Prospecting Araucaria-Associated Yeasts for Second-Generation Biorefineries

Abstract

Native yeasts are a promising microbial resource for the development of sustainable biorefineries. In this study, we isolated 30 yeast strains from soil, decaying wood, and tree bark in a preserved Araucaria Forest in Southern Brazil and characterized them phenotypically and taxonomically. All strains were able to grow on glucose, xylose, and cellobiose, and 50% of them could metabolize arabinose. Several isolates showed high growth rates on xylose (up to 0.47 h⁻¹) and cellobiose (up to 0.45 h⁻¹). Notably, 19 strains (63% of the analyzed yeasts) exhibited xylanase activity at 50 °C (up to 156.84 U/mL), and four strains (13%) showed significant cellulase production. β-Glucosidase activities were particularly high in permeabilized cells of CHAP-258, CHAP-277, and CHAP-278 (up to 584.33 U/mg DCW), with kinetic parameters indicating high enzymatic performance. Twelve strains (40% of the total) were classified as oleaginous, and three (10%) displayed both lipogenic and esterase activity. Lipase activity against p-nitrophenyl palmitate (pNPP) reached 55.55 U/mL in CHAP-260. Taxonomic identification revealed representatives of seven genera, including Meyerozyma, Papiliotrema, Scheffersomyces, and Sugiyamaella, with potential for biotechnological use. Overall, the biochemical diversity observed highlights the value of native yeasts from Araucaria Forests as biocatalysts for lignocellulose-based bioprocesses, particularly due to their ability to grow on pentoses, secrete hydrolytic enzymes, and accumulate lipids.

1. Introduction

Yeasts are unicellular fungi that play a pivotal role in ecosystems by contributing to the transformation of organic matter through enzymatic catalysis. Among the various enzyme classes produced by yeasts, hydrolases stand out due to their widespread industrial use. These enzymes catalyze the cleavage of covalent bonds through hydrolysis, involving water as a nucleophile, and can act on ester, glycosidic, peptide, C-N, and anhydride bonds [1,2]. Within this group, glycosyl hydrolases (GHs) are particularly significant for their application in lignocellulosic biomass degradation.

Cellulases and xylanases, two GH complexes, are essential for the hydrolysis of plant-based polymers such as cellulose and hemicellulose, respectively. These enzymes convert cellulose into cellooligosaccharides and ultimately glucose, while xylanases degrade hemicellulose into xylooligosaccharides and xylose—the major sugar constituent of xylan [3,4,5]. These enzymes are thus valuable for second-generation biofuels (2G), paper and pulp industries, and prebiotic food production [6,7,8]. In addition, recent advances have sought to integrate enzymatic routes with other technologies. For example, biomass pretreatment protocols assisted by microwaves and deep eutectic solvents have shown greater efficiency in lignin removal, thereby increasing the yield of enzymatic hydrolysis and reducing both costs and environmental impacts, while photo-thermal catalysts enable biodiesel production with a high yield and a negative carbon balance [9,10].

Despite the estimated existence of 2.2 to 3.8 million fungal species worldwide, only around 3500 yeast species have been identified, primarily within the phyla Ascomycota and Basidiomycota [11,12]. In natural environments, yeasts contribute to nutrient cycling by degrading organic matter and participating in mutualistic interactions with plants and insects [13]. In Brazil, a country spanning six major biomes—Amazon, Caatinga, Cerrado, Atlantic Forest, Pampas, and Pantanal—yeast biodiversity is both rich and underexplored. Research has shown that natural substrates such as rotting wood, plant surfaces, insect guts, and marine habitats harbor diverse yeast species with biotechnological potential [14,15,16,17].

The Atlantic Forest is one of the most studied Brazilian biomes in terms of yeast biodiversity; however, not all vegetative formations within it have received equal scientific attention. One such underexplored vegetation type in this biome is the Mixed Ombrophilous Forest, also known as the Araucaria Forest, which spans the southern states of Paraná, Santa Catarina, and Rio Grande do Sul. Previous studies have uncovered dozens of new yeast species in this biome, isolated from decomposing wood and animal feces [14,18], but none of them have covered Araucaria Forests. This ecosystem is a critically endangered biodiversity hotspot, with less than 13% of its original area remaining. It is a relict ecosystem characterized by a subtropical climate, high altitude, and winters with frequent frosts. Its floristic composition is unique, structured around the iconic Brazilian pine Araucaria angustifolia, which creates a specific microenvironment rich in decaying wood and plant litter [19]. This ecological uniqueness, combined with its conservation status, makes it a privileged but underexplored reservoir for microbial diversity. We hypothesized that yeasts adapted to this environment would possess robust metabolic capabilities for decomposing plant biomass, making them prime candidates for biotechnological applications.

Beyond saccharolytic potential, certain wild yeasts are also classified as oleaginous, capable of storing over 20% of their biomass as intracellular lipids. In this context, Yarrowia lipolytica is a notable example, with lipid contents reaching up to 70% [20,21]. Other genera known for oleaginous traits include Candida, Cryptococcus, Rhodotorula, Metschnikowia, and Lipomyces [22]. These yeasts metabolize sugars via glycolysis and the pentose phosphate pathway, producing acetyl-CoA and glycerol-3-phosphate as precursors for lipid biosynthesis. When extracellular triglycerides or long-chain fatty acids are present, hydrolysis by esterases and lipases is necessary before uptake and metabolism [23,24].

The presence of oxidative–reductive enzymes, desaturases, elongases, esterases, and lipases contributes to the lipid-modifying and degrading capacities of oleaginous yeasts. Esterases hydrolyze short-chain esters, while lipases target long-chain and water-insoluble substrates [25,26]. These enzymes are commercially relevant for applications ranging from wastewater treatment to food processing and biofuel production. Candida rugosa and Candida antarctica are currently exploited as industrial lipase sources [27,28,29].

Recent bioprospecting studies have revealed that wild yeast strains isolated from coffee fruits, cashew peels, and sugarcane straw exhibit esterase and lipase activities when cultivated on low-cost substrates [30,31]. Therefore, exploring natural environments such as the Araucaria Forest not only enriches our understanding of microbial biodiversity but also identifies promising candidates for biocatalysis and sustainable industrial applications [13].

Based on this context, the present study aimed to isolate, identify, and characterize wild yeasts from the Araucaria Forest, with a focus on screening glycosyl hydrolases, esterases, and lipases. The overarching goal was to evaluate their potential for lignocellulosic biomass valorization and their applicability in biotechnological processes aligned with the principles of the circular economy. To the best of our knowledge, our study represents the first comprehensive effort to isolate and characterize yeasts from the Mixed Ombrophilous Forests (Araucaria forests) of Brazil, highlighting its novelty in exploring this unique and underexplored ecosystem. Through this study, we identified strains with exceptional capabilities for pentose assimilation, high-temperature xylanase and β-glucosidase activity, and simultaneous oleaginous and lipolytic traits.

