Intraspecific priming of thermotolerance by heat-induced volatiles in Antarctic Colobanthus quitensis

Intraspecific priming of thermotolerance by heat-induced volatiles in Antarctic Colobanthus quitensis | Authorea try { document.documentElement.classList.add('js'); } catch (e) { } var _gaq = _gaq || []; _gaq.push(['_setAccount', 'G-8VDV14Y67G']); _gaq.push(['_trackPageview']); (function() { var ga = document.createElement('script'); ga.type = 'text/javascript'; ga.async = true; ga.src = ('https:' == document.location.protocol ? 'https://ssl' : 'http://www') + '.google-analytics.com/ga.js'; var s = document.getElementsByTagName('script')[0]; s.parentNode.insertBefore(ga, s); })(); Skip to main content Preprints Collections Wiley Open Research IET Open Research Ecological Society of Japan All Collections About About Authorea FAQs Contact Us Quick Search anywhere Search for preprint articles, keywords, etc. Search Search ADVANCED SEARCH SCROLL This is a preprint and has not been peer reviewed. Data may be preliminary. 11 September 2025 V1 Latest version Share on Intraspecific priming of thermotolerance by heat-induced volatiles in Antarctic Colobanthus quitensis Authors : Rodrigo Contreras 0000-0001-9970-3125 [email protected] and Gustavo Zuñiga Authors Info & Affiliations https://doi.org/10.22541/au.175760850.09198149/v1 Published Plant, Cell & Environment Version of record Peer review timeline 173 views 133 downloads Contents Abstract Supplementary Material Information & Authors Metrics & Citations View Options References Figures Tables Media Share Abstract Colobanthus quitensis , one of the only two native vascular plants in Antarctica, experiences highly variable summer temperatures that are intensifying under climate change. We investigated whether volatile organic compounds (VOCs) emitted from conspecifics exposed to lethal heat stress could prime adjacent plants to enhance thermotolerance. Plants were exposed to the median lethal temperature (LT50, 58.4°C for 3 h) or maintained as controls, while receiver plants in separate compartments were exposed only to the volatiles from either treatment. Integrated physiological, biochemical, hormonal, metabolomic, and RT-qPCR analyses showed that exposure to LT50 resulted in severe pigment depletion, impaired photosynthesis, oxidative injury, and hormonal reprogramming, accompanied by the accumulation of lipid- and carotenoid-derived VOCs and an induction of heat shock, antioxidant, and stress-responsive genes. Receivers of VOCs from heat-stressed plants displayed intermediate photosynthetic activity and oxidative stress, anticipatory activation of antioxidant enzymes, elevated basal levels of abscisic and jasmonic acids, increased accumulation of flavonoids and phenolic acids, and pre-induction of heat shock and defense-associated transcripts. The emitted VOCs included β-ionone, β-cyclocitral, 1-octen-3-ol, hexanal, and 2-pentylfuran, implicating apocarotenoid and oxylipin pathways in airborne priming. It was observed that, even in polar ecosystems, airborne signals can coordinate intraspecific preparation for acute thermal stress. This mechanism is suggested to contribute to improved physiological performance and to the resilience of plant communities under rapid regional warming. Intraspecific priming of thermotolerance by heat-induced volatiles in Antarctic Colobanthus quitensis Rodrigo A. Contreras 1, * and Gustavo E. Zúñiga 1,2, ** ¹ Laboratorio de Fisiología y Biotecnología Vegetal, Departamento de Biología, Facultad de Química y Biología, Universidad de Santiago de Chile, Santiago, Chile 2 Centro para el Desarrollo de la Nanociencia y la Nanotecnología (CEDENNA), Universidad de Santiago de Chile, Santiago, Chile *Correspondence: * [email protected] ; [email protected] ** [email protected] Funding : This work was supported by the National Agency for Research and Development (ANID, Chile) through FONDECYT project No. 3160274. Current address: Dr. Rodrigo A. Contreras, Scientific Research Unit; Research and Development Department, The Not Company. Quilín 3550, Macul, Santiago, Chile. ABSTRACT Colobanthus quitensis , one of the only two native vascular plants in Antarctica, experiences highly variable summer temperatures that are intensifying under climate change. We investigated whether volatile organic compounds (VOCs) emitted from conspecifics exposed to lethal heat stress could prime adjacent plants to enhance thermotolerance. Plants were exposed to the median lethal temperature (LT50, 58.4°C for 3 h) or maintained as controls, while receiver plants in separate compartments were exposed only to the volatiles from either treatment. Integrated physiological, biochemical, hormonal, metabolomic, and RT-qPCR analyses showed that exposure to LT50 resulted in severe pigment depletion, impaired photosynthesis, oxidative injury, and hormonal reprogramming, accompanied by the accumulation of lipid- and carotenoid-derived VOCs and an induction of heat shock, antioxidant, and stress-responsive genes. Receivers of VOCs from heat-stressed plants displayed intermediate photosynthetic activity and oxidative stress, anticipatory activation of antioxidant enzymes, elevated basal levels of abscisic and jasmonic acids, increased accumulation of flavonoids and phenolic acids, and pre-induction of heat shock and defense-associated transcripts. The emitted VOCs included β-ionone, β-cyclocitral, 1-octen-3-ol, hexanal, and 2-pentylfuran, implicating apocarotenoid and oxylipin pathways in airborne priming. It was observed that, even in polar ecosystems, airborne signals can coordinate intraspecific preparation for acute thermal stress. This mechanism is suggested to contribute to improved physiological performance and to the resilience of plant communities under rapid regional warming. Keywords: Antarctica; Colobanthus quitensis ; thermotolerance; volatile organic compounds INTRODUCTION Colobanthus quitensis is one of the two native vascular plant species in Antarctica. This species has developed traits that permit survival under the harsh polar climate. Its evolutionary origin, however, has long been debated. Some authors have argued that it may represent the survival of an ancient Gondwanan lineage, whereas others have suggested that it reached Antarctica through more recent long-distance dispersal. Fossil pollen attributable to Caryophyllaceae has been reported from the Southern Hemisphere since the Late Cretaceous, when Antarctica was still connected to South America and Australia, suggesting a possible vicariant origin. However, repeated glaciation cycles since the Eocene markedly reduced the likelihood of continuous survival, confining vascular plants to coastal refugia during interglacial periods. Genetic studies have shown slight variation among Antarctic populations of C. quitensis . Such a pattern is generally interpreted as evidence of secondary dispersal, probably mediated by birds and reinforced by episodic expansion during warmer intervals. Within this framework, C. quitensis is frequently considered a migratory relict. It was likely established in Antarctica during the Oligocene–Pliocene, when the continent was less isolated and climatic conditions were more favorable, and it has persisted since then in local refugia, with recent warming enabling opportunistic range expansión (Parnikoza et al., 2007). Antarctica is often described as uniformly cold, but the Antarctic