomics data were processed with MaxQuant for label-free quantification (\u003cspan citationid=\"CR152\" class=\"CitationRef\"\u003e152\u003c/span\u003e). All log\u003csub\u003e2\u003c/sub\u003e foldchange (\u0026ldquo;FC\u0026rdquo;) data correspond to differential expression analyses using the DEqMS package with MaxQuant LFQ intensities, unless otherwise specified as using the DEP package with normalized raw intensity values (\u003cspan citationid=\"CR153\" class=\"CitationRef\"\u003e153\u003c/span\u003e, \u003cspan citationid=\"CR154\" class=\"CitationRef\"\u003e154\u003c/span\u003e). For proteomics differential expression analysis, p-values were adjusted using the the Benjamini\u0026ndash;Hochberg procedure. Refer to \u003cem\u003eSI Appendix\u003c/em\u003e, Supplementary Methods for details regarding RStudio Bioconductor packages and annotation prediction tools used for bioinformatics analyses (\u003cspan citationid=\"CR155\" class=\"CitationRef\"\u003e155\u003c/span\u003e).\u003c/p\u003e \u003c/div\u003e "},{"header":"Declarations","content":"\u003ch2\u003eCompeting Interest Statement\u003c/h2\u003e \u003cp\u003eThe authors declare no competing interests.\u003c/p\u003e \u003ch2\u003eAuthor Contribution\u003c/h2\u003e\u003cp\u003eA.G. and B.Y. designed research; A.G., Y.P., D.H., and X.L. performed research; Y.P., D.H., X.C., A.J.R., T.Z., and W-J.Q. contributed new reagents/analytical tools; A.G., Y.P., D.H., and Z.J. analyzed data; A.G. wrote the original draft; B,Y, and W-J.Q assisted with conceptualization, supervision, and acquisition of experimental resources and project funding. all authors contributed to the writing and final edition of the final manuscript.\u003c/p\u003e\u003ch2\u003eAcknowledgement\u003c/h2\u003e\u003cp\u003eWe acknowledge the U.S. Department of Energy (DOE) Office of Energy Efficiency \u0026amp; Renewable Energy (DE-EE0008250, and DE-EE0009763), the U.S. Department of Agriculture (USDA) National Institute of Food and Agriculture (Hatch/Multi State project 1017904), and the Bioproducts, Science and Engineering Laboratory, Department of Biological Systems Engineering at Washington State University (WSU). Portions of this research were also supported by the Predictive Phenomics Initiative (TZ) conducted under the Laboratory Directed Research and Development Program at Pacific Northwest National Laboratory (PNNL) in addition to a project award from the Environmental Molecular Sciences Laboratory, a DOE Office of Science User Facility sponsored by the Biological and Environmental Research program under Contract No. DE-AC05-76RL01830. YP and AJR acknowledge partial support from the Center for Bioenergy Innovation (CBI) at Oak Ridge National Laboratory, a U.S. DOE Bioenergy Research Center supported by the Office of Biological and Environmental Research in the DOE Office of Science. AG is grateful for support from the PNNL-WSU Distinguished Graduate Research Program Fellowship. We are also thankful for the FAMEs protocol from PNNL scientists Mark G. Butcher and Angela M. Melville.\u003c/p\u003e\u003ch2\u003eData Availability\u003c/h2\u003e\u003cp\u003eThe data supporting this study\u0026rsquo;s findings are available from the corresponding author (Drs. Wen-Jun Qian, and Bin Yang) upon reasonable request.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\u003cli\u003e\u003cspan\u003eWang, H., Pu, Y., Ragauskas, A. \u0026amp; Yang, B. From lignin to valuable products\u0026ndash;strategies, challenges, and prospects. \u003cem\u003eBioresour Technol.\u003c/em\u003e \u003cb\u003e271\u003c/b\u003e, 449\u0026ndash;461 (2019).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eGrgas, D. et al. The Bacterial Degradation of Lignin\u0026mdash;A Review. \u003cem\u003eWater\u003c/em\u003e \u003cb\u003e15\u003c/b\u003e, 1272 (2023).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eDavis, R. E. et al. 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Methods\u003c/em\u003e. \u003cb\u003e12\u003c/b\u003e, 115\u0026ndash;121 (2015).