Ethanol perturbs cell wall and membrane homeostasis in Wickerhamomyces anomalus

Abstract

Background

Wickerhamomyces anomalus, a flavor-modulating non-Saccharomyces yeast, has garnered significant interest for its remarkable ability to shape wine flavor profiles. However, during fermentation, yeast cells are inevitably exposed to ethanol stress. The specific structural consequences of this stress, particularly its impact on cell wall and membrane homeostasis, remain unclear.

Results

In this study, we investigated the effects of ethanol stress on the cellular integrity of W. anomalus through a physiological analysis focused on the cell wall and membrane. Our results demonstrated that ethanol stress induced significant morphological and ultrastructural alterations, which were primarily attributed to the disruption of cellular homeostasis. Specifically, ethanol stress compromised cell wall integrity, activated the cell wall integrity (CWI) pathway, and increased the intracellular levels of β-glucan and chitin. Furthermore, ethanol stress disrupted membrane homeostasis by remodeling its composition, reducing integrity and fluidity, while increasing permeability and simultaneously enhancing ATPase activity and elevated intracellular K⁺ levels. Fatty acid profiling also demonstrated a decrease in the monounsaturated fatty acid C18:1 and an increase in very long-chain fatty acids (VLCFAs; C22:0 and C24:0) under ethanol challenge. Exogenous supplementation of these fatty acids was shown to enhance the ethanol stress tolerance of W. anomalus.

Conclusions

These findings provide crucial insights into the mechanistic basis of ethanol stress response in W. anomalus and offer a foundation for developing novel strategies to improve its industrial utilization under fermentative stress conditions.

Supplementary Information

The online version contains supplementary material available at 10.1186/s13068-026-02756-4.

Background

Wickerhamomyces anomalus, a non-Saccharomyces yeast, has garnered significant research interest in recent years due to its exceptional flavor-modulating capabilities, largely attributed to its glycosidase activity [1, 2]. This yeast species is also notable for its tolerance to a wide range of extreme environmental conditions, surviving pH levels from 2.0 to 12.4 and temperatures from 3 °C to 37 °C, which makes it particularly suitable for food fermentation processes, especially in winemaking [3, 4]. During wine fermentation, however, yeast cells are passively exposed to increasing ethanol stress in the middle to late stages, which can inhibit cell growth and lead to stuck fermentation [5]. Therefore, enhancing ethanol tolerance is a major objective for winemakers and a key challenge for researchers. While the mechanisms of ethanol stress have been extensively studied in Saccharomyces cerevisiae as a model organism [6], they remain poorly understood in non-Saccharomyces yeasts. Given the industrial value of W. anomalus, there is a compelling need to intensify research on its ethanol stress response mechanisms.

For yeast cells, the outer structures, primarily composed of the cell wall and membrane, serve as the primary barrier against external stresses and are also the main sites of attack by stress agents [7, 8]. Therefore, maintaining the integrity and function of the cell wall and membrane is crucial for cellular homeostatic regulation and stress adaptation. Numerous studies have demonstrated that stress tolerance can be effectively enhanced through the development of resistant strains via genetic modification or through exogenous nutrient supplementation—such as amino acids, peptides, and fatty acids—by improving the structural integrity of the cell wall and membrane [9–11]. For instance, Jin et al. reported that the addition of wheat gluten peptides enhanced osmotic stress tolerance in Saccharomyces pastorianus, promoting cell growth and survival [12]. This effect was attributed to an increased proportion of palmitoleic acid (C16:1), a key unsaturated fatty acid in the membrane, which improved membrane fluidity and reduced membrane permeability. Similarly, Wang et al. found that supplementation with specific fatty acids (C12:0, C16:0, C16:1, and C18:1) improved salt tolerance in Zygosaccharomyces rouxii by modulating membrane fatty acid composition, enhancing biofilm formation, and regulating the expression of key genes involved in fatty acid metabolism [11]. In another study, Qi et al. engineered the Mediator tail subunit Med15B in Candida glabrata, which modulated cell growth under acid stress [13]. Further analysis revealed that the enhanced acid stress tolerance in Med15B-overexpressing strains was associated with increased membrane integrity, fluidity, and H⁺-ATPase activity. In line with these findings, our previous study also demonstrated that exogenous arginine supplementation improved the ethanol stress tolerance of W. anomalus [9].

The cell wall, as the outermost structure of yeast cells, is a dynamic organelle that provides mechanical strength and shape, enabling the cell to withstand environmental stresses [14]. Structurally, the yeast cell wall is composed of two main layers: an outer layer rich in mannoproteins (mannose polymers and proteins) that provides a barrier function, and an inner layer primarily of β-glucan and chitin that confers mechanical stability [7]. This structure is essential for maintaining cellular homeostasis. However, under ethanol stress, the cell wall can sustain damage. In response, yeast cells activate the cell wall integrity (CWI) pathway and initiate wall remodeling processes [6]. Evidence has shown that exogenous supplementation with protective agents can help maintain cell wall homeostasis and function, thereby enhancing yeast ethanol stress tolerance [10]. In addition to the cell wall, the cell membrane plays a critical role in safeguarding cellular integrity by serving as a selective barrier [15]. Composed of a glycerophospholipid and sphingolipid bilayer embedded with sterols, glycolipids, and membrane proteins, the membrane is also a major target of ethanol stress [16]. As a small amphipathic molecule, ethanol readily incorporates into the lipid bilayer, disrupting lipid packing and inducing a harmful increase in membrane fluidity. This incorporation compromises membrane integrity, leading to elevated permeability and leakage of essential ions and cellular components, ultimately impairing vital cellular functions [17]. This mechanistic insight is consistent with findings from related studies, in which enhancing membrane integrity and modulating fluidity were key strategies for improving ethanol stress resistance [18].