Peninsula (the main distribution area of C. quitensis ) has undergone marked warming in recent decades (Cannone et al., 2022). C. quitensis has expanded in vegetated areas such as Signy Island (60°43′ S, 45°38′ W), where ground cover increased markedly between 2009 and 2018 (Cannone et al., 2022). It has been shown that experimental warming increases photosynthetic activity and growth in this species (Sáez et al., 2018). From this, it can be inferred that higher summer temperatures may favor demographic expansion and geographic spread. Plant priming was described as an enhanced, faster, or stronger response to a later stimulus following a previous non-damaging cue. Through heat priming, a mild thermal episode can establish a physiological and molecular memory, and this memory improves resilience when plants face later heat stress (Hilker & Schmülling, 2019). Recent studies have described mobile signals in the priming of neighboring individuals without direct exposure to stress. This plant-to-plant communication is mediated by volatile organic compounds (VOCs) (Engelberth & Engelberth, 2019). However, whether VOC-mediated priming occurs in C. quitensis , a species of disproportionate ecological importance and climatic sensitivity, remains unknown. Here, we evaluated the hypothesis that airborne cues emitted by C. quitensis exposed to LT 50 conditions prime conspecific neighbors for enhanced physiological and molecular resilience. Using a controlled hydroponic system combined with precise thermal treatments, we integrated physiological, biochemical, hormonal, metabolomic, and RT-qPCR analyses to resolve how VOCs orchestrate multiscale stress responses in this Antarctic vascular plant. Our results showed that VOCs released from thermally challenged individuals can precondition neighboring plants, aligning their physiological status and gene expression towards enhanced readiness even in one of Earth’s coldest frontiers. From these results, a new dimension of resilience in polar plant communities can be recognized. It is also suggested that VOC-mediated priming can influence ecosystem responses under conditions of accelerated regional warming (Cannone et al., 2022; Engelberth & Engelberth, 2019; Hilker & Schmülling, 2019; Sáez et al., 2018). MATERIALS AND METHODS Plant material and growth conditions Adult C. quitensis plants were manually collected during the 2016–2017 growing season in the vicinity of the Brazilian Antarctic Station Comandante Ferraz, King George Island, South Shetland Islands (62°05′0″ S; 58°23′28″ W), under permits issued by the Chilean Antarctic Institute (INACH) and Brazilian Antarctic Program (PROANTAR). Sampling was done with nitrile gloves and stainless-steel scissors sterilized in 70% (v/v) ethanol. Plant material was immediately placed in polyethylene bags with moist paper to maintain turgor and transported in insulated boxes at 4 °C to the Plant Physiology and Biotechnology Laboratory, University of Santiago, Chile. In the laboratory, plants were washed with sterile distilled water (5 min) and surface-sterilized with 70% ethanol (30 s), followed by 1.5% (v/v) sodium hypochlorite containing 0.1% (v/v) Tween-20 (15 min, gentle agitation). Explants were rinsed five times with sterile distilled water and established in 250 mL Magenta® glass jars containing 15 mL Murashige and Skoog medium (MS; Phytotechnology) (Murashige & Skoog, 1962) supplemented with 3% (w/v) sucrose, 2.0% (w/v) high-acyl gellan gum (Phytagel®, Sigma-Aldrich), 0.1 mg L -1 naphthaleneacetic acid (NAA), and 0.25 mg L -1 benzylaminopurine (BAP) at a 2:1 molar ratio. The medium was adjusted to pH 4.7 ± 0.1 before sterilization (Contreras et al., 2019; Zúñiga et al., 2009). Cultures