Integrative Analysis of Metabolic Signature and Phytohormone Response in Potato Under Heat, Drought and Salt Stresses

Abstract

Climate change-driven abiotic stresses threaten global potato production, yet stress-specific adaptive mechanisms remain poorly defined. We demonstrate that heat, drought and salt stresses induce fundamentally distinct physiological and biochemical responses in potato plants. Photosynthetic performance and gas exchange showed stress-specific patterns, with heat stress (HS) maintaining elevated carbon metabolism, drought stress (DS) causing severe photosynthetic suppression and water deficit, while salt stress (SS) exhibited intermediate physiological impairment. Secondary metabolite (SM) profiling revealed a corresponding stress-specific signature, where sesquiterpenes (caryophyllene, copaene, humulene) were dramatically elevated under HS but suppressed under DS (which specifically enhanced 1-hexanol and trans-sesquisabinene hydrate), while SS induced copaene and cis-β-farnesene but reduced caryophyllene. Phytohormone analysis demonstrated differential accumulation patterns across stresses: JA, JA-Ile, SA and ABA were maximally elevated under HS, moderately increased under DS, while SS uniquely maintained basal JA/JA-Ile with enhanced SA and ABA. Pharmacological intervention using hormone biosynthesis inhibitors (DIECA, SHAM, Jarin-1, AIP, ABT, fluridone) and exogenous ABA confirmed stress-specific regulatory networks. These findings establish a stress-specific hormone–metabolite regulatory framework, providing a molecular basis for developing climate-resilient potato genotypes.

1. Introduction

Climate change and environmental deterioration have intensified the frequency and severity of abiotic stresses, posing a significant threat to global food security and agricultural productivity [1,2,3]. Among these, heat, drought or salinity represent the most prevalent environmental constraints that limit crop growth and development. These abiotic stresses disrupt fundamental physiological processes in crops, including photosynthesis, water relations, nutrient uptake and cellular metabolism, ultimately leading to substantial yield loss [1,4,5]. Potato (Solanum tuberosum L.), the most important food crop, is particularly vulnerable to abiotic stress conditions due to its shallow root systems, high water requirement and sensitivity to temperature extremes [6]. Understanding how potato crops perceive, respond to and adapt to multiple abiotic stresses is important for developing climate-resilient genotypes and implementing effective stress management strategies to ensure sustainable potato production in the face of the changing climate.

Crops have evolved sophisticated defense mechanisms to deal with abiotic stress conditions, among which the release of secondary metabolites (SMs) represents a dynamic and multifaceted stress response strategy [7,8,9]. SMs are low molecular weight organic-compounds with high vapor pressure that can easily evaporate at ambient temperature and diffuse through air or water. Under abiotic stress conditions, crops significantly alter their SMs release both qualitatively and quantitatively as part of their adaptive response mechanism [10,11,12]. These stress-induced SMs serve multiple protective functions: they act as signaling molecules that prime defense responses in neighboring crops, function as reactive oxygen species (ROS) scavengers that mitigate oxidative damage, help stabilize cellular membranes under stress conditions and participate in thermo-tolerance mechanisms by dissipating excess heat energy [13,14,15]. Furthermore, stress-induced SMs can change the microclimate around crop tissues through evaporative cooling and provide protection against secondary stress that often accompanies primary abiotic stress conditions. The dynamic nature of SMs release under different stress types suggests that these compounds could serve as a sensitive, non-invasive indicator of plant stress status, potentially enabling early detection of stress conditions before visible symptoms appear and irreversible damage occurs [16,17,18].

The regulation of SMs biosynthesis and release under abiotic stress is intricately linked to the phytohormone signaling network, which functions as the central coordinator of crop stress responses [19]. Phytohormones such as abscisic acid (ABA), jasmonic acid (JA) and salicylic acid (SA) play a pivotal role in mediating crop adaptation to abiotic-stresses by modulating physiological processes and coordinating whole-plant response [20,21,22,23]. ABA, often termed the “stress hormone”, accumulates rapidly under drought and salinity stress and regulates stomatal closure, root hydraulic conductivity and the expression of stress-responsive genes [20,21]. JA and its derivatives mediate responses to various abiotic stresses by activating defense gene expression and modulating secondary metabolisms [22]. SA contributes to thermo-tolerance and oxidative stress management through its antioxidant properties and signaling function [23]. These phytohormones do not function individually but rather interact through complex crosstalk networks that fine-tune crops’ stress response [24,25]. However, the specific regulatory role of individual phytohormones in controlling SM release patterns under different abiotic stress types remains poorly understood.

To elucidate the regulatory mechanisms governing stress-induced SMs release, pharmacological intervention using specific phytohormone biosynthesis inhibitors offers valuable insights into hormone-SMs interactions [26]. Diethyldithiocarbamate (DIECA), Salicylhydroxamic acid (SHAM) and Jarin-1 inhibit JA biosynthesis, which target key enzymes in the octadecanoid pathway, allowing investigation of jasmonate-dependent VOC regulation [27,28,29]. ABA biosynthesis inhibitors such as fluridone block carotenoid biosynthesis and consequently prevent ABA accumulation, enabling examination of ABA-mediated stress responses [30]. SA biosynthesis inhibitors, such as 2-aminoindane-2-phosphonic acid (AIP) and 1-aminobenzotriazole (ABT), can inhibit the phenylalanine ammonia-lyase (PAL) activity as well as benzoic acid 2-hydroxylase (BA2H), which hydroxylate the SA biosynthesis [31]. Application of these inhibitors prior to or during stress exposure and subsequently monitoring changes in SMs release and phytohormone levels, it becomes possible to establish which hormonal pathways are necessary for specific SM responses and whether different abiotic stresses utilize distinct or overlapping hormonal regulatory mechanisms to control SMs release [32,33].

Despite growing recognition of the importance of SMs in plant stress responses, several critical knowledge gaps remain unaddressed, particularly regarding crop species of agricultural importance. First, while individual abiotic stresses have been studied in model plants [24,34,35], comparative analysis of how different stress types (heat, drought and salinity) affect SMs release in the same crop species is limited, making it difficult to identify stress-specific versus general stress response patterns. Second, the regulatory roles of specific phytohormones in controlling stress-induced SMs release remain largely unclear, especially regarding whether different stresses utilize distinct hormonal pathways to regulate SMs production [36,37]. Also, the relationship between SMs release patterns and conventional physiological stress indicators has not been systematically evaluated, leaving it uncertain whether SMs profiling can provide early, sensitive and specific detection of different stress types [38,39].

The objectives of the present study are to address knowledge gaps in understanding plant differential SM-regulated responses to abiotic stress types by conducting a comprehensive investigation of the physiological responses, SM release patterns and hormonal regulation of stress responses in potato plants exposed to three major abiotic stresses: heat, drought and salinity. Potato “scents” can provide a biochemical fingerprint of stress physiology. Current research holds significant implications for both fundamental plant stress biology and practical agricultural applications. Identification of stress-specific SM biomarkers could enable the development of non-invasive, real-time sensing technologies for precision agriculture applications, allowing farmers to detect stress conditions before visible symptoms appear and yield losses become irreversible.