2. Materials and Methods

2.1. Yeast Isolation

The sampling site selected for this study was the vegetation type known as Mixed Ombrophilous Forest (Araucaria Forest), located within the Chapecó National Forest. The sampling zone is located at coordinates 27°5′46.00″ S 52°47′14.60″ W and illustrated in Figure 1. Although this is not a primary forest area, it undergoes managed reforestation to prevent the spread of invasive species and ensure the continued planting of Araucaria angustifolia trees for ecological restoration [32]. Sampling was conducted on a single day (4 April 2023, autumn in the Southern Hemisphere) during the afternoon, under an average ambient temperature of 20 °C.

Samples were collected using sterile 50 mL Falcon tubes, sterile forceps, and disposable gloves. Material was obtained from three sources: (i) the litter layer (rotting leaves and twigs, including Araucaria cones), (ii) superficial soil, and (iii) bark fragments of A. angustifolia. Sampling was carried out at ten points surrounding the reference location shown in Figure 1. Samples were stored in Falcon tubes and immediately transported to the laboratory for processing.

For yeast isolation, 125 mL cotton-plugged Erlenmeyer flasks containing 25 mL of yeast nitrogen base (YNB, 6.7 g/L without amino acids, pH 5.0—Sigma-Aldrich, Milwaukee, WI, USA, cod. Y0626) supplemented with 10 g/L xylose and 0.02 g/L chloramphenicol were used. Approximately 1 g of each sample was inoculated into the flasks, which were then incubated at either 11 °C or 30 °C under agitation (145 rpm) until visible turbidity was observed. Isolation procedures followed the protocol described by Tadioto et al. [34]. The isolated yeast strains (listed in

Table 1. Yeast strains isolated in this study.

Strain a	Substrate and Temperature of Isolation	Species	LSU GenBank Code b or Reference
CHAP-254	Soil—11 °C	Peterozyma sp.	-
CHAP-255	Soil—11 °C	Peterozyma sp.	PV994468
CHAP-258	Soil—11 °C	Scheffersomyces coipomensis	-
CHAP-259	Soil—11 °C	Peterozyma sp.	-
CHAP-260	Araucaria’s bark—11 °C	Papiliotrema laurentii	-
CHAP-261	Araucaria’s bark—11 °C	Papiliotrema laurentii	PV994616
CHAP-262	Araucaria’s bark—11 °C	Yamadazyma dushanensis	PV994689
CHAP-263	Araucaria’s bark—11 °C	Peterozyma sp.	-
CHAP-265	Litter—11 °C	Peterozyma sp.	-
CHAP-266	Litter—11 °C	Peterozyma sp.	-
CHAP-267	Litter—11 °C	Peterozyma sp.	-
CHAP-268	Litter—11 °C	Peterozyma sp.	-
CHAP-270	Litter—11 °C	Papiliotrema terrestris	PX022965
CHAP-271	Litter—11 °C	Peterozyma sp.	-
CHAP-272	Litter—11 °C	Peterozyma sp.	-
CHAP-273	Litter—11 °C	Peterozyma sp.	-
CHAP-274	Litter—11 °C	Hannaella luteola	PV995078
CHAP-275	Litter—11 °C	Scheffersomyces stipitis	PV995079
CHAP-276	Soil—30 °C	Meyerozymacarpophila	PV995110
CHAP-277	Soil—30 °C	Meyerozyma caribbica	-
CHAP-278	Soil—30 °C	Scheffersomyces coipomensis	PV995111
CHAP-279	Araucaria’s bark—30 °C	Meyerozyma carpophila	-
CHAP-280	Araucaria’s bark—30 °C	Meyerozyma caribbica	-
CHAP-281	Araucaria’s bark—30 °C	Meyerozyma caribbica	-
CHAP-282	Litter—30 °C	Meyerozyma caribbica	PV995221
CHAP-283	Litter—30 °C	Sugiyamaella smithiae	PV995348
CHAP-284	Litter—30 °C	Sugiyamaella boreocaroliniensis	PV995352
CHAP-285	Litter—30 °C	Meyerozyma sp.	PV995365
CHAP-286	Litter—30 °C	Sugiyamaella sp.	PV995366
CHAP-287	Litter—30 °C	Meyerozyma carpophila	-

^a In the present study, we chose to sequence yeast strains from isolated branches and the best-performing strain from each cluster of the phylogenetic tree (underlined). The remaining strains were identified based on the formation of clusters in the dendrogram derived from DNA fingerprinting. For additional details, see Section 2.8, Section 2.9 and Section 3.4. ^b Sequences of the D1/D2 variable domains of the large subunit (LSU) rRNA gene deposited in GenBank (NCBI—https://www.ncbi.nlm.nih.gov/, accessed on 3 July 2025).

) were stored in 30% glycerol at −80 °C for long-term preservation.

2.2. Micro-Scale Cultivation

Prior to the experimental assays, the yeast isolates were pre-cultivated in 50 mL Erlenmeyer flasks containing 10 mL of YPD medium (10 g/L yeast extract, 20 g/L peptone, 20 g/L glucose) for 48 h at 30 °C under orbital shaking at 145 rpm. Subsequently, 5 µL of each pre-culture was inoculated into individual wells of 96-well microplates containing 100 µL of YNB medium supplemented with 20 g/L of a single carbohydrate.

The carbon sources tested were glucose, xylose, cellobiose, and arabinose. The plates were sealed and incubated in a Tecan GENios microplate reader (Tecan Group Ltd., Männedorf, Switzerland) at 30 °C with linear shaking at 160 rpm (1 mm amplitude). Optical density (OD600) readings were taken every 15 min for real-time monitoring of growth dynamics, following the protocol established by Albarello et al. [35].

The maximum specific growth rate (µmax) was calculated using geometric modeling in Microsoft Excel spreadsheets, as described by Albarello et al. [35].

2.3. Glycosyl Hydrolase Activities

2.3.1. Cellulase and Xylanase Activity

To assess the activity of hydrolytic enzymes, the yeast strains were cultivated in 50 mL Erlenmeyer flasks containing 10 mL of induction medium, under agitation at 145 rpm and a constant temperature of 30 °C for 96 h. The induction medium consisted of YP (10 g/L yeast extract and 20 g/L peptone, pH 5.0) supplemented with either 10 g/L carboxymethyl cellulose (CMC) or xylan to induce cellulase and xylanase production, respectively.