were maintained at 12 ± 1 °C under a 16 h light/8 h dark photoperiod with 150 µmol m -2 s -1 irradiance (white-spectrum LED lamps). Subcultures were performed every 28 days. Healthy and vigorous plantlets were placed in individual seedling trays (500 mL of substrate solution) with Hoagland n° 2 nutrient solution for the hydroponic acclimation. The trays were aerated using air pumps and ceramic diffusers. The solution was freshly prepared with distilled water, adjusted to pH 4.7 ± 0.1, and sterilized by autoclaving. Its electrical conductivity was 1.2 ± 0.05 mS cm -1 . The nutrient solution was replaced once per week (Contreras et al., 2019). Determination of the median lethal temperature (LT 50 ) The LT 50 was determined using whole C. quitensis plantlets previously acclimated to hydroponic conditions. Because of the small size of this species, leaves showing visible chlorosis or prior damage were removed to ensure that the assay was conducted with intact plants free of visible damage. Each plantlet was rinsed three times with ultrapure water and then placed in glass tubes containing 3 mL of the same water. The tubes were exposed to a temperature gradient from 12 to 80 °C on a dry heating plate for 3 h. After treatment, the samples were left to rest for 1 h at room temperature before the initial conductivity (C1) was recorded with a conductivity meter (Cortés-Antiquera et al., 2021). To determine the final conductivity (C 2 ), the same samples were incubated at 100 °C for 15 min. The percentage of damage was calculated according to Eq. (1): \(\text{Damage\ }\left(\%\right)=\ \left(\frac{C_{1}}{C_{2}}\right)\times 100\)(1) C 1 and C 2 refer to the initial and final conductivities, respectively. The values were then fitted to a four-parameter logistic model in GraphPad Prism 10.5 (GraphPad Software, San Diego, CA, USA). LT 50 was estimated as the inflection point of the curve at which 50% damage was observed on the y-axis. Experimental design for priming Adult C. quitensis plants previously acclimated to hydroponic conditions were randomly assigned to four independent experimental groups: 1. Control (CEU): untreated plants. 2. LT 50 : plants exposed to a heat shock of 58.4 °C for 3 h. 3. Volatile receiver (control; VRC): plants exposed to volatiles emitted by control plants. 4. Volatile receiver (primed; VRP): plants exposed to volatiles emitted by LT 50 -treated plants. Each treatment included six independent biological replicates. We carried out the exposures in borosilicate glass chambers with two compartments (1 L each). The compartments were linked by polytetrafluoroethylene (PTFE) tubing, 6 mm in inner diameter. Airflow was kept constant at 0.8 L min -1 . The air came from an oil-free pump, was passed through activated charcoal and a 0.22 µm polyvinylidene fluoride (PVDF) filter and then humidified before it reached the chambers. For the LT 50 group, plants were placed in one compartment and directly subjected to heat shock using a pre-equilibrated water bath. In the volatile receiver groups, plants remained in the opposite compartment, exposed only to the airflow carrying volatiles emitted by the plants in the emitter compartment (control or LT 50 , depending on treatment). The airflow was maintained throughout the exposure period, matching the duration of the heat treatment in the emitter compartment. Temperature in both compartments was continuously monitored with thermocouples. Photosynthetic pigment analysis Chlorophyll and total carotenoid contents were quantified from fresh leaves of C. quitensis. To do this, 50 mg of fresh tissue were weighed (fresh weight, FW) and reserved for pigment analysis. In parallel, a portion of tissue was oven-dried