\u003c/span\u003e\u003c/li\u003e\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":false,"highlight":"","institution":"","isAcceptedByJournal":true,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"scientific-reports","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"scirep","sideBox":"Learn more about [Scientific Reports](http://www.nature.com/srep/)","snPcode":"","submissionUrl":"","title":"Scientific Reports","twitterHandle":"","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"stoa","reportingPortfolio":"Scientific Reports","inReviewEnabled":true,"inReviewRevisionsEnabled":true},"keywords":"","lastPublishedDoi":"10.21203/rs.3.rs-6335743/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-6335743/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003cp\u003eThe current paradigm in synthetic biology for lignin bioconversion platforms includes primarily bacteria and filamentous fungi. Yeast are notoriously understudied for their role in lignin degradation and utilization, despite their ubiquity in saprophytic microbial communities. A few publications report ligninolytic yeasts, but investigations to date have relied on model aromatic compounds or lignin-containing substrates replete with other carbon sources. In this work, we use a suite of analytical tools to evaluate interactions between corn stover-extracted lignin and the oleaginous yeast \u003cem\u003eCutaneotrichosporon oleaginosum\u003c/em\u003e. Notably, 2D-NMR analysis showed a significant decrease in the H-lignin component as well as resinol (β-β) and phenylcoumaran (β-5) linkages. Using super-resolution fluorescence microscopy, we demonstrated that this yeast may uptake polymeric lignin. To explore mechanisms of lignin degradation, transport, and aromatics catabolism, extensive secretomics and proteomics analyses were conducted. Compared to carbon-limited glucose and “No Carbon” controls, several putative laccases, quinone reductases, superoxide dismutases, and glyoxal/oxalate oxidases were upregulated in the lignin condition. Excitingly, two ferric reductases and a oxalate exchanger were only observed on lignin. These results indicate that \u003cem\u003eC. oleaginosum\u003c/em\u003e may perform extracellular quinone redox cycling to generate lignin-modifying reactive oxygen species. These findings enhance our understanding of lignin utilization by yeast and provide valuable insights for metabolic engineering.\u003c/p\u003e","manuscriptTitle":"Exploring the Ligninolytic Capabilities of the Oleaginous Yeast Cutaneotrichosporon oleaginosum","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2025-05-07 06:08:23","doi":"10.21203/rs.3.rs-6335743/v1","editorialEvents":[{"type":"communityComments","content":0},{"type":"decision","content":"Revision requested","date":"2025-05-28T04:55:33+00:00","index":"","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-05-27T10:18:34+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"225245639953281291000184705354003915052","date":"2025-05-13T08:48:09+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-04-28T09:44:24+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"22520951774788385118477141999163317372","date":"2025-04-15T08:24:33+00:00","index":"hide","fulltext":""},{"type":"reviewersInvited","content":"","date":"2025-04-15T06:46:52+00:00","index":"","fulltext":""},{"type":"editorAssigned","content":"","date":"2025-04-09T09:04:56+00:00","index":"","fulltext":""},{"type":"editorInvited","content":"","date":"2025-04-09T09:00:44+00:00","index":"","fulltext":""},{"type":"checksComplete","content":"","date":"2025-04-07T07:36:34+00:00","index":"","fulltext":""},{"type":"submitted","content":"Scientific Reports","date":"2025-03-29T19:43:31+00:00","index":"","fulltext":""}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"scientific-reports","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"scirep","sideBox":"Learn more about [Scientific Reports](http://www.nature.com/srep/)","snPcode":"","submissionUrl":"","title":"Scientific Reports","twitterHandle":"","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"stoa","reportingPortfolio":"Scientific Reports","inReviewEnabled":true,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"9205b10a-d610-4f39-9dec-8862377dbfd0","owner":[],"postedDate":"May 7th, 2025","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"published-in-journal","subjectAreas":[{"id":47554441,"name":"Biological sciences/Biotechnology/Proteomics"},{"id":47554442,"name":"Biological sciences/Microbiology/Fungi/Fungal biology"}],"tags":[],"updatedAt":"2026-01-19T16:47:20+00:00","versionOfRecord":{"articleIdentity":"rs-6335743","link":"https://doi.org/10.1038/s41598-026-36483-5","journal":{"identity":"scientific-reports","isVorOnly":false,"title":"Scientific Reports"},"publishedOn":"2026-01-18 16:30:04","publishedOnDateReadable":"January 18th, 2026"},"versionCreatedAt":"2025-05-07 06:08:23","video":"","vorDoi":"10.1038/s41598-026-36483-5","vorDoiUrl":"https://doi.org/10.1038/s41598-026-36483-5","workflowStages":[]},"version":"v1","identity":"rs-6335743","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-6335743","identity":"rs-6335743","version":["v1"]},"buildId":"8U1c8b4HqxoKbykW_rLl7","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}