In our previous study, a flavor-modulating strain of W. anomalus, designated C11, was isolated from Rosa roxburghii Tratt—a perennial plant belonging to the Rosaceae family [4]. This strain has shown potential to enhance the flavor profile of fermented fruit wines, making it a promising candidate for developing distinctive alcoholic beverages [3]. However, ethanol, as a major and inevitable stress factor during wine fermentation, was observed to inhibit the growth and reduce the viability of W. anomalus [19]. Since the cell wall and membrane are primary targets of ethanol stress, maintaining their structural and functional homeostasis is essential for improving ethanol tolerance [10]. Nevertheless, how ethanol stress affects the homeostasis of the cell wall and membrane in W. anomalus remains unclear. Therefore, this study aims to investigate the impact of ethanol stress on the composition and molecular architecture of the cell wall and membrane, as well as the expression of key genes related to structural constituents and the CWI pathway in W. anomalus. The findings are expected to provide a theoretical basis for enhancing ethanol stress tolerance in this yeast species through reinforcement of its cell wall and membrane structures.

Materials and methods

Yeast strain and culture conditions

The W. anomalus strain C11, originally isolated from Rosa roxburghii Tratt fruit, was used in this study [4]. The cryopreserved strain was initially revived by streaking onto YEPD agar plates (containing 1% yeast extract, 2% peptone, 2% glucose, and 2% agar) and incubating statically at 28 °C for 72 h. Subsequently, a single colony was inoculated into YEPD broth (1% yeast extract, 2% peptone, and 2% glucose) and cultivated with shaking at 160 rpm for 8 h to obtain logarithmic-phase cells. For ethanol stress assessment, the experimental group was subjected to 9% (v/v) ethanol for 6 h. A control group was maintained under identical conditions in the absence of ethanol. Subsequently, cells from both groups were harvested for the analysis of various cell wall and membrane parameters, with the exception of lyticase sensitivity.

Scanning and transmission electron microscopy

Scanning electron microscopy (SEM) and transmission electron microscopy (TEM) analyses were performed according to established methods [19]. Briefly, cells were collected by centrifugation (4,000 × g, 10 min), washed three times with physiological saline (0.85% NaCl), and fixed with 2.5% glutaraldehyde at 4 °C overnight. After fixation, the cells were rinsed three times with phosphate-buffered saline (PBS; 0.1 M, pH 7.4). The samples were then dehydrated through a graded ethanol series (30%, 50%, 70%, 80%, 90%, 95%, and 100%). For SEM observation, the dehydrated samples were critical-point dried, sputter-coated with gold, and imaged using an FEI Quanta FEG 450 SEM system (FEI, Hillsboro, OR, USA). For TEM analysis, the samples were embedded in epoxy resin, ultrathin-sectioned (60–80 nm), stained with uranyl acetate and lead citrate, and examined using an FEI Tecnai Spirit TEM operated at 120 kV.

Quantification of β-1,3-glucan and chitin

The contents of β-1,3-glucan and chitin were quantified as previously described [8, 20]. Yeast cells were collected by centrifugation at 4,000 × g for 5 min, washed, and resuspended in TE buffer (10 mM Tris–HCl, 1 mM EDTA, pH 8.0). The cell suspension was adjusted to a final concentration of 1 M NaOH and incubated at 80 °C for 30 min. Thereafter, 1.05 mL of aniline blue solution (containing 0.18 M HCl, 0.03% aniline blue, and 0.49 M glycine, pH 9.5) was added to the mixture. After vortexing, samples were incubated at 50 °C for 30 min, followed by a further 30 min at 30 °C. Fluorescence intensity was measured at an excitation wavelength of 400 nm and an emission wavelength of 460 nm using a multimode microplate reader (Hitachi F-4700, Japan). The β-1,3-glucan content was calculated and expressed as a percentage relative to the control.

For chitin determination, yeast cells were harvested, washed three times with PBS, and resuspended in the same buffer. Calcofluor white stain was added to the cell suspension at a final concentration of 20 μg/mL, followed by incubation at 30 °C for 5 min in the dark. The stained cells were then washed twice with PBS and resuspended in 1 mL of PBS. Fluorescence intensity was recorded at excitation and emission wavelengths of 320 nm and 430 nm, respectively.

Yeast cell wall susceptibility to lyticase

Yeast cells were harvested by centrifugation and resuspended in TE buffer (10 mM Tris–HCl, 1 mM EDTA, pH 8.0). Lyticase was added to the cell suspensions at a final concentration of 100 μg/mL. The susceptibility of the cell wall to enzymatic digestion was assessed by measuring the decrease in optical density at 600 nm (OD₆₀₀). Readings were taken at 10 min intervals over a period of 70 min using a multimode microplate reader (Hitachi F-4700, Tokyo, Japan).