at 60 °C to constant weight to determine the dry weight (DW). Water content (WC) was calculated as shown in Eq. (2): \(\text{Water\ content\ }\left(\%\right)=\ \left[\frac{\left(FW-DW\right)}{\text{FW}}\right]\times 100\)(2) The fresh-to-dry weight conversion factor was calculated according to Eq. (3): \(Conversion\ factor=\ \frac{\text{FW}}{\text{DW}}\) (3) For pigment determination, the fresh tissue (50 mg) was ground in liquid nitrogen with a mortar and pestle and homogenized in 1 mL of cold 80% (v/v) acetone. Samples were incubated in the dark at 4 °C for 24 h to ensure complete pigment extraction. Extracts were centrifuged at 12,000 × g for 5 min at 4 °C, and the supernatant was recovered for spectrophotometric analysis. Absorbance of extracted pigments was measured at 662, 645, and 470 nm with an Agilent 8453 diode-array spectrophotometer (Agilent Technologies, Santa Clara, CA, USA). An 80% (v/v) acetone solution was used as the blank. The concentrations of chlorophyll a (C a ), chlorophyll b (C b ), and total carotenoids (C x+c ) were then calculated using Eqs. (4–6) (Lichtenthaler & Wellburn, 1983): \(C_{a}\ \left(\frac{\text{mg}}{L}\right)=12.25\ \times\ A_{662}-2.79\ \times\ A_{645}\ \)(4) \(C_{b}\ \left(\frac{\text{mg}}{L}\right)=21.50\ \times\ A_{645}-5.10\ \times\ A_{662}\)(5) \(C_{x+c}\left(\frac{\text{mg}}{L}\right)=\frac{\left(1000\ \times\ A_{470}-1.82\ \times\ \left[\text{Chlorophyll\ a}\right]-\ 85.02\times[Chlorophyll\ b]\right)}{198}\)(6) The pigment content was expressed in mg g -1 of dry weight, correcting for the final extract volume and applying the FW/DW factor obtained previously, according to Eq. (7). The ratios of C a /C b and (C a +C b )/C x+c were also calculated. \(\text{Pigment\ }\left(\frac{\text{mg}}{\text{g\ DW}}\right)=\frac{\left[\text{Concentration\ }\left(\frac{\text{mg}}{L}\right)\times Extract\ volume\ (L)\right]}{m_{\text{DW}}\ (g)}\)(7) where C is the pigment concentration, V is the extract volume, and m DW is the dry sample mass. Measurement of photosynthetic efficiency and electron transport rate (ETR) curves The maximum photochemical efficiency of photosystem II (F v /F m ) and electron transport rate (ETR) curves were determined using a Hansatech FMS-2+ portable modulated fluorometer (Hansatech Instruments Ltd., UK) equipped with a leaf clip and light intensity control. Measurements were performed on intact plants after a 30 min dark-adaptation period to allow complete oxidation of PSII reaction centers. Baseline fluorescence (F 0 ) was recorded with a low-intensity modulated light (<1 µmol photons m -2 s -1 ). A saturating pulse of white light (3000 µmol photons m -2 s -1 , 0.8 s) was then applied to record the maximum fluorescence (F m ). Maximum photochemical efficiency was calculated according to Eq. (8): \(Maximum\ photochemical\ efficiency=\frac{F_{v}}{F_{m}}=\frac{(F_{m}-F_{0})}{F_{m}}\)(8) ETR curves were obtained by stepwise increases in irradiance (0–1000 µmol photons m -2 s -1 ), maintaining each level for 60 s before recording steady-state and maximum fluorescence values. The apparent electron transport rate was calculated as shown in Eq. (9): \(ETR=\ \Phi_{\text{PS\ II}}\times PPFD\times\alpha_{\text{leaf}}\times\ f_{\text{PS\ II}}\)(9) where: • ETR = apparent electron transport rate (µmol e - m -2 s -1 ) • \(\Phi_{\text{PS\ II}}\ \)= effective quantum yield of PSII (dimensionless) • PPFD = incident photosynthetic photon flux density (µmol photons m -2 s -1 ) • \(\alpha_{\text{leaf}}\ \)= leaf absorptance (dimensionless, typically 0.84) • \(f_{\text{PS\ II}}\) = fraction of excitation energy distributed to PSII (dimensionless, typically 0.5) ETR curves were fitted to the exponential plateau model