2. Results

2.1. Abiotic Stresses Induce Distinct Physiological, Morphological and Biochemical Responses in Potato Plants

To assess the physiological responses of potato plants to different abiotic stresses, we first examined key photosynthetic parameters under heat stress (HS), drought stress (DS) and salt stress (SS) conditions. Gas exchange measurement revealed a distinct stress-specific pattern in photosynthetic efficiencies (Figure 1A–D). Transpiration rate (E) was significantly high under HS conditions compared to control plants (F_5,144 = 46.17, p = 0.0001), while both DS and SS treatments resulted in a marked reduction in E (Figure 1A). Similarly, stomatal conductance (gs) showed the highest values in HS-treated plants, while DS induced the most severe suppression of gs, with SS showing an intermediate effect (F_5,144 = 36.72, p = 0.0001) (Figure 1B). Internal CO₂ concentrations (Ci) displayed contrasting patterns, with HS-treated plants maintaining significantly higher Ci levels comparable to controls (F_5,144 = 149.7, p = 0.0001), while both DS and SS treatment caused a dramatic reduction in Ci (Figure 1C). This was accompanied by a corresponding change in net photosynthetic rate (A), where HS treatment sustained relatively high photosynthetic activity, whereas DS and SS treatments severely impaired carbon assimilation capacity (F_5,144 = 43.26, p = 0.0001) (Figure 1D). Notably, DS treatment induced the most severe photosynthetic inhibition among all stress conditions tested. SEM analysis of leaf surface ultrastructure provided visual evidence of stress-induced stomatal and epidermal modification. Under HS, leaf stomatal openings were enhanced, whereas DS and SS decreased stomata openings (Figure 1E–J).

Figure 1.

HS, DS and SS stress differentially affect photosynthetic gas exchange parameters and leaf surface ultrastructure in potato plants. (A) Transpiration rate (E), (B) stomatal conductance (gs), (C) internal CO₂ concentration (Ci) and (D) net photosynthetic rate (A) measured in potato leaves under control and stress conditions (n = 25 plants/treatment). Box plots show median, quartiles and individual data points. Different letters indicate significant differences among treatments (p < 0.05, ANOVA followed by Tukey’s HSD test). (E–J) Representative SEM images showing leaf surface ultrastructure (200 µm). (E) Control leaf surface under heat stress conditions, (F) heat-stressed leaf showing stomatal modifications, (G) control leaf surface under drought stress conditions, (H) drought-stressed leaf with closed stomata, (I) control leaf surface under salt stress conditions, (J) salt-stressed leaf.

Given the profound effect on gas exchange, we next evaluated plant water status and cellular membrane stability under stress conditions (Figure S1). Leaf water potential measurement revealed that all three stress treatments induced significant water deficit, with DS causing the most severe decline in water potential (F_5,24 = 6.98, p = 0.0003), followed by HS and SS (Figure S1A). The difference between stress treatments and their respective controls was highly significant, indicating substantial disruption of plant water homeostasis. Relative water contents analysis corroborated these findings, showing that DS treatment reduced leaf water contents to approximately 45% compared to the control level, representing the most dramatic dehydration response among all treatments (F_5,24 = 8.218, p = 0.0001) (Figure S1B). Salt stress also significantly decreased RWC, albeit to a lesser extent than DS, while HS showed a moderate effect on tissue hydration status. Interestingly, water use efficiency remained relatively stable across all treatments (Figure S1C), suggesting compensatory physiological adjustment to maintain water–carbon balance under stress conditions (F_5,24 = 0.488, p = 0.782).

To assess cellular membrane integrity under stress conditions, we measured cell membrane stability and chlorophyll content as indicators of stress-induced damage (Figure S2). Cell membrane stability was significantly compromised under HS and DS treatment, with DS showing the most pronounced membrane destabilization (F_5,18 = 19.77, p = 0.0001) (Figure S2A). Salt stress also induced moderate membrane damage, indicating oxidative stress-mediated lipid peroxidation across all stress treatments. Chlorophyll contents analysis revealed treatment-specific pigment dynamics (Figure S2B). Heat-stress-treated plants exhibited enhanced chlorophyll levels compared to the control, potentially reflecting a compensatory response to maintain photosynthetic capacity. In contrast, DS treatment resulted in the most significant reductions in chlorophyll content (F_5,18 = 4.677, p = 0.0065), consistent with the severe photosynthetic impairment observed in gas exchange measurement (Figure 1D). Salt stress induced moderate chlorophyll degradation, further supporting the oxidative damage hypothesis. To further characterize stress-induced oxidative damage, we quantified proline accumulation and MDA contents as key biochemical markers (Figure S3). Proline contents, a well-established osmo-protectant and stress indicator, showed dramatic accumulation under DS treatments (F_5,30 = 13.08, p = 0.0001) (Figure S3A). Both HS and SS treatments also induced significant proline accumulations, though to a lesser extent than DS, reflecting differential osmotic adjustment strategies across stress types. Malondialdehyde contents, an indicator of lipid peroxidation and oxidative membrane damage, were significantly increased under DS conditions, reaching the highest levels among all treatments (F_5,24 = 5.447, p = 0.0017) (Figure S3B). Heat stress also induced substantial MDA accumulation, consistent with the membrane stability data (Figure S2A). Salt stress showed moderate increases in MDA contents, suggesting intermediate levels of oxidative damage. Collectively, these biochemical markers demonstrate that drought stress imposed the most severe oxidative burden on potato plants, followed by heat and salt stresses.

2.2. Stress-Specific Secondary Metabolite Signatures and Phytohormone Dynamics Define Distinct Stress Response Pathways in Potato

Having established distinct physiological responses to different abiotic stresses, we next investigated whether these stresses induced characteristic changes in secondary metabolite profiles that could serve as specific stress biomarkers. Comprehensive metabolomic analysis identified 23 major volatile and semi-volatile metabolites across all treatments, revealing striking stress-specific accumulation patterns (Figure 2A). Hierarchical clustering analysis clearly separated the metabolite profile by stress type, indicating profound metabolic reprogramming under stress conditions. Heat stress induced a unique metabolite signature characterized by dramatic up-regulation of a specific volatile compound. Notably, caryophyllene, humulene and copaene showed pronounced accumulation under HS conditions compared to control. In contrast, most other metabolites, including cis-β-farnesene, ylangene, cedrol and various aldehydes (hexanal, octanal, heptanal, 1-hexanol), also showed variable changes in accumulation across stress type (Figure 2A). In contrast, DS led to more generalized metabolite suppressions, with the majority of detected compounds showing reduced accumulations. Salt-stress exhibited an intermediate profile, with selective up-regulation of certain metabolites, including cis-β-farnesene and moderate changes in sesquiterpene level. Principal component analysis (PCA) of the metabolites data further confirmed the stress-specific metabolic signature (Figure 2B). The first two principal components accounted for 67.3% of the total variance (PC1: 38.7%, PC2: 28.6%), effectively discriminating among control and stress-treated samples. Heat-stress samples separated distinctly along PC1 and PC2, driven primarily by elevated levels of humulene, copaene, camphor and caryophyllene. Drought stress samples occupied a unique position in the PCA space, associated with increased 3-carene and 1-hexanol. Salt stress samples showed intermediate positioning, with cis-β-farnesene and heptanal as key contributing metabolites. The magnitude of PCA separation, quantified as the Euclidean distance from the control sample, was greatest for DS treatment, followed by SS and HS (Figure 2C), indicating that DS induced the most extensive metabolic reprogramming.

Figure 2.