Cell wall integrity assay

The integrity of the cell wall was assessed by quantifying the release of intracellular proteins resulting from cell lysis [8]. The extent of cell lysis was evaluated based on the amount of cytosolic proteins released into the extracellular medium. After centrifugation at 4,000 × g for 10 min, the supernatant was collected, and its protein content was determined using a BCA assay kit (Beyotime, Shanghai, China), according to the manufacturer's instructions.

Assessment of cell membrane integrity, permeability, and fluidity

Cell membrane integrity was assessed by staining with fluorescein diacetate (FDA) and propidium iodide (PI) [8]. Briefly, cells were incubated with 20 μM FDA or PI at 25 °C with shaking at 150 rpm for 25 min. The stained cells were then visualized using an Olympus BX51 fluorescence microscope (Tokyo, Japan) and quantified with a Hitachi F-4700 fluorescence spectrophotometer (Tokyo, Japan). Cell membrane permeability was evaluated using a relative electrical conductivity assay, as previously described [10]. Membrane fluidity was determined by measuring the generalized polarization (GP) of the fluorescent probe Laurdan, as previously described by Jin et al. [12].

Analysis of cell membrane fatty acid composition

The fatty acid composition of cell membranes was quantified using GC–MS fatty-acid profiling. Yeast cells were harvested by centrifugation at 4,000 × g for 5 min at 4 °C. The cell pellet was mixed with a dichloromethane and methanol mixture (1:1, v/v) and homogenized in a frozen grinder at 50 Hz for 3 min. Subsequently, the sample was subjected to low-temperature ultrasonication for 15 min and allowed to stand at − 20 °C for another 15 min. After centrifugation at 10,000 × g for 10 min at 4 °C, the supernatant was collected and evaporated to dryness under a stream of nitrogen gas. A mixed internal standard was added to the residue, followed by vortexing for 30 s and incubation in a water bath at 60 °C for 30 min. The mixture was cooled, centrifuged again under the same conditions (10,000 × g, 10 min, 4 °C), and the final supernatant was transferred to a glass insert in a sample vial for GC–MS analysis.

GC–MS analysis was performed on an Agilent 7890B gas chromatography system coupled with an Agilent 5977A mass spectrometer (Agilent Technologies, Santa Clara, CA, USA). Chromatographic separation was achieved on an Agilent DB-FastFAME capillary column (20 m × 0.18 mm, 0.2 μm) using high-purity helium (purity ≥ 99.99%) as the carrier gas at a constant flow rate of 1.0 mL/min. The injector temperature was set at 230 °C, and 1 μL of sample was injected in split mode (split ratio 50:1) with a solvent delay of 1.0 min. The oven temperature was programmed as follows: initial temperature of 80 °C held for 0.5 min, increased to 175 °C at 70 °C/min, then raised to 230 °C at 8 °C/min and held for 1 min. The post-run temperature was maintained at 80 °C for 2 min. The mass spectrometer was operated with an electron impact ion source at 230 °C. The quadrupole and transfer line temperatures were set at 150 °C and 240 °C, respectively. The electron energy was 70 eV, and data were acquired in selected ion monitoring mode. Fatty acids were identified and quantified by comparison with authentic standards.

Spot assay

Cell viability was assessed via spot assay [19]. Treatments included: a control (DMSO only), an ethanol treatment [9% (v/v)], and fatty acid supplementation groups [9% (v/v) ethanol plus 70 μmol/L C18:1n9c, 60 μmol/L C22:0, or 55 μmol/L C24:0 (all dissolved in DMSO)]. After 6 h of incubation at 28 °C, cells were harvested (4,000 × g, 5 min), washed twice with distilled water, and resuspended to OD₆₀₀ = 1.0. The suspension was serially diluted (10⁰ to 10⁻⁴), spotted onto YEPD plates, and incubated at 28 °C for 36 h. Colonies were subsequently imaged using a digital microscope (Olympus, Tokyo, Japan).

Determination of intracellular K⁺ concentration and Na⁺/K⁺-ATPase activity

Intracellular K⁺ concentration was measured using a commercial assay kit (C001-2–1; Jiancheng Bioengineering, Nanjing, China). Na⁺/K⁺-ATPase activity was assayed with a separate kit (A070-2; Jiancheng Bioengineering), following the manufacturer’s protocols with reference to previous studies [11, 12]. Briefly, yeast cells were collected by centrifugation at 4,000 × g for 5 min. The pellet was washed twice with PBS (pH 7.4) and resuspended in 1 mL of PBS. The cell suspension was then sonicated on ice using an ultrasonic cell disruptor (150 W) with a cycle of 5 s on and 5 s off for a total duration of 4 min. The resulting lysate was centrifuged at 12,000 × g for 10 min at 4 °C, and the supernatant was collected for subsequent assays.

Fourier transform-infrared spectroscopy

Molecular structural alterations in the yeast cell wall and membrane were analyzed using Fourier transform-infrared (FTIR) spectroscopy, as previously described [20]. Briefly, yeast cells from both ethanol-treated and control groups were collected by centrifugation, washed with distilled water, and lyophilized. The lyophilized samples were then mixed with potassium bromide powder at a ratio of 1:100 and compressed into transparent pellets. FTIR spectra were acquired using a VERTEX 70 V Fourier Transform Infrared Spectrometer (Bruker, Bremen, Germany) across the wavenumber range of 4000 to 400 cm⁻¹ with a resolution of 4 cm⁻¹.