in GraphPad Prism v10.5, following Eq. (10): \(y=y_{0}+(plateau-\ y_{0})\times(1-e^{-K\times x})\) (10) Quantification of oxidative damage The total content of reactive oxygen species (ROS) and the extent of membrane lipid peroxidation were quantified in fresh leaves of C. quitensis . Determination of total ROS The total ROS content was quantified with a fluorometric assay that relies on the oxidation of 2′,7′-dichlorodihydrofluorescein diacetate (DCFH-DA). The method was adapted for small sample volumes in 96-well plates (Contreras et al., 2019). Fresh tissue (100 mg) was incubated for 60 min at room temperature in the dark in 10 µM DCFH-DA prepared with 50 mM Tris–HCl buffer (pH 8.0). The incubation was carried out with gentle agitation to improve dye uptake. After this step, the samples were rinsed with 50 mM EDTA to remove excess probe, surface-dried, and ground in liquid nitrogen. The resulting powder was homogenized in the same buffer, and the homogenate was centrifuged at 10,000 × g for 10 min at 4 °C. The supernatant was filtered through Miracloth and dispensed into black, flat-bottom 96-well plates. Fluorescence was recorded using a Tecan Infinite® M200 Pro microplate reader (Tecan Group Ltd., Switzerland) with excitation at 488 nm and emission at 525 nm. Quantification was performed by interpolation against a standard oxidized dichlorofluorescein (DCF) curve, and results were expressed as nmol DCF equivalents g -1 DW. Determination of lipid peroxidation (TBARS) The membrane lipoperoxidation was evaluated by quantifying thiobarbituric acid-reactive substances (TBARS) with microscale modifications (Contreras et al., 2019). Fresh tissue (50 mg) was homogenized in 1% (v/v) trichloroacetic acid (TCA) and centrifuged at 10,000 × g for 5 min at 4 °C. From the supernatant, 250 µL were mixed with 1 mL of 0.5% (w/v) thiobarbituric acid (TBA) prepared in 20% (v/v) TCA. Samples were incubated in a water bath at 100 °C for 30 min, rapidly cooled on ice, and centrifuged to remove precipitates. Absorbance was recorded at 532 nm, with nonspecific absorbance corrected at 600 nm. According to Eq. (11), the malondialdehyde (MDA) concentration was calculated using the molar extinction coefficient 155 mM -1 cm -1 . \(\text{MDA}\left(\text{nmol\ }g^{-1}\text{DW}\right)=\frac{(A_{532}-A_{600})}{\varepsilon\times l}\times\frac{10^{6}}{m_{\text{DW}}}\)(11) where A 532 -A 600 is the corrected absorbance, ε is the molar extinction coefficient (155 mM -1 cm -1 ), l is the path length (cm), and m DW is the dry weight of the sample (g). Results were expressed as nmol MDA g -1 DW. Protein extraction and enzyme activity assays Total soluble proteins were extracted from 100 mg of fresh C. quitensis tissue, previously ground in liquid nitrogen, was homogenized using 50 mM sodium phosphate buffer (pH 7.5) at 4 °C. The suspension was centrifuged at 10,000 × g for 10 min at 4 °C, and the resulting supernatant was used for enzyme activity assays. Protein concentration was determined using the Bradford method (Bradford, 1976), using bovine serum albumin (BSA) as a standard and expressed as mg protein g -1 DW. Superoxide dismutase (SOD; EC 1.15.1.1) SOD activity was assessed through its ability to inhibit the photochemical reduction of nitro blue tetrazolium (NBT) in the presence of methionine and riboflavin. The reaction mixture contained 50 mM phosphate buffer (pH 7.5), 0.16 mM EDTA, 21.7 mM methionine, 0.5 mM NBT, 33 µM riboflavin, and 20 µg of protein. The samples were exposed to white light, and NBT reduction was monitored spectrophotometrically at 560 nm. A dark-incubated reaction was included as a control. One unit of SOD was defined as the amount of enzyme required to inhibit