Secondary metabolite profiling reveals stress-specific metabolic signature in potato leaves. (A) Heatmap showing hierarchical-clustering of volatile and semi-volatile metabolites across control and stress-treated samples. Colors represent log₂-transformed metabolite level. (B) Principal component analysis (PCA) biplot illustrating the separation of samples based on metabolite profile. Colored circles represent individual abiotic stress types: black (control), red (HS), blue (DS), green (SS). Loading vectors (arrows) indicate metabolite contribution to sample separation. (C) Magnitude of metabolic reprogramming quantified as the Euclidean distance in PCA space (PC1 + PC2) from the respective control sample. Bars represent mean distances, demonstrating the extent of stress-induced metabolic changes across treatment (n = 7 plants/treatment). * indicate log₂-transformed values.

Next, we quantified the specific metabolites that showed the most pronounced stress-responsive pattern (Figure 3). Hexanal level remained relatively stable across most treatments, with both HS and DS causing a significant reduction compared to their respective control (F_5,36 = 32.47, p = 0.0001) (Figure 3A). In contrast, humulene accumulation was dramatically and specifically elevated under HS conditions, while both DS and SS treatments failed to induce significant changes (F_5,36 = 59.35, p = 0.0001) (Figure 3B). This stress-specific response pattern establishes humulene as a potential biomarker for HS in potatoes. Copaene exhibited a similar heat-specific induction pattern, with HS treatment resulting in approximately a 2.3-fold increase compared to control, whereas DS and SS treatments caused a significant reduction in copaene level (F_5,36 = 24.14, p = 0.0001) (Figure 3C). Heptanal showed a unique response profile, with dramatic accumulation specifically under SS conditions, while HS and DS treatments maintained relatively stable levels (F_5,36 = 87.78, p = 0.0001) (Figure 3D), suggesting heptanal as a potential SS biomarker. Caryophyllene accumulation paralleled the humulene pattern, showing specific and substantial elevation under HS treatment, while DS and SS caused marked suppression of caryophyllene biosynthesis (F_5,36 = 40.21, p = 0.0001) (Figure 3E). Similarly, camphor levels were significantly elevated under HS conditions, while DS and SS treatments resulted in a dramatic reduction (F_5,36 = 71.23, p = 0.0001) (Figure 3F). Collectively, these data demonstrate that HS uniquely activates sesquiterpene and monoterpene biosynthesis pathways, while drought and salt stresses generally suppress secondary metabolite production.

Figure 3.

Quantification of stress-responsive volatile metabolites in potato leaves. (A) hexanal, (B) humulene, (C) copaene, (D) heptanal, (E) caryophyllene and (F) camphor content measured in control and stress-treated potato plants (n = 7 plants/treatment). Bars represent mean ± standard error; individual data point is overlaid. Different letters indicate significant differences among treatments (p < 0.05, ANOVA followed by Tukey’s HSD test).

Given the distinct metabolic signatures associated with different stress types, we hypothesized that these responses might be mediated by differential phytohormone signaling. Comprehensive phytohormone profiling revealed stress-specific hormone accumulation patterns that strongly correlated with the observed physiological and metabolic phenotypes (Figure 4). JA, a key regulator of stress response and secondary metabolism, showed the most dramatic accumulation under HS treatment (F_5,36 = 35.57, p = 0.0001) (Figure 4A). Drought stress also induced significant but moderate JA accumulation, while SS treatment showed no significant change. The bioactive JA conjugate, JA-Ile, displayed a nearly identical response pattern, with HS inducing the highest accumulation, while SS showed minimal effects (F_5,36 = 25.49, p = 0.0001) (Figure 4B). This strong correlation between JA/JA-Ile levels and sesquiterpene accumulation under HS suggests that JA signaling drives heat-specific secondary metabolite biosynthesis. SA, another critical defense hormone, exhibited stress-specific accumulation, with HS inducing the most pronounced increase, followed by SS, while DS showed moderate elevation (F_5,36 = 83.54, p = 0.0001) (Figure 4C). ABA, the primary stress hormone regulating water balance and stomatal function, displayed a distinct accumulation pattern that aligned with the severity of water deficit. Heat stress and DS induced dramatic ABA accumulation, whereas SS showed moderate ABA elevation (F_5,36 = 131.6, p = 0.0001) (Figure 4D). These ABA dynamics directly correlate with the stomatal closure patterns observed in gas exchange measurement (Figure 1B) and the reduced RWC under DS and HS (Figure S1B). Interestingly, IAS, the major auxin, showed significant suppression under all stress conditions, with HS, DS and SS treatments causing 73%, 54% and 47% reductions in IAA levels, respectively, compared to their controls (F_5,36 = 50.8, p = 0.0001) (Figure 4E). This generalized IAA suppression likely contributes to the stress-induced growth inhibition observed across all treatments. CK level exhibited the most dramatic stress-induced reduction, with HS, DS and SS causing 51%, 63% and 80% decreases, respectively (F_5,36 = 72.98, p = 0.0001) (Figure 4F). The severe CK depletion, particularly under SS conditions, correlates with reduced chlorophyll contents (Figure S2B) and impaired photosynthetic performances (Figure 1D), consistent with the essential role of CK in maintaining chloroplast function and delaying senescence.

Figure 4.

Phytohormone profiling reveal stress-specific hormonal dynamics in potato plant. (A) JA (B) JA-Ile (C) SA (D) ABA (E) IAA and (F) CK content in potato leaves under control and stress conditions (n = 7 plants/treatment. Bars represent mean ± standard error; individual data point is shown. Different letters denote significant differences among treatments (p < 0.05, ANOVAs followed by Tukey’s HSD test).

2.3. Pharmacological Dissections of Hormone Metabolite Network Reveals Stress-Specific Regulatory Mechanism

Comprehensive metabolomics profiling of inhibitor-treated plants revealed striking stress-specific and inhibitor-specific metabolic reprogramming (Figure 5). Under HS conditions, the metabolite landscape was profoundly altered by various hormone pathway manipulations. The most dramatic metabolic shifts under HS were observed with DIECA and Jarin-1 treatment, which strongly suppressed the accumulation of sesquiterpenes—particularly copaene, caryophyllene and humulene—that were otherwise highly induced by HS alone (Figure 5A). This suppression suggests that both ethylene and JA pathways are critical for heat-induced sesquiterpene biosynthesis. In contrast, SHAM treatment showed a more nuanced effect, moderately reducing the sesquiterpene level while maintaining relatively higher levels of certain volatile aldehydes. ABT treatment, which inhibits the activity of benzoic acid 2-hydroxylase (BA2H) involved in multiple hormone biosynthesis pathways, caused widespread metabolite suppression, indicating the central role of BA2H in stress-induced secondary metabolism. Interestingly, fluridone treatments under HS resulted in partial maintenance of sesquiterpene level, suggesting that while ABA accumulates substantially under HS (Figure 3D), it plays a less direct role in regulating sesquiterpene biosynthesis compared to the JA and ethylene pathways. Exogenous ABA applications under HS induced concentration-dependent effects, with lower doses (50 μM) causing moderate metabolic changes, while higher doses (100 μM) resulted in more pronounced alterations in metabolite profiles, particularly affecting volatile aldehyde and monoterpenes levels (Figure 5A).

Figure 5.