Enzymatic activities were measured following the protocol described by Albarello et al. [35]. Since these enzymes are secreted by the cells to act extracellularly, culture medium aliquots were collected and centrifuged (9000 rpm, 3 min). Subsequently, 10 μL of the supernatants from each condition was transferred to 96-well microplates in triplicate. Negative controls consisted of sterile, uninoculated medium aliquoted similarly. Then, 15 μL of 0.15 M succinate-Tris buffer (pH 5.0) containing 10 g/L of either CMC or xylan was added to each well to initiate hydrolysis.

The plates were sealed and incubated for 1 h at either 30 °C or 50 °C in a thermostatically controlled water bath. After incubation, the concentration of reducing sugars released was determined using the dinitrosalicylic acid (DNS) method, following Santos et al. [36]. Calibration curves were assembled in separate wells using increasing concentrations of glucose or xylose (0–2 g/L). Subsequently, 25 μL of DNS reagent (1% 3,5-dinitrosalicylic acid and 30% sodium potassium tartrate in 0.4 M NaOH) was added to each well.

The plates were resealed and incubated for 5 min at 100 °C to complete the redox reaction. After cooling, 330 μL of distilled water was added to each well, and absorbance was read at 490 nm using a microplate reader. Enzymatic activity was expressed in international units (U), defined as the amount of enzyme required to release 1 nmol of reducing sugar per minute under the assay conditions.

2.3.2. β-Glucosidase Activity in Permeabilized Yeast Cells

All 30 yeast strains were cultivated in 50 mL Erlenmeyer flasks containing 10 mL of YP medium supplemented with 20 g/L of cellobiose, under agitation at 145 rpm and incubation at 30 °C. Cultures were maintained until an optical density (OD570) of approximately 10 was reached. Subsequent steps followed the protocol described by Barrilli et al. [37]. For the enzymatic assay, 100 μL of a cell suspension containing 20 g/L biomass from the culture was used and subjected to permeabilization. Approximately 2 mg of cells (10 μL) was washed with 0.5 mL of 0.1 M MOPS-NaOH buffer, pH 6.8 (buffer A), followed by permeabilization with 200 μL of buffer A supplemented with 20% glycerol, 1 mM EDTA, 1 mM DTT (buffer B), and 12 μL of a toluene/ethanol/Triton X-100 10% (1:4:1 v/v) solution. Cells were vigorously vortexed for 1 min. After permeabilization, cells were washed with buffer B and resuspended in 1 mL of buffer A.

For the assay, 50 μL of permeabilized cells was transferred into five microtubes (two of which were boiled at 100 °C to serve as negative controls). Then, 400 μL of buffer A (1.25×) and 50 μL of 20 mM p-nitrophenyl-β-D-glucopyranoside (pNPβG) were added to all tubes.

Reactions were incubated at 30 °C for 10 min, then stopped by boiling the samples at 100 °C followed by centrifugation at 9000 rpm for 3 min. For pNPβG assays, 500 µL of distilled water was added to each supernatant, and absorbance was measured at 400 nm (ε = 7.28 mM⁻¹∙cm⁻¹) to estimate p-nitrophenol release.

For kinetic analyses, pNPβG was replaced with cellobiose at varying concentrations (12.5, 25, 50, 100, 200, and 400 mM) in buffer A. For assays using cellobiose, 10 µL of supernatant was combined with 1 mL of reagent from the Gold Analysa commercial kit (Belo Horizonte, Brazil) to determine the glucose release. Kinetic parameters were calculated using the non-linear regression Michaelis–Menten model available in GraphPad Prism^® v8.0.2 (StatSoft Inc., Tulsa, OK, USA).

One unit of enzymatic activity (U) was defined as the amount of enzyme that releases 1 nmol of p-nitrophenol or glucose per minute.

2.3.3. β-Glucosidase Activity in Culture Supernatants

For the extracellular β-glucosidase activity assay, 1 mL of culture medium was centrifuged at 9000 rpm for 3 min. Then, 50 μL of supernatant was transferred into five microtubes (two of which were boiled at 100 °C as negative controls). To each tube, 50 μL of 0.3 M succinate-Tris buffer (pH 5.0) and 50 μL of 6 mM pNPβG solution were added.

Reactions were incubated at 50 °C for 10 min. After incubation, reactions were terminated via boiling. Then, 200 µL of 2 M sodium bicarbonate was added, followed by transfer of 200 µL of this mixture to tubes containing 1 mL of distilled water. Absorbance was measured at 400 nm (ε = 7.28 mM⁻¹∙cm⁻¹) to quantify p-nitrophenol.

One unit of enzymatic activity (U) was defined as the amount of enzyme that releases 1 nmol of p-nitrophenol per minute.

2.4. Screening of Oleaginous Yeasts

For lipid accumulation screening, 1 μL of pre-cultured yeast cells was spot-inoculated onto Petri dishes containing solid medium composed of 23 g/L glucose, 0.3 g/L peptone, 0.5 g/L yeast extract, 7 g/L monobasic potassium phosphate, 1.06 g/L dibasic potassium phosphate (anhydrous), 3.07 g/L magnesium sulfate heptahydrate, and 20 g/L agar. The medium was supplemented with a 100× stock solution of Rhodamine B (1 g/L in absolute ethanol). The plates were incubated at 30 °C for 48 h. Oleaginous yeasts developed pink-colored colonies due to the dye’s affinity for intracellular lipids, while non-oleaginous strains remained uncolored [38].

The strain Saccharomyces cerevisiae PE-2, a known non-oleaginous yeast, was used as a negative control, and the oleaginous Yarrowia lipolytica UFMG-CM-Y6114 served as the positive control. ImageJ software (v1.54g, http://imagej.org) was used to measure the average color intensity of the colony area. The value for strain PE-2 was assigned as 0, and UFMG-CM-Y6114 as 1. Based on linear interpolation, the oleaginous score (Ya) was calculated:

Ya = Y1 + [(Y2 − Y1) × ((Xa − X1)/(X2 − X1))]

where:

• Ya: Sample’s lipid index (0–1 scale);
• Y1: Value assigned to the negative control (0);
• Y2: Value assigned to the positive control (1);
• Xa: Mean color intensity of the sample;
• X1: Mean color intensity of the negative control;
• X2: Mean color intensity of the positive control.

Based on this scale, yeast strains were classified into four categories: 0.00–0.25, non-oleaginous; 0.26–0.50, slightly oleaginous; 0.51–0.75, moderately oleaginous; 0.76–1.00, highly oleaginous.