Real-time quantitative PCR (RT-qPCR) analysis

Total RNA was extracted from yeast cells using Trizol reagent (Invitrogen, Carlsbad, CA, USA) and further purified with a commercial kit (Takara, Kyoto, Japan). Subsequently, complementary DNA (cDNA) was synthesized from the purified RNA using the PrimeScript RT Reagent Kit (Takara) according to the manufacturer's instructions. RT-qPCR was performed on a LightCycler 96 system (Roche, Basel, Switzerland). The reaction protocol consisted of an initial denaturation at 95 °C for 30 s, followed by 40 cycles of denaturation at 95 °C for 10 s and annealing/extension at 60 °C for 30 s. A melting curve analysis was conducted immediately after amplification to verify the specificity of the products by heating from 95 °C to 65 °C over 10 s, holding at 65 °C for 60 s, and then ramping to 97 °C with continuous fluorescence acquisition. The primer sequences used are listed in Table S1. The ACT1 gene was used as an internal reference for normalization, and the relative gene expression levels were calculated using the 2^−ΔΔCT method.

Statistic analysis

All experiments were performed in three independent biological replicates, and data are presented as the mean ± standard deviation (SD). Statistical analysis was performed using SPSS 21.0 software. Differences between two groups were assessed using unpaired two-tailed Student’s t-test, while comparisons among multiple groups were analyzed by one-way analysis of variance (ANOVA) followed by Duncan’s post hoc test. A P-value of less than 0.05 was considered statistically significant. Data visualization was conducted using GraphPad Prism 10.1.2 (GraphPad Software, USA) for column graphs and TBtools v2.0 for heatmaps and hierarchical cluster analysis (HCA).

Results and discussion

Ethanol stress-induced morphological and ultrastructural alterations in W. anomalus

Electron microscope analysis could clearly reveal morphological and structural alterations in yeast cells under various environmental stress conditions [21, 22]. The morphological and structural characteristics of ethanol-treated W. anomalus cells, used to assess the effects of ethanol stress, are shown in Fig. 1. In the absence of ethanol, the cells exhibited a plump, elliptical morphology with smooth surfaces (Fig. 1A). Under ethanol stress, however, cells showed significant morphological alterations, including swelling and noticeable increase in surface roughness (Fig. 1B). Furthermore, TEM observations of the yeast ultrastructure showed that normal cells possessed intact cell walls and cytoplasmic membranes, whereas ethanol stress led to appeared thinner and coarser in cell walls, along with visible damage to the cytoplasmic membrane (Fig. 1C, D). These results indicate that ethanol stress compromises the structural integrity of the cell wall and membrane, thereby altering the morphology of W. anomalus. Similar detrimental effects have also been observed in other yeast species (e.g., S. cerevisiae and Z. rouxii) under different stress conditions (e.g., salt stress) [10, 22]. The cell wall and membrane play crucial roles in yeast stress resistance by dynamically maintaining their structural integrity [10]. The ethanol-induced impairment of these structures observed here, which likely contributes to the loss of cellular function and ultimately cell death, highlights the critical importance of structural integrity for yeast survival under stress.

Fig. 1.

SEM and TEM analysis of the morphological changes in W. anomalus under ethanol stress. Representative SEM images of cells from the control (A) and ethanol-treated (B) groups. A red arrow highlights surface irregularities, indicating swelling or increased roughness. Representative TEM images of cells from the control (C) and ethanol-treated (D) groups. A red arrow indicates regions of cell wall thinning or membrane damage. Scale bars: 5 μm (A, B); 500 nm (C, D). Abbreviations: CM: cell membrane; CW: cell wall

Ethanol stress disrupts cell wall homeostasis in W. anomalus

The cell wall is essential for maintaining yeast cell integrity, as it provides structural strength, defines cell shape, and offers protection against environmental stress [7]. Therefore, sustaining cell wall homeostasis is a key strategy for enhancing cellular stress tolerance [23]. Given the observed reduction in cell wall thickness via TEM, we further investigated the cell wall homeostasis of W. anomalus by analyzing its composition and function. The β-1,3-glucan content, which constitutes the fundamental inner structural framework of the cell wall, is quantified in Fig. 2A. Under ethanol stress, the β-1,3-glucan content increased significantly, reaching approximately twice that of the control. Chitin, a linear polymer of β-1,4-linked N-acetylglucosamine, forms the inner layer of the cell wall together with β-glucan [24]. It has been reported that chitin levels can increase by up to 20% of the cell wall dry weight in response to stress [7]. In our study, the chitin content in ethanol-stressed W. anomalus cells also increased approximately twofold, showing a significant difference from the control group (Fig. 2B). These compositional changes indicate structural remodeling of the cell wall under high ethanol stress. A key and paradoxical finding is the concurrent increase in β-1,3-glucan and chitin content alongside a decrease in overall wall thickness. This suggests that ethanol triggers not mere synthesis or degradation, but a comprehensive architectural reorganization. The increased glucan and chitin likely constitute a compensatory strengthening response, whereas the reduced thickness may stem from altered polymer assembly, cross-linking, or shifts in the proportions of other constituents (e.g., mannoproteins), resulting in a more compact architecture. Future studies quantifying the full suite of wall components are essential to fully elucidate this adaptive remodeling process.