Comprehensive metabolomic profiling reveals stress-specific and hormone pathway-dependent metabolite reprogramming in potato. Heatmap showing hierarchical clustering of major volatiles and semi-volatile metabolites across control, stress and hormone inhibitor-treated samples under three abiotic-stress conditions (n = 7 plants/treatment. Colors represent log₂-transformed metabolite level. (A) Left panel (heat stress), (B) middle panel (drought stress) and (C) right panel (salt stress). Metabolites are ordered by hierarchical clustering based on Euclidean distances and the complete linkage method. * indicate log₂-transformed values.

Under DS conditions, the metabolite response to inhibitor treatments showed a distinctly different pattern compared to HS. Fluridone treatment, which blocks ABA biosynthesis, caused the most dramatic metabolic alterations under drought, with widespread suppression of multiple metabolite classes. This contrasts sharply with its effects under HS, highlighting the central and dominant role of ABA in orchestrating drought-specific metabolic response. Exogenous ABA applications under DS induced dose-dependent metabolite accumulation, with 100 μM ABA treatment substantially elevating the level of several metabolites that were otherwise suppressed by drought alone, suggesting that optimal ABA levels are crucial for maintaining metabolic homeostasis under water-deficit conditions (Figure 5B). Salt stress metabolite profiles showed intermediate sensitivity to inhibitor treatment. Fluridone treatment resulted in moderate metabolic suppression, while high-dose ABA applications (100 μM) induced substantial metabolite accumulations, particularly for sesquiterpene and certain aldehydes. This pattern suggests that salt stress-induced metabolic responses are regulated by a more balanced interplay between multiple hormone pathways, with no single pathway exerting dominant control (Figure 5C).

2.3.1. Pharmacological Manipulation of Hormone Pathways Reveals Differential Regulation of Sesquiterpene Biosynthesis Across Stress Conditions

We conducted detailed quantitative analysis of copaene, caryophyllene and humulene—the three sesquiterpenes most dramatically induced by abiotic stresses (Figure 6). Under HS, copaene level increased and this heat-induced copaene accumulation was dramatically suppressed by DIECA treatment, demonstrating that JA biosynthesis is essential for heat-stimulated copaene production (F_9,60 = 20.33, p = 0.0001). Similarly, SHAM treatment reduced copaene, while Jarin-1 treatment caused an even more pronounced suppression, indicating that JA signaling is the primary driver of copaene biosynthesis under heat stress. Remarkably, ABT and AIP treatments resulted in an unexpected increase in copaene levels, which, while lower than that of untreated HS, remained significantly elevated compared to control, suggesting complex regulatory interactions. Exogenous ABA applications showed dose-dependent response, with 100 μM ABA treatment resulting in significantly lower levels, suggesting that excessive ABA may antagonize JA-mediated sesquiterpene biosynthesis (Figure 6A). In contrast to HS, DS-induced metabolite responses were predominantly controlled by ABA signaling. Copaene level under DS decreased, representing 32% reduction. DS treatment alone reduced copaene, while fluridone treatment under drought partially restored copaene level, significantly lower than DS alone, indicating that basal ABA is necessary for maintaining even the reduced copaene level observed under droughts (F_4,30 = 28.29, p = 0.0001). Remarkably, exogenous ABA application significantly reduced copaene accumulations, demonstrating ABA’s capacity to regulate induction when present at an appropriate concentration. Under SS conditions, copaene levels were relatively stable across treatment (F_4,30 = 26.64, p = 0.0001). The modest effect of fluridone and the moderate reduction by exogenous ABA suggest that multiple hormone pathways contribute to copaene regulation under SS, with no single pathway being dominant (Figure 6A).

Figure 6.

Hormone pathway inhibitors and exogenous ABA differentially regulate key sesquiterpene accumulation across abiotic stress types. (A) Copaene (B) caryophyllene and (C) humulene contents in potato leaves under control, stress and hormone inhibitor/ABA-treated conditions (n = 7 plants/treatment. Left panels (heat stress): Yellow bars represent various treatments under heat stress conditions. Middle panels (drought stress): Blue bars show drought stress treatments. Right panels (salt stress): Green bars display salt stress treatments. Bars represent means ± standard error; individual data points are overlaid as open circles. Different letters indicate significant differences among treatments within each stress type (p < 0.05, ANOVAs followed by Tukey’s HSD test).

Caryophyllene exhibited even more dramatic responses to hormone pathway manipulation under HS (Figure 6B). DIECA treatment potently suppressed caryophyllene induction, as did SHAM and Jarin-1 treatment, collectively demonstrating that the JA pathway synergistically regulates caryophyllene biosynthesis. Interestingly, ABT and AIP treatments resulted in the intermediate caryophyllene level, suggesting that the SA and JA pathways work synergistically to induce metabolic induction. Fluridone showed surprisingly robust caryophyllene accumulation, supporting that ABA signaling is not directly required for heat-induced caryophyllene induction. Low-dose ABA (50 μM) reduced caryophyllene, while high-dose ABA (100 μM) caused even greater suppression, reinforcing the antagonistic relationship between ABA, SA and JA-mediated sesquiterpene production under HS (F_9,60 = 27.45, p = 0.0001). Our results indicate that ABA acts as a negative regulator of caryophyllene biosynthesis during drought stress (Figure 6B). This is evident from the partial recovery of caryophyllene in fluridone-treated plants, where ABA synthesis was inhibited, and the further significant reduction observed when exogenous ABA was applied (F_4,30 = 32.56, p = 0.0001). Under salt stress, caryophyllene levels showed a moderate reduction, but its regulation appeared to be less ABA-dependent (Figure 6B). Although a trend was observed where fluridone slightly maintained caryophyllene levels and exogenous ABA modestly reduced them, these changes were not statistically significant (F_4,30 = 25.41, p = 0.0001).

Humulene also showed the most pronounced heat-induced accumulation among all sesquiterpenes analyzed (Figure 6C). The regulatory pattern closely paralleled those observed for caryophyllene, with DIECA, SHAM and Jarin-1 treatment all significantly suppressing heat-induced humulene biosynthesis. ABT and AIP treatments resulted in decreased humulene level, providing further evidence for the complex SA-mediated regulatory network controlling sesquiterpene metabolisms. Fluridone treatments caused elevated humulene levels, while exogenous ABA application dose-dependently suppressed humulene accumulation (F_9,60 = 29.67, p = 0.0001). Collectively, these data establish that heat-induced sesquiterpene biosynthesis is primarily regulated by coordinated JA, SA and ABA, playing an antagonistic modulatory role. Similarly, the emission of humulene was regulated by an ABA-dependent pathway under both drought and salt stress, mirroring the response observed for caryophyllene (Figure 6C). This was demonstrated by the significant recovery of humulene levels with fluridone treatment and their further suppression upon the application of exogenous ABA (F_4,30 = 24.49, p = 0.0001). In contrast to its clear ABA-dependent regulation under drought, humulene levels under salt stress (SS) showed no significant variation across treatments (F_4,30 = 0.976, p = 0.254) (Figure 6C). The comparable humulene levels in fluridone- and ABA-treated plants suggest its biosynthesis under salt stress is not primarily governed by ABA, but may involve a balance with other hormonal pathways.