NBT photoreduction by 50%. Specific activity was expressed as enzymatic units (EU) per gram of protein according to Eq. (12): \(\text{SOD\ activity\ }\left(\text{EU\ }g^{-1}\text{\ protein}\right)=\frac{U}{m_{\text{protein}}}\)(12) where U corresponds to the units of enzyme (50% inhibition of NBT photoreduction) and m protein is the total protein mass (mg) in the reaction mixture (Contreras et al., 2018). Ascorbate peroxidase (APX; EC 1.11.1.11) APX activity was determined by monitoring the decrease in absorbance at 290 nm associated with ascorbate oxidation. The final reaction mixture contained 50 mM phosphate buffer (pH 7.5), 0.4 mM ascorbate, 0.15 mM hydrogen peroxide, and 50 µg of protein. Enzyme activity was calculated using the molar extinction coefficient of ascorbate (ε = 2.8 mM -1 cm -1 ) according to Eq. (13): \(\text{APX\ activity\ }\left(\text{µmol\ }\min^{-1}\ \text{mg}^{-1}\text{\ protein}\right)=\frac{\mathrm{\Delta}A_{290}/(\varepsilon\times l)}{m_{\text{protein}}}\)(13) where ΔA 290 is the change in absorbance per minute, ε is the molar extinction coefficient of ascorbate (2.8 mM -1 cm -1 ), l is the path length of the cuvette (cm), and m protein is the total protein mass (mg) in the reaction mixture (Contreras et al., 2018). Catalase (CAT; EC 1.11.1.6) CAT activity was determined by monitoring the degradation of H 2 O 2 at 240 nm. The final reaction mixture contained 50 mM phosphate buffer (pH 7.5), 0.15 mM H 2 O 2 , and 20 µg of protein. Enzyme activity was calculated using the molar extinction coefficient of H 2 O 2 (ε = 39.4 M -1 cm -1 ) and expressed as µmol H 2 O 2 decomposed min -1 mg -1 protein according to Eq. (14): \(\text{CAT\ activity\ }\left(\text{µmol\ }\min^{-1}\ \text{mg}^{-1}\text{\ protein}\right)=\frac{\mathrm{\Delta}A_{240}/(\varepsilon\times l)}{m_{\text{protein}}}\)(14) where ΔA 240 is the change in absorbance per minute, ε is the molar extinction coefficient of H 2 O 2 (39.4 M -1 cm -1 ), l is the cuvette path length (cm), and m protein is the total protein mass (mg) in the reaction mixture (Contreras et al., 2018). Total peroxidases (Guaiacol peroxidase; POD; EC 1.11.1.7) POD activity was determined by measuring tetraguaiacol formation at 470 nm. The final reaction mixture contained 50 mM phosphate buffer (pH 7.5), 0.5 mM guaiacol, 0.15 mM H 2 O 2 , and 20 µg of protein. Enzyme activity was calculated using the molar extinction coefficient of tetraguaiacol (ε = 26.6 mM⁻¹ cm⁻¹) and expressed as µmol tetraguaiacol formed min -1 mg -1 protein according to Eq. (15): \(\text{POD\ activity\ }\left(\text{µmol\ }\min^{-1}\ \text{mg}^{-1}\text{\ protein}\right)=\frac{\mathrm{\Delta}A_{470}/(\varepsilon\times l)}{m_{\text{protein}}}\)(15) where ΔA 470 is the change in absorbance per minute, ε is the molar extinction coefficient of tetraguaiacol (26.6 mM - ¹ cm - ¹), l is the cuvette path length (cm), and m protein is the total protein mass (mg) in the reaction mixture (Contreras et al., 2018). Glutathione reductase (GR; EC 1.6.4.2) GR activity was determined by monitoring the oxidation of NADPH at 340 nm for 3 min. The reaction mixture (final volume 1 mL) contained 50 mM phosphate buffer (pH 7.5), 2 mM EDTA, 0.15 mM NADPH, 0.5 mM oxidized glutathione (GSSG), and 100 µg of protein. The decrease in absorbance was recorded spectrophotometrically, and enzyme activity was calculated using the molar extinction coefficient of NADPH (ε = 6.2 mM -1 cm -1 ) according to Eq. (16): \(\text{GR\ activity\ }\left(\text{µmol\ }\min^{-1}\ \text{mg}^{-1}\text{\ protein}\right)=\frac{\mathrm{\Delta}A_{340}/(\varepsilon\times l)}{m_{\text{protein}}}\)(16) where ΔA 340 is the change in absorbance per minute, ε is the molar extinction coefficient of