2.5. Screening for Extracellular Esterase-Producing Yeasts

Extracellular esterase activity was assessed on Petri dishes containing 20 g/L agar, 10 g/L peptone, 5 g/L sodium chloride, 4 g/L calcium chloride dihydrate, and 10 g/L Tween 80. The plates were sectioned into quadrants, and strains were spot-inoculated using an inoculation needle. Positive esterase activity was indicated by the formation of a white halo around colonies, caused by the precipitation of calcium salts resulting from the hydrolysis of ester bonds in Tween 80 [39]. Halo diameters were measured using ImageJ software (v1.54g, http://imagej.org).

The enzymatic index (EI) was calculated as the ratio of the halo diameter (D) to the colony diameter (EI = D_halo/D_colony).

2.6. Screening for Extracellular Lipase Activity

Strains with esterase enzymatic index values greater than 2 were cultivated for 96 h at 30 °C with shaking (145 rpm) in 125 mL Erlenmeyer flasks containing 25 mL of medium composed of 20 g/L peptone, 10 g/L yeast extract, 2 g/L glucose, and 2% (v/v) olive oil. At 48 h and 96 h, 1 mL samples were collected and centrifuged. Supernatants were transferred to fresh tubes, and 100 µL was distributed into four microtubes. One tube received 500 µL of absolute ethanol to deactivate the enzyme and served as a negative control.

Subsequently, 900 µL of a pre-prepared emulsion was added. This emulsion was prepared by mixing 10 mL of 3 mg/mL p-nitrophenyl palmitate (pNPP) in isopropanol with 90 mL of a solution containing 100 mg of gum arabic and 400 mg of Triton X-100 in 2.5 mM Tris-HCl buffer (pH 8.0), following Bussamara et al. [25].

Reactions were incubated at 37 °C for 30 min in a thermostatically controlled water bath. The reaction was stopped by adding 500 µL of absolute ethanol. Absorbance was measured at 410 nm (ε = 18.3 mM⁻¹∙cm⁻¹) to quantify p-nitrophenol release. One unit (U) of lipase activity was defined as the amount of enzyme releasing 1 nmol of p-nitrophenol per minute under the assay conditions.

2.7. Statistical Analysis

The enzymatic activity data were subjected to one-way analysis of variance (ANOVA) to test the null hypothesis, using GraphPad Prism^® v8.0.2 (StatSoft Inc.), in order to determine whether significant differences existed among the tested groups (p < 0.05). When the ANOVA results indicated statistical significance, Tukey’s post-hoc multiple comparison test was performed to identify which pairwise comparisons exhibited significant differences (p < 0.05). The results are presented as averages (based on triplicates) along with the corresponding standard deviation.

2.8. Genetic Fingerprinting of Yeast Isolates and UPGMA-Based Similarity Analysis

Yeast cells were pre-cultivated in YPD medium at 30 °C under agitation (145 rpm) for 12 h. For genomic DNA extraction, a 100 μL aliquot of culture was transferred to three microtubes and centrifuged at 9000 rpm for 15 min at 4 °C. The supernatant was discarded, and cells were washed with Milli-Q water. After a second centrifugation, the residual water was discarded. DNA extraction followed the protocol described by Lõoke, Kristjuhan, and Kristjuhan [40], using 0.2 M lithium acetate with 1% SDS to permeabilize cells, followed by precipitation with absolute ethanol and 70% ethanol.

The extracted DNA was subjected to a polymerase chain reaction (PCR) for amplification of simple sequence repeats (SSRs), also known as microsatellites. Amplification was carried out in PCR tubes containing 0.5 µL of the (GTG)₅ primer (5′-GTGGTGGTGGTGGTG-3′), 2.5 μL of 10× Mg²⁺ Plus PCR Buffer (Sinapse Biotecnologia, São Paulo, Brazil), 0.5 μL of 10 mM dNTPs, 0.25 μL of 500 U Taq Polymerase (Sinapse Biotecnologia), 19.5 μL of Milli-Q water, and 1 μL of yeast genomic DNA.

The PCR conditions included an initial denaturation step at 94 °C for 3 min, followed by 32 cycles of denaturation at 94 °C for 30 s, annealing at 60 °C for 30 s, and extension at 72 °C for 90 s. A final extension step at 72 °C for 10 min was performed after the last cycle. PCR products were analyzed via electrophoresis [on 1.5% agarose gels prepared in 0.5× TBE buffer (pH 8.0) containing 3 μL of 10,000× SafeDye Nucleic Acid Stain (Cellco, San Francisco, CA, USA) per gel [41] and compared to the previously identified yeast strains presented in Table S1.

Band pattern analysis and annotation were performed using Bio-Rad’s Image Lab software (version 6.1.0). Band sizes were estimated using molecular weight markers of 100 bp (Ludwig, Alvorada, Brazil) and 1000 bp (Sinapse Biotecnologia). Based on the observed electrophoretic profiles, a binary presence/absence matrix was constructed for all strains, assigning “1” to indicate presence and “0” to indicate absence of a band [42].

This binary matrix was converted into the FASTA format and submitted to the UPGMA clustering algorithm available on the DendroUPGMA website (http://genomes.urv.es/UPGMA/index.php, accessed on 30 June 2025) developed by Garcia-Vallvé and Puigbo. The similarity matrix, based on the Jaccard index, was used to generate a dendrogram in the Newick format. The resulting phylogenetic tree was visually constructed using the Interactive Tree of Life (iTOL v7) platform (https://itol.embl.de, accessed on 30 June 2025).

2.9. Taxonomic Analysis of Yeasts

The yeast strains from individual branches, as well as the best-performing strain from each cluster of the phylogenetic tree, were taxonomically identified by analyzing the D1/D2 variable domains of the large subunit (LSU) rRNA gene, amplified by the primers pair LR0R (5′-ACCCGCTGAACTTAAGC-3′) and LR16 (5′-TTCCACCCAAACACTCG-3′) [43]. The amplified DNA was concentrated, purified, and sequenced in an automatic capillary DNA sequencer 3500 × l (Applied Biosystems, Foster City, CA, USA) using Big Dye 3.1 reagent. The obtained sequences were compared with those from holotype strains in the NCBI Genbank using the basic local alignment search tool (BLAST, v2.16.0, at http://www.ncbi.nlm.nih.gov, accessed on 3 July 2025), considering only hits with a query cover of ≥99%, as described in a previous study [44]. After identification, the LSU sequences were deposited in the NCBI GenBank with the accession numbers presented in Table 1.