2.3.2. Hormone Inhibitor Treatments Induce Stress-Specific Alterations in Endogenous Phytohormone Networks and Cross-Talk Dynamics

To comprehensively understand how pharmacological interventions altered endogenous hormone homeostasis and to validate the specificity of inhibitor effects, we conducted extensive phytohormone profiling across all treatments and stress conditions (Figure 7). Hierarchical clustering of hormone profiles revealed distinct grouping patterns that corresponded to both stress type and inhibitor treatment (Figure 7A). Under HS, IAA and CK levels clustered together, showing coordinated suppression across most treatments, while JA, JA-Ile, SA and ABA formed a separate cluster characterized by stress-induced elevation and differential responses to inhibitors. Principal component analysis of the hormones dataset (Figure 7B) revealed that the first two dimensions captured 82.6% of the total hormonal variance (Dim1: 62.9%, Dim2: 19.7%), effectively discriminating among different treatments. The loading plots demonstrated that JA and JA-Ile contributed most strongly to Dim1, while SA and ABA were the primary drivers of Dim2. IAA and CK showed a negative correlation with stress hormone accumulations, consistent with the antagonistic relationships between growth-promoting and stress-response hormones.

Figure 7.

Integrative phytohormone profiling revealing stress-specific hormone network dynamics and inhibitor induced perturbation. (A) Heatmap with hierarchical clustering showing endogenous hormone levels across all treatments and stress conditions. Colors represent normalized hormone level (red: elevated; blue: reduced) (B) PCA biplot of hormone profile. Loading vectors indicate hormone contribution to sample separation (C) JA content, (D) SA content and (E) ABA contents quantified across treatment. Left panels: Heat stress treatment (yellow bars) (n = 7 plants/treatment). Middle panel: Drought stress treatments (blue bars). Right panel: Salt stress treatment (green bars). Bars represent means ± standard error with individual data points. Different letters denote significant difference among treatments within each stress condition (p < 0.05, ANOVAs followed by Tukey’s HSD test).

The efficacies of the JA biosynthesis inhibitors were confirmed by treatments with Jarin-1, DIECA and SHAM, which collectively reduced JA level by 68% compared to CK_HS. In contrast, inhibitors targeting other pathways (ABT, AIP, fluridone) did not affect JA accumulations. Crucially, exogenous ABA applications under HS significantly induced JA biosynthesis. This provides direct evidence that ABA acts as a positive regulator of JA under HS. These positive regulatory relationships between ABA and JA were consistently observed under DS and SS as well, where JA levels were enhanced and further increased by exogenous ABA (Figure 7C).

SA profiling revealed a complex regulatory pattern distinct from other hormones. SA accumulations under HS were largely independent of JA signaling, as all JA biosynthesis inhibitors maintained significantly elevated SA levels. In contrast, ABA signaling positively regulated SA. This was demonstrated by the inductions of SA following exogenous ABA application, which resulted in the highest observed accumulations. Conversely, inhibition of ABA biosynthesis with fluridone also led to unexpectedly high SA, suggesting a potential feedback mechanism or compensation. Treatment with ABT and AIP significantly reduced SA levels below the HS control, implicating their respective pathways in SA promotion. This positive ABA-SA interaction was conserved under DS and SS, where ABA application consistently further elevated stress-induced SA levels (Figure 7D).

ABA analysis confirmed its central role in orchestrating the stress responses. Under HS, ABA accumulations occurred independently of JA and SA signaling, as their inhibition did not reduce ABA levels. The critical role of ABA biosynthesis was confirmed by fluridone, which suppressed ABA by 64%, while exogenous ABA applications not only restored but dose-dependently elevated the endogenous pool, demonstrating effective uptake and a potential positive feedback loop. This core regulatory module—where stress enhances ABA, fluridone suppresses it and exogenous applications amplify it—was consistently observed under both DS and SS, solidifying ABA as a master regulator of osmotic stress responses (Figure 7E).

3. Discussion

Understanding how crops discriminate among different abiotic stresses and activate appropriate adaptive responses is fundamental for developing climate-resilient varieties [37,40]. Our comprehensive investigation reveals that potato plants employ fundamentally distinct physiological, metabolic and hormonal strategies when confronted with heat, drought and salt stresses, challenging the notion of universal stress response. These stress-specific signatures reflect different adaptive priorities: HS favors continued metabolic activities with an active cooling mechanism; DS prioritizes water conservation through metabolic suppression and SS balances osmotic adjustments with ionic toxicity management.

The contrasting photosynthetic strategies illuminate divergent stress perceptions and response mechanisms [41,42]. HS plants maintained higher gas exchange with enhanced stomatal conductance, enabling evaporative cooling while sustaining carbon assimilation necessary for synthesizing protective proteins and metabolites. These thermo-tolerance strategies, though water-intensive, prove adaptive when HS occur transiently or when water availability remains sufficient [43]. The elevated internal CO₂ despite maintained photosynthesis suggests biochemical limitation in Calvin cycle enzymes, particularly heat-sensitive Rubisco, becomes more constraining than stomatal conductance [44]. Drought stress, conversely, induced severe stomatal closure prioritizing water conservation over carbon gain, representing a fundamental growth and defense trade-off where survival supersedes productivity. The correlation between depleted water contents, collapsed photosynthetic capacity and massive proline accumulations confirms that drought triggers coordinated physiological retrenchment extending beyond simple stomatal limitations to include non-stomatal impairments such as photosystem damages, chlorophyll degradation and metabolic dysfunctions [45,46,47]. Salt stress exhibited intermediate responses, reflecting dual challenges of osmotic stress and ionic toxicities, where crops must balance water conservation with maintaining sufficient transpiration for ion-exclusion while managing accumulated sodium and chloride that disrupt enzymatic activities and membrane integrity [48,49].

The discovery of stress-specific SMs signature represents a significant advance, establishing that different environmental challenges activate distinct biosynthetic programs rather than generalized metabolic responses [10]. The dramatic heat-induced accumulations of sesquiterpenes—particularly humulene, caryophyllene and copaene—suggest these volatile isoprenoids play a functional role in thermo-tolerance, potentially acting as membrane fluidizers counteracting heat-induced rigidification or serving as an airborne signal for systemic acquired thermo-tolerances. The evolutionary conservation of sesquiterpene biosynthesis across crop lineages and its frequent association with stress responses support functional significance beyond mere metabolic by-product [50,51,52]. Drought and salt stress, despite sharing an osmotic component, induced distinctly different metabolite profiles: drought specifically enhanced 1-hexanol and trans-sesquisabinene hydrate while suppressing sesquiterpene, whereas SS elevated copaene and cis-β-farnesene while reducing caryophyllene. These divergent patterns likely reflect different resource allocation strategies and specific biochemical adjustments to osmotic versus ionic challenges, with drought inducing generalized metabolic suppression to conserve resources [53], while SS maintains selective biosynthetic capacities for compounds potentially involved in ion compartmentalization or oxidative-stress mitigations.