Fig. 2.

Cell wall homeostasis analysis of W. anomalus under ethanol stress. A The β-1,3-glucan content. B The chitin content. C The sensitivity to lyticase. D The extracellular protein leakage. *P < 0.05, **P < 0.01, ***P < 0.001 compared to the control

Lyticase is an enzyme complex that primarily hydrolyzes β-1,3-glucan, a major structural component of the yeast cell wall. Therefore, its activity is routinely used to probe the integrity of the glucan layer [10, 20]. Under ethanol stress, we assessed the cell wall susceptibility to lyticase in W. anomalus. As shown in Fig. 2C, cells exposed to ethanol for 0 to 70 min exhibited significantly increased lyticase susceptibility, indicating compromised cell wall integrity. Moreovere, the impact of ethanol on CWI was evaluated by measuring the release of intracellular proteins due to cell lysis [10]. Ethanol stress significantly increased the release of intracellular proteins compared to the control, suggesting that the cell wall was damaged (Fig. 2D). Taken together, ethanol stress induced cell wall damage, which subsequently impaired its integrity, ultimately leading to cell death. It is noteworthy that although the contents of β-1,3-glucan and chitin increased in ethanol-treated cells, CWI was decreased. This may be attributed to the high concentration of ethanol and the severe damaging effects of ethanol stress.

To further elucidate ethanol-induced alterations in the cell wall composition of W. anomalus, we analyzed the expression levels of key genes involved in cell wall biosynthesis using RT-qPCR. Four genes—FKS1, KRE6, CHS3, and MNN9—were selected based on their crucial roles in this process. FKS1 encodes β-1,3-glucan synthase, catalyzing the production of β-1,3-glucan [25]. KRE6 encodes β-1,6-glucan synthase, responsible for synthesizing β-1,6-glucan [26]. CHS3 encodes chitin synthase, which participates in chitin formation [27], and MNN9 encodes a mannosyltransferase essential for mannan biosynthesis [28]. As shown in Fig. 3, the expression levels of FKS1, KRE6, CHS3, and MNN9 were significantly upregulated under ethanol stress compared to the control cells. This upregulation strongly suggests enhanced biosynthesis of glucans, chitin, and mannan within the cell wall under ethanol stress. These results are consistent with the compositional changes observed in Fig. 2A, B.

Fig. 3.

Expression levels of genes involved in cell wall synthesis of W. anomalus under ethanol stress. A Relative expression of the FSK1 gene. B Relative expression of the GSH3 gene. C Relative expression of the KRE6 gene. D Relative expression of the MNN9 gene. *P < 0.05, **P < 0.01 compared to the control

Ethanol stress regulates the expression of CWI-related genes in W. anomalus

The CWI pathway upregulates the expression of cell wall biosynthesis-related genes, thereby remodeling the cell wall structure and maintaining its stability [29, 30]. Studies have demonstrated that yeast cells activate the CWI pathway to remodel the cell wall, preserve its structural integrity, and adapt to stress conditions, ultimately enhancing cell survival [31]. To investigate the impact of ethanol stress on the expression of key genes involved in the CWI pathway and cell wall remodeling, RT-qPCR was performed. The selected genes included core components of the CWI signaling cascade: BCK1 (MAPKKK), MKK2 (MAPKK), SLT2 (MAPK), and RLM1, which encodes a transcription factor responsible for downstream transcriptional activation [6]. As shown in Fig. 4, ethanol stress significantly up-regulated the expression of BCK1, MKK2, SLT2, and RLM1 compared to the control. These results indicate that the CWI pathway is activated in response to ethanol stress, facilitating cell wall remodeling and maintaining structural integrity of W. anomalus.

Fig. 4.

Expression levels of genes involved in CWI pathway in W. anomalus under ethanol stress. A Relative expression of the BCK1 gene. B Relative expression of the MKK2 gene. C Relative expression of the SLT2 gene. D Relative expression of the RLM1 gene. *P < 0.05, ***P < 0.001 compared to the control

It is plausible that this ethanol stress response mechanism is conserved across both S. cerevisiae and non-Saccharomyces yeasts [32, 33]. This hypothesis is also supported by the observed upregulation of key CWI pathway genes (e.g., BCK1, SLT2, RLM1) in W. anomalus under ethanol stress—a transcriptional response characteristic of the well-established CWI signaling in S. cerevisiae. The consequent remodeling of cell wall composition, notably the enhanced synthesis of β-1,3-glucan and chitin, further mirrors the canonical adaptive strategy employed by S. cerevisiae to reinforce cell wall architecture.

Ethanol stress disrupts cell membrane homeostasis in W. anomalus

Cell wall variations inevitably lead to alterations in the cell membrane's function against environmental stress [34]. As a natural barrier, the cell membrane forms a protective boundary between the cell and its external environment by maintaining structural integrity and selective permeability. This thin, flexible lipid bilayer serves as a dynamic shield, regulating the passage of substances while keeping harmful agents out [35]. Its ability to maintain this protective role is crucial for cellular homeostasis, allowing the cell to thrive under diverse and often harsh conditions [36]. To evaluate the impact of ethanol treatment on the cell membrane of W. anomalus, we assessed membrane integrity, permeability, and fluidity. As shown in Fig. 5A, B, nearly all control cells were stained green, indicating intact membranes. In contrast, the number of stained cells decreased significantly under ethanol stress, suggesting impaired membrane integrity. Similar results were confirmed using the fluorochrome PI. A significantly higher proportion of cells exhibited red fluorescence after ethanol treatment, indicating membrane damage (Fig. 5C, D).