3. Results

3.1. Specific Growth Rates

The performance of yeasts in second-generation biorefineries—which use lignocellulosic residues as feedstock—depends, among other factors, on their ability to assimilate the main sugars present in plant biomass, particularly glucose, xylose, cellobiose, and arabinose. Therefore, in order to assess their phenotypic profiles in response to these key carbon sources, the thirty yeast strains isolated from the Araucaria Forest (see Table 1) were cultivated in microscale assays using minimal media containing these individual sugars. All strains were able to grow on the first three sugars (Figure 2).

Strains CHAP-255, -258, -265, -266, -277, and -281 exhibited the highest specific growth rates on xylose, with μmax values ranging from 0.42 to 0.47 h⁻¹. Given that the isolation procedures employed xylose-containing media (since xylose is the second-most abundant sugar in lignocellulosic biomass after glucose), this result was anticipated, as it indicates the selective enrichment of pentose-assimilating yeasts. Nonetheless, some strains also stood out for their performance in cellobiose-based media, an important disaccharide commonly found in lignocellulosic hydrolysates [45,46]. This was the case for strains CHAP-263, -265, -267, and -287, which presented μmax values between 0.40 and 0.45 h⁻¹ on cellobiose. Moreover, fifteen strains were able to grow on arabinose. Among them, isolates CHAP-261 and CHAP-276 exhibited the highest specific growth rates, reaching 0.39 and 0.35 h⁻¹, respectively. It is important to note that these two strains were also able to grow on glucose, xylose, and cellobiose. However, their maximum specific growth rates on these sugars (0.29, 0.27, and 0.25 h⁻¹ for strain CHAP-261; and 0.29, 0.31, and 0.23 h⁻¹ for strain CHAP-276, on glucose, xylose, and cellobiose, respectively) were lower than those observed on arabinose.

3.2. Prospecting Yeasts with High GH Activities

To assess the cellulolytic activity of yeast isolates from the Araucaria Forest, culture supernatants obtained from media containing carboxymethylcellulose (CMC) were used as enzymatic extracts. These extracts were added to buffer solutions containing 1% CMC and incubated at 30 °C and 50 °C. No cellulolytic activity was detected at 30 °C, but nineteen isolates exhibited promising results at 50 °C (Figure 3A). The highest cellulolytic activities were observed in strains CHAP-255, -282, -285, and -286, which showed activities of 31.39 ± 11.24, 37.38 ± 11.68, 29.10 ± 11.30, and 39.99 ± 13.41 U/mL, respectively.

Regarding xylanolytic activity, most yeast strains displayed similar activity levels, with enhanced performance at 50 °C (Figure 3B). At 30 °C, the same strains also exhibited activity, though on average ~30% lower than at the higher temperature (Figure 3C). At 50 °C, CHAP-274 stood out with the highest activity (156.84 ± 39.58 U/mL), which was 50% higher than the second highest (CHAP-276, 98.04 ± 4.05 U/mL) and double the value of the third highest (CHAP-255, 78.76 ± 34.25 U/mL). Notably, CHAP-274 also showed outstanding activity at 30 °C, with 100.27 ± 24.44 U/mL.

Given the importance of cellobiose hydrolysis for efficient utilization of lignocellulosic residues, β-glucosidase activity was also evaluated. Culture supernatants and permeabilized yeast cells were used as enzyme extracts to hydrolyze pNPβG, a chromogenic analog of cellobiose. The supernatants allowed identification of strains capable of secreting β-glucosidase into the medium, as previously reported for Cyberlindnera rhodanensis [47]. Permeabilized cells were used to assess intracellular and/or periplasmic β-glucosidase activity, as described for Sungouiella (Candida) pseudointermedia and Yarrowia lipolytica [37,48]. The enzymatic activity in supernatants was tested at 50 °C, while assays with permeabilized cells were performed at 30 °C. Only a few strains secreted enzymes into the medium (Figure 4A), but all exhibited β-glucosidase activity associated with the cells (Figure 4B).

Among the permeabilized cells, only three strains showed activity below 100 U/mg DCW. One of these was CHAP-270, which also lacked extracellular β-glucosidase activity, possibly explaining its low specific growth rate in cellobiose (µmax = 0.15 ± 0.01 h⁻¹). In contrast, although CHAP-283 had a higher enzymatic activity (202.02 ± 73.68 U/mg DCW), it still displayed a low µmax (0.14 ± 0.01 h⁻¹), suggesting possible inefficiencies in β-glucoside transport. CHAP-283 may harbor low-affinity transporters, limiting cellobiose uptake and thus growth rate. In fact, previous studies from our group have already reported that disaccharide transport across the membrane can be the limiting factor in carbohydrate metabolism. These studies show that intracellular hydrolytic activity can be 20 to 100 times higher than the rate at which the sugar crosses the plasma membrane [4,37].

Conversely, CHAP-258, CHAP-277, and CHAP-278—showing the highest β-glucosidase activities (531.15 ± 41.90, 474.72 ± 40.47, and 584.33 ± 22.96 U/mg DCW, respectively)—also achieved high µmax values on cellobiose (0.35 ± 0.02, 0.29 ± 0.01, and 0.29 ± 0.07 h⁻¹, respectively). Interestingly, all three isolates were obtained from soil samples. CHAP-265 presented the highest µmax (0.45 ± 0.01 h⁻¹) and notable β-glucosidase activity (303.96 ± 28.81 U/mg DCW).

Given the outstanding performance of CHAP-258, CHAP-277, and CHAP-278, these strains were selected for β-glucosidase kinetic assays using cellobiose as the substrate (cellobiose concentrations ranged from 12.5 to 400 mM). As shown in Figure 5, the three strains displayed distinct enzymatic profiles, reflected in their Michaelis–Menten kinetic parameters. Vmax values differed significantly (p < 0.05), with CHAP-278 reaching the highest rate (723.40 ± 79.09 U/mg DCW). Km values indicated similar substrate affinities between CHAP-278 and CHAP-258 (23.93 ± 5.16 and 20.72 ± 6.34 mM, respectively).

3.3. Oleaginous and Lipolytic Yeasts

In the context of second-generation biorefineries, the ability to accumulate lipids and hydrolyze ester bonds is highly desirable. Therefore, this study also focused on screening oleaginous yeast strains capable of producing esterases and lipases. Initially, lipid accumulation was assessed by quantifying the color intensity of colonies grown for 48 h on rhodamine B-containing agar plates. Colonies accumulating lipids were stained pink, while non-oleaginous strains remained white. Color intensity values were quantified using ImageJ software. Y. lipolytica UFMG-CM-Y6114 was used as a positive control due to its well-documented oleaginous nature, while S. cerevisiae PE-2 served as the negative control. Among the thirty isolates, seven were classified as highly oleaginous and five as moderately oleaginous, totaling twelve lipogenic strains (Figure 6A). CHAP-263, CHAP-265, and CHAP-270 stood out with scale values of 0.88, 0.92, and 1.00, respectively.