Phytohormone profiling revealed a sophisticated stress-discrimination mechanism where hormone-network configurations serve as a molecular signature distinguishing environmental challenges [54,55,56]. The maximal elevations of JA, JA-Ile, SA and ABA under HS, with particularly pronounced JA accumulations, establish jasmonate signaling as central to thermo-tolerance despite JA’s traditionally recognized role in herbivore-defense and wound-responses. This heat-JA connection likely evolved from an ancestral defense pathway that became co-opted for abiotic-stress responses, providing integrated protections against multiple challenges often co-occurring in the natural environment where high temperatures coincide with increased herbivore activities and pathogens pressure [57]. Drought stress induced substantial but moderate hormone elevation compared to heat, with ABA showing the highest relative increase consistent with its canonical role as the primary drought-hormone mediating stomatal closure, osmo-protectant synthesis and dehydration-responsive gene expressions. Also, the unique SS hormonal signature—maintained basal JA/JA-Ile with elevated SA and ABA—suggests qualitatively different signaling requirements where JA-mediated responses become less critical while SA and ABA coordinate osmotic-adjustment and antioxidant defenses against ionic toxicities [58]. Furthermore, the suppression of growth-promoting hormones IAA and CK across all stresses, with CK showing particularly dramatic reductions under SS, reflects fundamental growth defense antagonism where crops temporarily suspend development to reallocate resources toward survival [59]. This trade-off, evolutionarily conserved across plant lineage, represents adaptive-plasticity maximizing survival probability under adverse conditions at the cost of reduced productivity. The severe CK depletion correlating with chlorophyll loss and photosynthetic decline, especially under SS, highlights the essential role of CK in chloroplast maintenance and senescence delay, suggesting that sustaining CK levels through genetic or chemical intervention could mitigate stress-induced yield penalties while maintaining adaptive stress responses [60].

Pharmacological dissections using hormone biosynthesis inhibitors definitively established a causal relationship between specific hormone pathways and stress induced metabolic reprogramming, revealing stress-specific regulatory architecture [31,61]. Under HS, JA biosynthesis inhibitor dramatically suppressed sesquiterpene accumulations, establishing these pathways as the primary regulators of heat-responsive secondary metabolisms. The paradoxical sesquiterpene hyper-accumulations following SA pathway inhibition suggest complex metabolic networks where blocking competing biosynthetic pathway redirect carbon flux toward sesquiterpenes, or where impaired sesquiterpene catabolism causes accumulations [31]. Under DS, ABA biosynthesis inhibition caused the most severe metabolic suppression, while exogenous ABA partially restored metabolite levels, demonstrating ABA’s dominant regulatory roles under water-deficit [62]. This stress-specific regulatory switches—from JA dominance under heat to ABA dominance under drought—represent a sophisticated discrimination mechanism enabling stress-appropriate metabolic programs through differential transcription factor activation and hormone-responsive element engagement [63].

The antagonistic ABA-JA interaction under HS, where high-dose ABA suppressed both endogenous JA accumulation and sesquiterpene biosynthesis, contrasts sharply with the synergistic ABA-JA relationship under DS, revealing context-dependent hormones cross-talk that fine-tunes response based on stress type and intensities [64]. This dynamic interaction likely involves complex feedback loops and competitive transcription factor binding, where elevated ABA under HS may prevent excessive resource diversion to secondary metabolism when water status becomes limiting, while under drought, coordinated ABA-JA signaling optimizes both water conservation and antioxidant defense [65]. From an evolutionary and ecological perspective, these distinct adaptive strategies likely reflect different selective pressures and stress characteristics in the natural environment. Functional validations of stress-specific metabolites through genetic manipulations or exogenous application would clarify whether compounds like sesquiterpenes causally contribute to stress-tolerance or serve as correlated biomarkers [66,67]. Also, the application of plant hormone inhibitors can have off-target effects, potentially influencing other metabolic pathways, which may complicate the interpretation of hormone-specific responses. Field validations under realistic agricultural condition including combined stresses and variable intensities remains essential for translating laboratory insights into practical application.

In conclusion, this study establishes that heat, drought and salt stresses activate fundamentally different adaptive programs in the potato plant governed by stress specific hormone-metabolite regulatory network (Figure 8). Heat stress shows a JA-associated metabolic activation pattern, drought stress exhibits an ABA-driven metabolic suppression and salt stress displays an intermediate multi-hormonal response. These stress-specific signatures provide molecular frameworks for developing climate-resilient cultivars through targeted genetic improvement and precision agriculture strategies. Overall, the findings highlight the key hormonal signatures that differentiate each stress type, without reiterating mechanistic detail, and offer a focused basis for future breeding and stress-management approaches.

Figure 8.

Heat, drought and salt stresses induce distinct metabolic signatures in potato plants through stress specific phytohormones regulation. Pharmacological dissection reveal stress specific hormone-metabolite networks, with differential stomatal responses reflecting underlying physiological adaptation.

4. Materials and Methods

4.1. Plant Culture and Growing Conditions

Potato (Solanum tuberosum L. cv. Diamant) plants were regenerated in vitro using tissue culture techniques following previously established protocols [68]. Plantlets were maintained under controlled conditions at 22 ± 1 °C with a 16 h photoperiod (150 μmol m⁻² s⁻¹ photosynthetic photon flux density; LED T724w, Prime Agrilight Technology, Shenzhen, China). After 3–4 weeks of subculture, plantlets were acclimatized and transplanted into plastic pots (110 × 105 mm) containing peat moss substrate. The transplanted plants were subsequently grown in a greenhouse under the following environmental conditions: 16 h light period at 22 ± 1 °C (day) and 8 h dark period at 18 ± 1 °C (dark), with supplemental lighting providing 500–700 μmol m⁻² s⁻¹ photosynthetic photon flux density (800 W LED grow lights, Lucgrow, Guildford, UK). Plants were watered daily to maintain substrate moisture. After 30 days of acclimatization and growth in the greenhouse, uniform plants were selected for abiotic stress experiments.

4.2. Stress Treatment

4.2.1. Heat Stress Treatments

For heat stress (HS) experiments, plants were divided into two groups and transferred to separate climate-controlled growth chambers (MRC-450C-LED, Prandt, Ningbo, China). Chamber conditions were maintained at 70% relative humidity, a 16 h photoperiod and photosynthetic photon flux density of 500 μmol m⁻² s⁻¹ (FL40SS-W/37 R fluorescent lamps, Osaka, Japan). Control stress (CK_HS) plants were maintained at 22 °C during the light period and 18 °C during the dark period. Heat-stressed plants were exposed to 35 °C during the light period and 22 °C during the dark period, following the same 16 h/8 h light/dark cycle. The elevated temperature regime was selected based on the thermal sensitivity of potato (S. tuberosum), a cool-season crop that exhibits significant physiological stress responses at temperatures exceeding 30 °C [69,70]. Plants were maintained under these conditions for 2 days prior to sampling.

4.2.2. Drought Stress Treatments

For drought stress (DS) experiments, plants were separated into two treatment groups. Well-watered control stress (CK_DS) plants received 150 mL of water per pot every 5 days to maintain soil moisture at field capacity. Drought-stressed plants (DS) were subjected to progressive water deficit by receiving 50 mL of water per pot every 5 days for an initial 20-day period, after which watering was further restricted to maintain soil moisture content at 30–40% of field capacity throughout the remainder of the experiment. To confirm a consistent soil-moisture level across treatments, volumetric water content was monitored using ML3 ThetaProbe soil moisture sensors (Delta-T Device, Cambridge, UK) in all pots [71]. Moreover, leaf osmotic potential was determined in both treatment groups to confirm the establishment of water-deficit stress.