Fig. 5.

Cell membrane homeostasis analysis of W. anomalus under ethanol stress. A, B FDA staining for membrane integrity. C, D PI staining for membrane integrity. E Membrane permeability via relative electrical conductivity assay. F Membrane fluidity represented by the generalized polarization (GP) value. *P < 0.05, ***P < 0.001 compared to the control

Ethanol-induced disruption of membrane homeostasis was further demonstrated by assessing cell permeability using a well-established relative electrical conductivity assay, a method frequently employed in yeast stress response studies [10]. As shown in Fig. 5E, ethanol stress significantly increased membrane permeability, resulting in a 25.06% elevation in relative electrical conductivity compared to the control. Membrane fluidity, an important parameter for assessing membrane homeostasis, plays a vital role in adapting to environmental stress and maintaining normal physiological function. It can be evaluated using the generalized polarization (GP) value, where a higher GP corresponds to lower fluidity [10]. Under ethanol stress, the GP value increased significantly by 2.19-fold compared to the control, indicating a reduction in membrane fluidity (Fig. 5F). Previous research has suggested that reduced membrane fluidity can enhance ethanol tolerance [37]. Consistent with this concept, our observation of decreased membrane fluidity is associated with the adaptive response of W. anomalus to ethanol stress.

Fatty acids, as crucial constituents of the cellular membrane, play a fundamental role in maintaining membrane integrity, fluidity, and stress tolerance [38, 39]. Specifically, saturated fatty acids (SFAs) enhance membrane rigidity, while unsaturated fatty acids (UFAs) contribute to fluidity by introducing kinks in their hydrocarbon chains [40]. Using GC–MS fatty-acid profiling, we determined changes in the fatty acid composition of W. anomalus under ethanol stress. As shown in Fig. 6A, B a total of 22 fatty acids were identified—13 saturated fatty acids (SFAs) and 9 unsaturated fatty acids (UFAs). Under ethanol stress, four fatty acids (C13:0, C18:3n3, C22:0, and C24:0) were significantly increased, whereas 11 fatty acids (C12:0, C14:0, C14:1, C15:0, C16:0, C16:1, C17:0, C18:1n9c, C18:2n6c, C20:1, and C20:2) were significantly decreased (Fig. S1).

Fig. 6.

Membrane fatty acids composition analysis of W. anomalus under ethanol stress. A Volcano plot of fatty acids. B Cluster heatmap of fatty acid profiles. C UFAs/SFAs ratio. D C16/C18 fatty acid ratio. E Relative expression of the OLE1 gene. F Relative expression of the ACC1 gene. G Relative expression of the HFA1 gene. H Relative expression of the ELO2 gene. I Effect of exogenous C18:1, C22:0, and C24:0 supplementation on ethanol tolerance. *P < 0.05, ***P < 0.001 compared to the control

Structurally, the kinks in UFAs chains prevent tight packing between molecules, unlike the straight conformations of SFAs, thereby inhibiting the formation of a solid-like structure and helping maintain membrane fluidity [41]. Notably, the UFAs/SFAs ratio was significantly lower in the ethanol-stressed group compared to the control (Fig. 6C), suggesting a reduction in membrane fluidity. This result is consistent with data obtained from GP value analysis (Fig. 5F), confirming that ethanol stress decreases the membrane fluidity of W. anomalus. Previous studies have shown that salt stress also reduces membrane fluidity in Z. rouxii [11]. Our findings under ethanol stress similarly demonstrate increased fatty acid saturation, further supporting the observed decrease in fluidity. This shift may represent an adaptive mechanism to mitigate ethanol-induced membrane disruption and maintain cellular function in W. anomalus.

Comparative analyses have linked oxidative stress tolerance in yeast to monounsaturated fatty acids (MUFAs), particularly C16:1 and C18:1 [42]. Our study found that ethanol stress significantly reduced the levels of both C16:1 and C18:1, as well as the C16/C18 ratio, in W. anomalus (Fig. 6D, S1), further supporting their crucial function under such conditions [43, 44]. Intriguingly, although the expression of OLE1—encoding the Δ9-fatty acid desaturase that produces C16:1 and C18:1—was upregulated approximately 1.7-fold by ethanol stress (Fig. 6E), the levels of C16:1 and C18:1 decreased. This discrepancy suggests a potential inhibition of Δ9-desaturase activity under ethanol stress, a hypothesis that warrants further investigation. Additionally, our results align with prior findings that higher proportions of C12:0, C14:0, and C18:2 are associated with lower ethanol tolerance [45].