Esterase activity was evaluated based on halo formation on solid media containing Tween 80 as the substrate. After 48 and 96 h of incubation, colony and halo diameters were measured using ImageJ, and enzymatic index (EI) values were calculated. Only three isolates—CHAP-260, CHAP-261, and CHAP-270—showed halos at 48 h. However, after 96 h, esterase-secreting strains increased to nine (Figure 6B), and the EI also increased in the three strains already positive at 48 h.

Among the highly oleaginous strains (Figure 6A), only three—CHAP-260, CHAP-270, and CHAP-274—showed esterase activity (Figure 6B). This was unexpected, as lipid-accumulating yeasts are presumed to produce esterases to metabolize stored lipids. Thus, to evaluate their potential on a long-chain ester substrate, eight of the nine esterase-positive strains (EI ≥ 2) were subjected to p-nitrophenyl palmitate (pNPP) assays on days 2 and 4 of cultivation.

At 48 h, only two strains hydrolyzed pNPP: CHAP-260 and CHAP-261, with activities of 6.08 ± 0.96 and 4.51 ± 1.37 U/mL, respectively (Figure 7). By 96 h, all eight strains showed lipase activity, with CHAP-260 and CHAP-261 again being the top producers—now with activities of 55.55 ± 4.06 and 45.06 ± 2.45 U/mL, respectively.

3.4. Taxonomic Identification of Yeasts

To cluster the isolates by genetic similarity, PCR was performed using the primer (GTG)₅, followed by electrophoresis to generate DNA fingerprints in comparison with previously identified strains (Figure S1 and Table S1). Band patterns were analyzed using the UPGMA method, resulting in the dendrogram shown in Figure 8. The dendrogram revealed five clusters: Cluster I with three isolates, Cluster II with eleven, Cluster III with four isolates plus Meyerozyma caribbica CHAP-087, Cluster IV with two, and Cluster V with two isolates plus Papiliotrema laurentii CHAP-158. The remaining isolates were individually positioned on distinct branches.

Following this clustering, we selected one representative strain from each branch of the phylogenetic tree for sequencing (Figure 8). Within each cluster, the strain chosen was the one that exhibited the best overall performance, considering specific growth rate, enzymatic activities, and lipid accumulation. The D1/D2 domains of the LSU rDNA from these selected strains were sequenced, and the resulting sequences were deposited in GenBank (Table 1). Subsequently, they were compared with reference sequences from holotype strains previously available in GenBank. The top three identity matches for each strain are displayed in Table S2. Based on these analyses, strain CHAP-261 was assigned to Papiliotrema laurentii, CHAP-262 to Yamadazyma dushanensis, CHAP-270 to Papiliotrema terrestris, CHAP-274 to Hannaella luteola, CHAP-275 to Scheffersomyces stipitis, CHAP-276 to Meyerozyma carpophila, CHAP-278 to Scheffersomyces coipomensis, CHAP-282 to Meyerozyma caribbica, CHAP-283 to Sugiyamaella smithiae, and CHAP-284 to Sugiyamaella boreocaroliniensis. Conversely, we did not assign species-level epithets to strains CHAP-255, CHAP-285, and CHAP-286 due to either a percent identity below 99% (as in the case of CHAP-255 and CHAP-286) or a high similarity to two or more species (as observed for CHAP-285). Therefore, these strains were identified only at the genus level: Peterozyma sp. (CHAP-255), Meyerozyma sp. (CHAP-285), and Sugiyamaella sp. (CHAP-286). Future studies may further analyze these strains to confirm their species identity or determine whether they represent novel taxa.

Interestingly, strains CHAP-282 and CHAP-261—identified in this study as M. caribbica and P. laurentii, respectively—clustered with other strains previously identified as belonging to these same species by our group (see Clusters III and V in Figure 8) [34]. This finding confirms, as expected, the effectiveness of the clustering approach based on simple sequence repeats (SSRs). Indeed, the same method has been successfully employed by other authors [49,50]. Accordingly, we classified the three strains in Cluster I as Meyerozyma carpophila, the eleven strains in Cluster II as Peterozyma sp., all strains in Cluster III as M. caribbica, both strains in Cluster IV as Scheffersomyces coipomensis, and the strains in Cluster V as P. laurentii (Table 1).

4. Discussion

This study aimed to prospect yeast strains with phenotypes suitable for second-generation biorefineries. We specifically sought to investigate whether native yeasts from the Araucaria Forest could potentially transform residual plant biomass. To this end, thirty yeast strains were isolated from a preserved forest area within a conservation unit in southern Brazil. The biochemical characterization began with the determination of maximum specific growth rates in minimal media containing key sugars found in lignocellulosic hydrolysates.

Although all isolates were capable of utilizing xylose as a carbon source, five strains (CHAP-255, -258, -265, -266, -277, and -281) stood out, with μmax values ranging from 0.42 to 0.47 h⁻¹—nearly twice the value reported for the xylose-fermenting species Scheffersomyces stipitis (0.23 h⁻¹) in batch cultures [51]. In media containing the other tested pentose (arabinose), fifteen strains could grow. Among them, CHAP-261 and CHAP-276 achieved the highest specific growth rates, at 0.39 and 0.35 h⁻¹, respectively. These results suggest potential application of these strains in lignocellulosic residue valorization, especially for the production of compounds such as arabitol (a sweetener) or 2G ethanol [52,53].

Microbial application in biorefineries also depends on enzymatic performance, particularly glycosyl hydrolases (GHs) that catalyze the breakdown of plant polysaccharides. Four isolates (CHAP-255, -282, -285, and -286) showed cellulolytic activities consistent with prior reports: 31.39 ± 11.24, 37.38 ± 11.68, 29.10 ± 11.30, and 39.99 ± 13.41 U/mL, respectively. A previous study from our group that screened 46 yeast isolates from the gut of Spodoptera frugiperda reported similar results, with values up to 60 U/mL [35]. Similarly, Ko and Park [54] genetically modified Kluyveromyces lactis to express an endoglucanase from Monochamus saltuarius, achieving 30 U/mL after 30 h of cultivation. However, as expected, these yeast enzymatic activities are generally much lower than those of filamentous fungi; for example, Trichoderma reesei can reach over 1200 U/mL using CMC as substrate [55].