4.2.3. Salt Stress Treatments

In salt stress (SS) treatments, plants were divided into two treatment groups. Control stress (CK_SS) plant received 150 mL of distilled water applied to the basis of each pot, while SS plants received 150 mL of 200 mM NaCl solution applied similarly. Sodium chloride (NaCl) was selected as it represents the most prevalent salt contributing to soil salinization in agricultural systems [72,73]. The 200 mM concentration was chosen based on previous studies demonstrating significant stress responses in solanaceous crops at this salinity level [74,75,76]. Salt treatments were applied every day for 3-days unless specified. Plants were sampled on the fifth day following the initiation of salt stress treatment. Due to the repeated application schedule and evapotranspiration, actual soil salt concentrations may have exceeded the applied concentration by the third day of each application cycle.

4.3. Exogenous Spray of Inhibitors

We employed a comprehensive pharmacological approach using specific inhibitors targeting key hormone biosynthesis and signaling pathways. We applied DIECA (Sigma-Aldrich, MO, USA; an ethylene biosynthesis inhibitor), SHAM (Sigma-Aldrich, MO, USA (an alternative oxidase inhibitor), Jarin-1 (MedChemExpress, Shanghai, China; a JA biosynthesis inhibitor), ABT (MedChemExpress, Shanghai, China; inhibits the activity of benzoic acid 2-hydroxylase), AIP (MedChemExpress, Shanghai, China; inhibitor of phenylalanine ammonia-lyase) and fluridone (MedChemExpress, Shanghai, China; an ABA biosynthesis inhibitor) under heat, drought and salt stress conditions. The stressed plants were individually sprayed with biosynthetic inhibitors. All chemicals were diluted in distilled-water containing 0.02% Tween-20. Distilled water containing 0.02% Tween-20 and 0.01% ethanol was used as a control treatment. The plants were sprayed for 24 h after a 6 h interval before the experiments. Solutions were sprayed with a hand-sprayer onto both surfaces of the leaves until run-off and transferred into separate incubators.

4.4. Physiological Measurements

4.4.1. Gas Exchange Measurements

Photosynthetic gas exchange parameters were measured using a portable infrared gas analyzer system (LI-6400XT, LI-COR Biosciences, Lincoln, NE, USA). Measurements were conducted on fully expanded, mature leaves from the middle canopy of each plant. Leaves were enclosed in the standard 2 cm² leaf chamber under the following controlled conditions: photosynthetic photon flux density of 1500 μmol m⁻² s⁻¹, airflow rate of 500 μmol s⁻¹, ambient CO₂ concentration of 400 μmol mol⁻¹ and vapor pressure deficit maintained at 2.0 kPa. The following parameters were recorded for each treatment: net photosynthetic rate (A), transpiration rate (E), intercellular CO₂ concentration (Ci), stomatal conductance (gs) and instantaneous water use efficiency (WUE, calculated as A/E). Measurements were taken between 09:00 and12:00 to minimize diurnal variation.

4.4.2. Cell Membrane Stability

Cell membrane stability (CMS) was assessed by measuring electrolyte leakage following established protocols [69]. Fully expanded mature leaves were harvested from each treatment group and leaf discs (1 cm diameter) were excised. Initial electrical conductivity (C₁) was measured in deionized water at 25 °C using a compact conductivity meter (B-771, Horiba Ltd., Kyoto, Japan). Samples were then incubated at 25 °C for 24 h with gentle shaking, after which the conductivity was recorded. Subsequently, samples were autoclaved at 121 °C for 20 min to achieve complete membrane disruption, cooled to 25 °C and the final conductivity (C₂) was measured. Cell membrane stability was calculated using the equation: CMS (%) = [1 − (C₁/C₂)] × 100, where higher values indicate greater membrane integrity.

4.4.3. Chlorophyll Content

Total chlorophyll content was determined using a commercial assay kit (Cat# BC0995, Solarbio Science & Technology Co., Beijing, China) according to the manufacturer’s instructions. Fresh leaf samples were rinsed with distilled water, blotted dry and cut into small pieces. Approximately 0.2 g of fresh tissue was homogenized in 2 mL of 80% (v/v) acetone under dim light conditions. The homogenate was transferred to a 15 mL volumetric flask and brought to volume with the extraction solution. Following incubation in darkness for 3 h at 4 °C, samples were centrifuged at 13,000× g for 5 min and the absorbance of the supernatant was measured at 663 nm and 645 nm using a spectrophotometer (ThermoFisher Scientific, Waltham, MA, USA). Total chlorophyll content was calculated according to the equations provided by the kit manufacturer and expressed as mg g⁻¹ fresh weight.

4.4.4. Relative Water Content

Leaf relative water content (RWC) was determined gravimetrically. Fully expanded leaves were excised and immediately weighed to obtain fresh weight (FW). Leaf samples were then immersed in deionized water at room temperature for 5 h under low-light conditions to achieve full turgor, after which they were blotted dry and weighed to determine turgid weight (TW). Subsequently, samples were oven-dried at 75 °C for 24 h until constant weight was achieved and dry weight (DW) was recorded. Relative water content was calculated using the equation: RWC (%) = [(FW − DW)/(TW − DW)] × 100.

4.4.5. Leaf Water Potential

Leaf water potential (Ψ) was measured using the psychometric method. Leaf discs (6 mm in diameter) were excised from fully expanded mature leaves and immediately transferred to sample chambers (C-52, Wescor Inc., Logan, UT, USA) to prevent water loss. After a 30 min equilibration period at room temperature, water potential was measured using a microvoltmeter (HR-33T, Wescor Inc., Logan, UT, USA) operating in hygrometric dew-point mode.

4.5. Measurement of Prolines and MDA Contents

Proline content was determined following a modified ninhydrin-based colorimetric method [69]. Fresh leaf tissue (approximately 0.5 g) was homogenized in 5 mL of 5% (w/v) sulfosalicylic acid and the homogenate was filtered through Whatman No. 1 filter paper. The filtrate (2 mL) was mixed with 2 mL of acid ninhydrin reagent and 2 mL of glacial acetic acid. The reaction mixture was incubated at 100 °C for 1 h in a water bath, then immediately cooled on ice to terminate the reaction. The chromophore was extracted by adding 4 mL of toluene and vigorously mixing for 15–20 s. After phase separation, the absorbance of the toluene phase (upper layer) was measured at 520 nm using a UV-visible spectrophotometer (Beijing Puxi General Instrument Co., Ltd., Beijing, China). Proline concentration was determined using a standard curve prepared with L-proline (0–100 μg mL⁻¹) and expressed as μmol g⁻¹ fresh weight.

Lipid peroxidation was assessed by measuring malondialdehyde (MDA) content using the thiobarbituric acid reactive substances (TBARS) assay. Fresh leaf tissue (0.5 g) was homogenized in 5 mL of 10% (w/v) trichloroacetic acid (TCA) and incubated overnight at 4 °C. The homogenate was centrifuged at 10,000× g for 10 min at 4 °C and 1 mL of the supernatant was mixed with 4 mL of 0.5% (w/v) thiobarbituric acid (TBA) prepared in 20% TCA. The reaction mixture was heated at 95 °C for 30 min in a water bath, then immediately cooled on ice. Following centrifugation at 10,000× g for 10 min, the absorbance of the supernatant was measured at 532 nm and 600 nm using a UV-visible spectrophotometer. The non-specific absorbance at 600 nm was subtracted from the absorbance at 532 nm to eliminate interference from turbidity. MDA concentration was calculated using an extinction coefficient of 155 mM⁻¹ cm⁻¹ and expressed as nmol g⁻¹ fresh weight.