We also examined the expression of ACC1 and HFA1, which encode acetyl-CoA carboxylase—an enzyme that catalyzes the conversion of acetyl-CoA to malonyl-CoA, a key initial step in fatty acid synthesis and a substrate supplier for very long-chain fatty acid (VLCFA) production [46]. Both ACC1 and HFA1 were upregulated under ethanol stress (Fig. 6F, G). Cluster analysis indicated that ethanol stress significantly increased the abundance of VLCFAs such as C22:0 and C24:0 in W. anomalus (Fig. 6B). Thus, under ethanol stress, W. anomalus appears to upregulate ACC1 to supply substrates for VLCFA synthesis, leading to increased VLCFA incorporation into the membrane, which may help stabilize membrane structure and enhance ethanol tolerance. Consistently, the expression of ELO2, which encodes an elongase involved in VLCFA synthesis, was also significantly enhanced under ethanol stress (Fig. 6H).

Moreover, exogenous supplementation with C18:1, C22:0, or C24:0 improved yeast ethanol tolerance (Fig. 6I), further supporting the important roles of MUFA C18:1 and VLCFAs C22:0 and C24:0 in the adaptive response of W. anomalus to ethanol stress.

However, it should be noted that the ethanol-induced remodeling of fatty acid composition in W. anomalus differs from that in other yeasts. In the model yeast S. cerevisiae, ethanol stress typically increases oleic (C18:1) and palmitoleic (C16:1) acids while decreasing lauric (C12:0) and palmitic (C16:0) acids [44]. In contrast, our study on W. anomalus revealed a decrease in both C18:1 and C16:1, and an increase in C12:0. Furthermore, Issatchenkia orientalis responds differently, with an increase in stearic acid (C18:0) and a decrease in C18:1 under ethanol stress [44]. These distinct patterns suggest that fatty acid remodeling under ethanol stress exhibits significant species specificity.

Ergosterol, the principal sterol in the yeast cell membrane, plays a critical role in maintaining membrane stability and function by regulating its thickness, integrity, and permeability [47, 48]. The biosynthesis of ergosterol relies on key enzymes such as squalene epoxidase (encoded by ERG1) and C-8 sterol isomerase (encoded by ERG2), both of which are essential for yeast survival under stress conditions [49]. To evaluate the effect of ethanol stress on ergosterol biosynthesis, we measured the transcript levels of ERG1 and ERG2 (Fig. 7). The results demonstrated that ethanol stress significantly upregulated the expression of both ERG1 and ERG2 genes, with transcript levels increasing to 1.84-fold and 3.92-fold of the control group, respectively (Fig. 7A, B). This finding is consistent with previous reports by Swan et al., which revealed that higher cellular sterol levels confer greater tolerance to environmental stress, and that ergosterol concentrations directly correlate with survival rates [50]. Similarly, Yu et al. observed that a 1.6-fold increase in ergosterol concentration enhanced the stress tolerance of S. cerevisiae compared to its parent strain [51]. Therefore, it can be inferred that W. anomalus enhances ergosterol biosynthesis to maintain cell membrane stability and mitigate ethanol-induced damage.

Fig. 7.

Ethanol stress-induced expression of ergosterol biosynthesis genes in W. anomalus. A Relative expression of the ERG1 gene. B Relative expression of the ERG2 gene. *P < 0.05, **P < 0.01 compared to the control

Ethanol stress induces intracellular K⁺ accumulation and enhances membrane ATPase activities in W. anomalus

Maintaining high intracellular K⁺ levels is crucial for yeast viability, as it influences cell volume, plasma membrane potential, and metabolic processes [52]. Moreover, K⁺ has been shown to play a vital role in the response to various environmental stresses in both plants and yeast [53, 54]. Previous research demonstrated that K⁺ supplementation enhanced the tolerance of S. pastorianus to osmotic and ethanol stress by restoring cell membrane and mitochondrial function [55]. In this study, intracellular K⁺ levels and cell membrane ATPase activity were measured in W. anomalus under ethanol stress (Fig. 8). The intracellular K⁺ content increased significantly in ethanol-stressed cells, reaching approximately 30 times that of the control (Fig. 8A). Previous studies have suggested that maintaining high intracellular K⁺ concentrations can improve ethanol tolerance in S. cerevisiae [56, 57]. Therefore, further experiments involving exogenous K⁺ supplementation under ethanol stress are warranted to confirm the role of intracellular K⁺ in the ethanol tolerance of W. anomalus.

Fig. 8.

Intracellular K⁺ (A) and membrane ATPase activity (B) analysis of W. anomalus under ethanol stress. **P < 0.01, ***P < 0.001 compared to the control

Na⁺/K⁺-ATPase is a transmembrane protein essential for cellular ion homeostasis, regulating the Na⁺ and K⁺ gradients across the membrane [43, 54, 58]. As shown in Fig. 8B, Na⁺/K⁺-ATPase activity in the ethanol-stressed group was significantly enhanced, reaching 4.28 times that of the control. This result is consistent with previous reports indicating that yeasts modulate Na⁺/K⁺-ATPase activity and maintain transmembrane K⁺ gradients as a common strategy to cope with environmental stress [54, 58]. Additionally, several studies have shown that exogenous nutrient supplementation, such as nitrogen sources, can effectively enhance Na⁺/K⁺-ATPase activity and consequently improve cellular ethanol tolerance [12].