Xylanase activity, on the other hand, was notably high, especially at 50 °C, where activities were up to 56% greater than at 30 °C. Although 50 °C is not optimal for yeast growth, it is commonly cited as the optimal temperature for this enzyme activity [56]. Xylanolytic activities in our supernatants were comparable to those reported for Papiliotrema laurentii (205.50 U/mL) isolated from the gut of caterpillars [35]; Sugiyamaella smithiae (110.00 U/mL), Vishniacozyma taibaiensis (150.00 U/mL), and P. laurentii (110.00 U/mL) isolated from termites [57]; and for Sugiyamaella valenteae (137.00 U/mL) and Sugiyamaella xylolytica (102.00–111.00 U/mL) isolated from decaying wood and sugarcane biomass [58]. On the other hand, similar to what was found for cellulolytic activities, our isolates exhibited xylanolytic activities lower than those of some filamentous fungi, such as the Aspergillus species A. tubingensis and A. fumigatus, isolated from soil and plant residues—the xylanolytic activity for these fungi varied between 18,900 and 32,290 U/mL at 60 °C [59].

Despite lower enzymatic activity values compared to filamentous fungi, yeast-derived xylanases and cellulases have potential biotechnological applicability in consolidated bioprocesses (CBPs). In this process, yeasts must not only produce enzymes to hydrolyze lignocellulose but also be capable of fermenting diverse carbohydrates [60]. For example, our isolated M. carpophila CHAP-276 not only demonstrated that it could grow on the four carbohydrates that we analyzed (glucose, xylose, cellobiose, and arabinose—all present in lignocellulosic biomass), but also showed considerable xylanolytic activity (98.04 ± 4.05 U/mL), which makes it interesting for a consolidated bioprocess using hemicellulose as raw material. For comparison, an industrial S. cerevisiae (derived from the PE-2 strain) expressing an endoxylanase from the fungus Trichoderma reesei [at an enzymatic activity of 140 μmol/min/L or 140 nmol/min/mL (140 U/mL)] was able to hydrolyze 48.7% of a corncob hemicellulosic liquor, generating ~12 g/L xylose [61]. Furthermore, yeast xylanases have been applied to generate xylooligosaccharides with prebiotic properties and to produce xylitol from agricultural residues [62].

Among the GHs analyzed, β-glucosidases were particularly significant, outperforming prior results from our group, such as those in Barrilli et al. [37], which reported activities around 200 U/mg DCW for two S. pseudointermedia strains from rotting wood. In a more recent study, we overexpressed a periplasmic β-glucosidase (BGL1) from Y. lipolytica and an intracellular β-glucosidase (BGL7) from Spathaspora passalidarum in S. cerevisiae and obtained 300–450 U/mg DCW [4]. In comparison, it is worth noting that CHAP-258, CHAP-277, and CHAP-278 reached 531.15 ± 41.90, 474.72 ± 40.47, and 584.33 ± 22.96 U/mg DCW, respectively. Furthermore, kinetic analyses revealed exceptionally high Vmax values—up to 3.9 times greater than those reported by Barrilli et al. [37]. These results thus reinforce the potential of Araucaria Forest yeasts for bioprocesses based on lignocellulosic residues.

Interestingly, the three most promising β-glucosidase-producing strains were isolated from soil samples collected near Araucaria angustifolia trees. Soil is indeed a rich matrix for enzyme prospection due to its high content of plant-derived organic matter, which is decomposed by microbial enzymes releasing glucose for microbial metabolism [63,64]. For example, metagenomic studies of Amazonian soils led to the identification and cloning of a β-glucosidase with high activity and affinity [63]. Similarly, Candida sorbosivorans, isolated from vineyard soil, exhibited elevated β-glucosidase activity, reinforcing soil as a promising source for enzyme discovery [65].

Oleaginous yeasts are also highly desirable for biorefineries. The screening method used here was proposed by Niehus et al. [38] as a rapid way to detect lipid-accumulating yeasts. In their study, 124 isolates were tested, and 50 were classified as oleaginous—31 highly and 19 moderately. This proportion is similar to ours. Although not all strains were taxonomically identified in the study by Niehus et al. [38], some belonged to Candida, Debaryomyces, and Yarrowia. Notably, some of our isolates accumulated lipids to the same extent as our positive control, Y. lipolytica.

In a circular bioeconomy context, esterases and lipases are also valuable, with applications ranging from wastewater treatment to biofuel and food production [66,67]. Souza et al. [39], using the same esterase screening method that we employed, identified Leucosporidium muscorum and Mrakia blollopis among Antarctic yeasts, reporting enzymatic indices comparable to ours. Likewise, Chaib et al. [68] screened isolates from soil, hot springs, olive pomace oil, Pistacia lentiscus resin, and rinse water, identifying 17 strains with moderate to high esterase activity, especially Geotrichum candidum P027.

Strains CHAP-260 and CHAP-261, which showed the highest lipase activities (55.55 ± 4.06 U/mL and 45.06 ± 2.45 U/mL, respectively), also ranked among the top esterase producers. In contrast, CHAP-270 and CHAP-278, which showed high enzymatic indices in the Tween 80 assay, exhibited low lipolytic activity when tested with pNPP, a fatty acid analog. This suggests their esterases have limited lipolytic activity. Furthermore, only three of the twelve yeasts which we found to be lipogenic had esterase activity. These contrasting profiles highlight the diversity of lipase and esterase activities among oleaginous yeasts, a feature closely related to their ability to metabolize lipids through different pathways. Oleaginous yeasts can synthesize lipids directly from simple carbon sources, such as glucose, in a process called de novo synthesis; these lipids are stored as triacylglycerols in lipid droplets. Alternatively, these yeasts can absorb and accumulate lipids or fatty acids directly from the environment, in a process called ex novo synthesis [69]. For this latter mechanism, oleaginous yeasts release extracellular lipases/esterases that break down lipid substrates into free fatty acids that can be absorbed by cells [70]. On the other hand, for the degradation of lipids stored by cells, intracellular lipases and esterases are required [71]. The yeast Y. lipolytica, for example, can produce intracellular, extracellular, and membrane-bound lipases [71,72]. Regarding esterases, Freitas et al., through flow cytometry analysis, observed that a strain of the oleaginous yeast Rhodosporidium toruloides exhibits intracellular esterase activity until the end of yeast cultivation [73]. Moreover, Bučková et al. isolated several yeast strains from ice wine must samples, some of which (including strains of the oleaginous yeast Metschnikowia pulcherrima) demonstrated intracellular rather than extracellular esterase activity [74]. Therefore, it would be interesting to evaluate in the future whether our strains also exhibit intracellular lipase and esterase activities to further broaden the understanding of the biotechnological potential of these isolates.