4.6. Quantification of Secondary Metabolites

Secondary metabolite profiling of stressed plant leaves was conducted following an established protocol with minor modifications [77,78]. Fresh leaf samples were harvested from each treatment group, precisely weighed (2.0 g fresh weight), and immediately flash-frozen in liquid nitrogen to preserve metabolite integrity. Samples were stored at −80 °C until extraction.

4.6.1. Metabolites Extraction

Frozen tissue samples were ground to a fine powder under liquid nitrogen using a pre-chilled mortar and pestle. The powdered tissue was extracted with HPLC-grade n-hexane (≥98%, Sigma-Aldrich, St. Louis, MO, USA) at a tissue-to-solvent ratio of 1:10 (w/v) in sealed 22 mL glass vials. Nonyl acetate (≥97%, Sigma-Aldrich) was added as an internal standard at a final concentration of 10 μg mL⁻¹ to enable quantitative analysis. Extractions were performed at room temperature (22 ± 2 °C) for 24 h with continuous agitation on an orbital shaker at 150 rpm. Following extraction, samples were centrifuged at 5000× g for 10 min at 4 °C and the supernatant was filtered through 0.22 μm nylon syringe filters (Millipore, Burlington, MA, USA) to remove particulate matter prior to instrumental analysis.

4.6.2. GC-MS Analysis

Gas chromatography-mass spectrometry (GC-MS) analysis was performed using an Agilent 7890A gas chromatograph coupled to a 5975C inert XL EI/CI mass spectrometer with triple-axis detector (Agilent Technologies, Santa Clara, CA, USA). Chromatographic separation was achieved using an HP-5MS capillary column (30 m × 0.25 mm i.d., 0.25 μm film thickness; Agilent Technologies) with ultra-high purity helium (99.999%) as the carrier gas at a constant flow rate of 1.0 mL min⁻¹. One microliter of sample extract was injected in splitless mode with the injector temperature maintained at 250 °C and a purge flow of 50 mL min⁻¹ activated after 1 min. The oven temperature program was as follows: initial temperature of 40 °C held for 2 min, ramped to 280 °C at 4 °C min⁻¹ and held at 280 °C for 5 min (total run time: 67 min). The transfer line temperature was maintained at 280 °C. Mass spectrometric detection was performed in electron ionization (EI) mode at 70 eV with an ion source temperature of 230 °C and a quadrupole temperature of 150 °C. Data acquisition was performed in full-scan mode over the mass range m/z 35–550 with a scan rate of 3.2 scans s⁻¹ and a solvent delay of 3 min.

4.6.3. Data Processing and Metabolite Identification

Metabolite identification was accomplished by comparing experimental mass spectra and retention indices with reference spectra in the NIST Mass Spectral Database. Tentative identifications required a minimum spectral similarity match of ≥80% and a difference in retention index of ≤20 units when available. Quantification was performed using the internal standard method, whereby metabolite concentrations were calculated based on peak area ratios relative to nonyl acetate and expressed as ng g⁻¹ fresh weight.

4.7. Plant Hormones Analysis

Phytohormone levels in stressed plants were analyzed following a previously established protocol with minor modifications [79,80,81]. Freshly harvested leaf samples were immediately flash-frozen in liquid nitrogen and stored at −80 °C until extraction. Frozen tissue (0.20 g fresh weight) was ground to a fine powder under liquid nitrogen using a pre-chilled mortar and pestle. Phytohormones were extracted using a methanol-water solvent system. Ground tissue was transferred to 2 mL micro-centrifuge tubes and extracted with 1 mL of ice-cold extraction solvent consisting of methanol: water (70:30, v/v) containing 400 μL of ethyl acetate (final ratio 100:20, v/v methanol: ethyl acetate). Deuterium-labeled internal standards were added to each sample at a concentration of 10 ng per sample as follows: D₆-jasmonic acid (D₆-JA), d₆-abscisic acid (d₆-ABA), D₆-salicylic acid (D₆-SA), D₆-jasmonoyl-isoleucine (D₆-JA-Ile), ¹³C₆-indole-3-acetic acid (¹³C₆-IAA) and ²H₆-cytokinin (²H₆-CK). Samples were vortexed vigorously for 30 s and extracted at 4 °C with continuous shaking at 150 rpm in darkness. Following extraction, samples were centrifuged at 12,000× g for 15 min at 4 °C. The supernatant was carefully transferred to clean micro-centrifuge tubes and dried under a gentle stream of nitrogen gas at room temperature. The dried residue was reconstituted in 200 μL of methanol: water (50:50, v/v), vortexed for 30 s and centrifuged at 12,000× g for 10 min at 4 °C. The final supernatant was transferred to 2 mL HPLC auto-sampler vials with crimp caps and stored at −20 °C until LC-MS/MS analysis. Phytohormone concentrations were quantified using the internal standard method, whereby endogenous hormone levels were calculated based on the peak area ratio of each hormone to its corresponding deuterium-labeled internal standard. Quantification was performed using calibration curves prepared from authentic standards at concentrations ranging from 0.1 to 100 ng mL⁻¹. Final hormone concentrations were normalized to sample fresh weight and expressed as ng g⁻¹ FW.

4.8. Statistical Analysis

Differences between treatments were determined using ANOVAs (p ≤ 0.05) and the Tukey HSD test was used to separate means. SMs heatmaps were generated with hierarchical cluster analysis (Euclidean distance, Ward variance methods) via the R package “pheatmap”. PCA analysis was performed using the prcomp function in R. Data visualization and graphical outputs were generated using the ggplot2 package. Illustrations were created using BioRender (https://www.biorender.com/). All analysis was performed using R software (version 4.×; R Core Team).

5. Conclusions

This study establishes that heat, drought and salt stresses activate fundamentally different adaptive programs in the potato plant governed by stress specific hormone-metabolite regulatory network. Heat stress maintains metabolic activity through JA-mediated sesquiterpene biosynthesis, DS induces ABA-dominant metabolic suppression and SS exhibits intermediate multi-hormonal regulation. These stress-specific signatures provide molecular frameworks for developing climate-resilient cultivars through targeted genetic improvement and precision agriculture strategies. The dynamic hormone cross-talk mechanisms revealed here, where regulatory hierarchies shift dramatically depending on stress type, advance our understanding of plant adaptive plasticity and establish foundations for systems-level approaches to engineer optimized stress tolerance in potato and other economically important crops facing increasingly variable environmental conditions.

Abbreviations

The following abbreviations are used in this manuscript:

VOC	Volatile organic compounds
SS	Salt stress
DS	Drought stress
HS	Heat stress
DIECA	Diethyldithiocarbamate
SHAM	Salicylhydroxamic acid
AIP	2-aminoindane-2-phosphonic acid
ABT	1-aminobenzotriazole

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/plants15050844/s1, Figure S1: Water relations in potato plants subjected to different abiotic stresses; Figure S2: Cell membrane integrity and photosynthetic pigment content under abiotic stress conditions; Figure S3: Biochemical markers of osmotic adjustment and oxidative damage in stressed potato plants.

Data Availability Statement

The original contributions presented in this study are included in the article; further inquiries can be directed to the corresponding author.

Conflicts of Interest

The author declares no conflicts of interest.

Funding Statement

This research was supported by Shaqra University, Dawadmi, Saudi Arabia.