Ethanol stress induces structural changes in W. anomalus as assessed by spectral assay

FTIR spectroscopic analysis is an effective technique for monitoring changes in the structural components of the cell wall and cell membrane, enabling rapid assessment of the impact of stress factors on cellular architecture [59, 60]. A comparison of the FTIR absorption peaks between the control group and the ethanol-stressed group of W. anomalus revealed largely consistent peak positions, indicating similar chemical compositions. However, the ethanol-stressed group exhibited a significant increase in the absorption intensity of several characteristic peaks, reflecting notable alterations in cellular components upon ethanol exposure.

As shown in Fig. 9, the FTIR spectrum of W. anomalus is characterized by several key absorption bands: the O–H stretching vibration at 3278.6 cm⁻¹, characteristic protein peaks at 1635.4 cm⁻¹ (amide I), 1540.9 cm⁻¹ (amide II), and 1238.1 cm⁻¹ (amide III), as well as methyl and methylene group vibrations at 2921.8 cm⁻¹ and 1400.1 cm⁻¹. The peak at 1043 cm⁻¹ is tentatively assigned to C–O stretching of carbohydrates in the cell wall, while the peak at 516.9 cm⁻¹ may correspond to C–O–C stretching vibrations in nucleic acids and phospholipids. Following ethanol stress treatment, the O–H stretching vibration peak at 3278.6 cm⁻¹ showed no noticeable shift but exhibited a 43.4% increase in absorption intensity compared to the control. Similarly, the CH₃ and CH₂ absorption peaks did not shift significantly, though their intensities rose by 45.5% and 29.3%, respectively. In contrast, the C = O stretching vibration of amide I shifted from 1635.4 cm⁻¹ to 1633.5 cm⁻¹, with a 71.6% increase in intensity. The amide II peak, associated with N–H bending and C–N stretching, shifted from 1540.9 cm⁻¹ to 1535.1 cm⁻¹ and showed a 61.8% intensity increase. The amide III band, also related to N–H bending and C–N stretching, shifted from 1238.1 cm⁻¹ to 1236.2 cm⁻¹, with a 37.03% rise in intensity. The shifts and intensity changes in the amide I and II bands suggest alterations in protein conformation, which may affect membrane integrity and function, potentially due to ethanol disrupting hydrogen bonding within proteins.

Fig. 9.

FT-IR spectral features analysis of W. anomalus under ethanol stress

Similarly, under ethanol stress, the C–O stretching vibration of carbohydrates shifted from 1043 cm⁻¹ to 1041.4 cm⁻¹, with intensity rising to 88.3% of the control level. The C–O–C stretching vibration of nucleic acids and phospholipids shifted from 516.9 cm⁻¹ to 511.1 cm⁻¹, with intensity increasing to 116.6% of the control. These spectral changes provide molecular-level insights into the structural perturbations of the cell wall and membrane in W. anomalus under ethanol stress.

Based on the aforementioned results, a schematic diagram summarizing the homeostatic regulation of the cell wall and cell membrane in W. anomalus under ethanol stress is presented in Fig. 10. Ethanol stress activates the CWI pathway, regulates the expression of genes involved in cell wall synthesis (e.g., FKS1, GHS3, KRE6, and MNN9), and thereby remodels the cell wall structure. On the other hand, ethanol stress also targets the cell membrane, upregulating the expression of ERG1 and ERG2, which enhances ergosterol synthesis. Additionally, it alters the fatty acid composition by reducing the content of monounsaturated fatty acid C18:1 and increasing very-long-chain fatty acids (VLCFAs) such as C22:0 and C24:0. Concurrently, it induces intracellular K⁺ accumulation and enhances membrane ATPase activity. These collective damages result in morphological and ultrastructural alterations, which likely further inhibit cellular growth and reduce viability.

Fig.10.

Schematic diagram of the homeostatic regulation of cell wall and cell membrane in W. anomalus in response to ethanol stress. Components highlighted in red indicate up-regulation or increase, whereas those in blue represent down-regulation or decrease. SFAs: saturated fatty acid; UFAs: unsaturated fatty acid; VLCFAs: very long-chain fatty acid

Conclusions

The present study elucidates the regulatory mechanisms of W. anomalus in response to ethanol stress from the perspective of the cell wall and cell membrane. Ethanol stress induced morphological and ultrastructural alterations, which were attributed to the disruption of cell wall and membrane homeostasis. Although the contents of β-glucan and chitin increased and the expression of related synthesis genes was up-regulated, ethanol stress ultimately impaired the integrity of the cell wall. In addition, ethanol stress disturbed cell membrane homeostasis by modifying its composition, reducing integrity and fluidity, while increasing permeability and enhancing ATPase activity. The findings of this study may contribute to the development of ethanol-tolerant industrial strains of W. anomalus.

Author contributions

X.L., Y.L., and X.Z. designed the research. Y.W., Z. K., and X. L. conducted the experiments. M.W., X.Z., Z.K., X.L., Y.L., and X. G. analyzed the data. X.L. and Y.L. wrote and revised the original draft. All authors contributed to reviewing the manuscript and approved the final version.

Funding

This work was supported by the National Natural Science Foundation of China (32160557) and Guizhou Provincial Science and Technology Foundation (Qiankehejichu MS [2026] 244; [2025]192).

Data availability

All data generated or analyzed during this study are included in this published article and its supplementary information files.

Declarations

Ethics approval and consent to participate

Not applicable.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.