Comparing well with our results, similar lipolytic activity levels were observed in Pseudozyma sp. HB27B and Sporobolomyces salmonicolor HB75 when cultured with beef tallow (66.10 ± 0.23 and 37.11 ± 1.11 U/mL, respectively) and in Papiliotrema laurentii HB18 with soybean oil (33.93 ± 3.1 U/mL) [25]. Likewise, Y. lipolytica W29 exhibited 78.0 ± 6.0 U/mL lipase activity during treatment of olive mill wastewater [75]. These variations reflect both the nature of the lipid used during cultivation and the specificity of the substrate in enzymatic assays. For instance, the extracellular lipase from Aureobasidium pullulans showed 29.8% higher activity with peanut oil than with olive oil. Enzyme activity is also influenced by pH: for A. pullulans, the optimal activity was observed at pH 7.0 [76], whereas in genetically modified Y. lipolytica strains overexpressing lipase genes, pH 6.0 yielded the highest activity [77]. Furthermore, at pH 6.0, the Brazilian yeasts isolated in Serro Minas cheese, Debaryomyces hansenii and Kodamaea ohmeri, also exhibited lipolytic activity similar (60 U/mL and 30 U/mL, respectively) to the maximum levels seen by our strains [78].

Regarding yeast lipases, the best known and most extensively studied are those derived from the yeast Candida antarctica. In a recombinant Pichia pastoris expression system, the activity of C. antarctica lipase B (CALB) reached approximately 57.9 U/mL in flask culture and up to 11,900 U/mL in fed-batch fermentation [79]. However, it is important to highlight that C. antarctica CALB lipase, although widely used, is sensitive to non-ideal environments (high temperatures, extreme pHs, or organic solvents), which reduces its activity and limits its industrial use [80]. Therefore, isolating these enzymes from novel microorganisms remains important, as new sources may provide enzymes with unique catalytic characteristics, greater stability under extreme conditions, or even different substrate specificities.

The sequencing and analysis of the D1/D2 domains of the LSU rDNA from the representative isolates revealed considerable taxonomic diversity: the thirteen identified strains were assigned to seven distinct genera—Peterozyma, Papiliotrema, Yamadazyma, Hannaella, Scheffersomyces, Meyerozyma, and Sugiyamaella. These genera encompass yeasts with broad ecological distribution and have been isolated from various natural environments. For instance, Papiliotrema species have frequently been associated with insects, plant leaves, and flowers, as well as aquatic environments [35,44,81,82], while yeasts of the genus Yamadazyma have been reported from fruits [83], mushrooms [84], food products [85], soil samples [86], and decaying wood [87]. The genus Scheffersomyces is well known for harboring several xylose-fermenting species that are commonly isolated from decaying wood [88,89] and wood-digesting insects [90]. Meyerozyma and Sugiyamaella, in turn, are broadly distributed genera, with some Sugiyamaella species reported as xylanase producers [58,91], and certain Meyerozyma strains described as capable of fermenting xylose [34,35]. Lastly, representatives of the genus Hannaella have been primarily isolated from plant surfaces [92,93], whereas yeasts of the genus Peterozyma, although less frequently reported, have been associated with flies and beetles [94,95].

Regarding our Papiliotrema isolates, the results showed that the strain CHAP-261 (P. laurentii) exhibited the highest growth rate on arabinose and the second-highest lipase activity among the analyzed strains, while strain CHAP-270 (P. terrestris) showed the highest esterase activity. In fact, strains of P. laurentii have already been shown to assimilate lignocellulosic biomass sugars such as arabinose and to produce a variety of extracellular enzymes, including lipases, proteases, amylases, and xylanases [96,97,98,99]. Some strains of P. terrestris have been used as biocontrol agents against both field and postharvest phytopathogenic fungi [100,101], with lytic enzyme production and secretion identified as one of the molecular mechanisms underlying their biocontrol effect [100].

Finally, it is worth noting that three of our sequenced isolates (Peterozyma sp. CHAP-255, Meyerozyma sp. CHAP-285, and Sugiyamaella sp. CHAP-286) did not exhibit D1/D2 domains with sufficient homology to confidently assign a specific epithet. In this regard, we consider the possibility that these isolates may represent novel species. Indeed, when yeasts are sought in natural environments, particularly in underexplored habitats, it is estimated that up to 15% of the total isolates correspond to previously undescribed species [102]. In future work, our group will focus on these yeasts, initially sequencing additional barcodes (beyond D1/D2) to confirm their status as new species. Upon such confirmation, the research will be directed toward the formal description of these novel taxa.

5. Conclusions

This study reinforces the potential of native yeast biodiversity as a key resource for the advancement of sustainable biotechnologies. The isolation and functional characterization of 30 yeast strains from a preserved Araucaria Forest revealed diverse metabolic and enzymatic traits directly aligned with the needs of second-generation biorefineries. Several strains showed high specific growth rates on sugars derived from lignocellulosic biomass, while others stood out for their capacity to secrete hydrolytic enzymes (e.g., xylanases and β-glucosidases) and accumulate intracellular lipids. The identification of strains such as Papiliotrema laurentii CHAP-261, Hannaella luteola CHAP-274, Meyerozyma carpophila CHAP-276, and Scheffersomyces coipomensis CHAP-278—with high enzymatic activity and robust sugar metabolism—underscores their potential for integration into circular and low-carbon production systems. Specifically, strains CHAP-261 and CHAP-276 were able to grow on sugars derived from lignocellulosic biomass, including glucose, xylose, cellobiose, and arabinose, demonstrating potential applicability in lignocellulosic hydrolysates. In contrast, strains CHAP-274 and CHAP-278 exhibited high xylanase and β-glucosidase activities, respectively, suggesting their relevance for the depolymerization of hemicellulose and cellulose. Notably, CHAP-274 displayed elevated xylanase activity at 50 °C, making this strain particularly promising for biomass hydrolysis processes conducted at higher temperatures. Similarly, M. carpophila CHAP-276 not only utilized all four sugars but also showed considerable xylanolytic activity, highlighting its potential for consolidated bioprocesses using hemicellulose as a substrate. The discovery of oleaginous and esterase-producing strains further broadens the applicability of these isolates in processes ranging from bioplastic precursors to enzyme-assisted waste valorization. Taxonomic analyses confirmed that these strains belong to genera with known industrial relevance, such as Scheffersomyces, Meyerozyma, Papiliotrema, and Sugiyamaella. In particular, Papiliotrema terrestris CHAP-270 was shown to be both oleaginous and highly active in esterase production. By tapping into the microbial wealth of conserved forest ecosystems, this work highlights the importance of biodiversity conservation as a pillar of sustainable innovation. These native yeasts represent promising biological tools for transforming agroindustrial residues into high-value bioproducts, thereby contributing to resource-efficient and environmentally responsible biorefining strategies.