NADPH (6.2 mM - ¹ cm - ¹), l is the cuvette path length (cm), and m protein is the total protein mass (mg) in the reaction mixture (Contreras et al., 2018). Phenylalanine ammonia-lyase (PAL; EC 4.3.1.5) The PAL activity was determined by spectrophotometric monitoring of the conversion of L-phenylalanine into trans-cinnamic acid at 290 nm. The reaction mixture contained 100 mM sodium borate buffer (pH 8.8), 2 mM L-phenylalanine, 1% (w/v) PVPP, 1 mM DTT, and 50 µg of protein. Reactions were incubated at 37 °C for 2 h and stopped with 0.6 N HCl. Parallel reactions were performed using D-phenylalanine as substrate to correct for non-enzymatic background absorbance. Enzyme activity was calculated from the difference in absorbance between L- and D-phenylalanine reactions (ΔA 290 ), using the molar extinction coefficient of trans-cinnamic acid (ε = 9,630 M -1 cm -1 ), and expressed as Mkat kg -1 protein according to eq. (17): \(\text{PAL\ activity\ }\left(\text{Mkat\ }\text{kg}^{-1}\text{\ protein}\right)=\frac{\mathrm{\Delta}A_{290}/(\varepsilon\times l)}{m_{\text{protein}}}\)(17) where ΔA 290 is the L–D difference in absorbance per minute, ε is the molar extinction coefficient of trans-cinnamic acid (9,630 M -1 cm -1 ), l is the cuvette path length (cm), and m protein is the total protein mass (mg) in the reaction mixture (Contreras et al., 2019). Phytohormone analysis Endogenous phytohormones were quantified from 50 mg of fresh C. quitensis leaves, finely ground in liquid nitrogen using a pre-chilled mortar and pestle. Each sample was spiked with a mixture of deuterated internal standards (final concentration 50 nM), followed by 500 µL of cold 2-propanol: water: HCl (2:1:0.002, v/v/v). The mixture was shaken at 100 rpm for 30 min at 4 °C, then 1 mL of dichloromethane was added, and samples were centrifuged at 13,000 × g for 5 min at 4 °C. The lower organic phase was collected, evaporated under a nitrogen stream, and the dry residue was reconstituted in 100 µL of LC–MS grade methanol (Pan et al., 2010). Separation and detection were performed using an Agilent 1260 HPLC system coupled to an Agilent 6410 triple quadrupole mass spectrometer (Agilent Technologies, Santa Clara, CA, USA) equipped with a Zorbax Eclipse C18 column (150 × 4.6 mm, 5.0 µm). The mobile phases consisted of water (A) and acetonitrile (B), both containing 0.1% formic acid, at a flow rate of 0.6 mL min⁻¹. The elution program was as follows: 0–2 min, 30% B; 2–20 min, linear increase to 100% B; 20–22 min, 100% B; 22–25 min, return to 30% B. Detection was performed in multiple reaction monitoring (MRM) mode by electrospray ionization (ESI), in either positive or negative mode depending on the analyte. The specific MRM transitions were: m/z 263.0 → 153.0 for abscisic acid (ABA), 209.1 → 59.0 for jasmonic acid (JA), 175.0 → 130.0 for indole-3-acetic acid (IAA), 331.1 → 213.0 for gibberellin A 4 (GA 4 ), 137.0 → 93.0 for salicylic acid (SA), 102.0 → 56.0 for 1-aminocyclopropane-1-carboxylic acid (ACC), and 220.2 → 136.2 for trans-zeatin (tZ). All concentrations were expressed as nmol g -1 DW and corrected for recovery using the internal standards (Contreras et al., 2025). VOCs VOCs were collected by solid phase microextraction (SPME) using 65 µm PDMS/DVB fibers (Supelco, Bellefonte, PA, USA). Preconditioned SPME fibers were exposed for 2 h inside the emitter chamber (control or LT 50 plants). Chromatographic analysis was performed using a GC–MS 7890B–5977B system (Agilent Technologies, Santa Clara, CA, USA) equipped with an HP-5MS UI capillary column (30 m × 0.25 mm × 0.25 µm). Helium (99.999% purity) was used as the carrier gas at a constant flow of 1.0 mL min -1 . The injector was maintained at 250 °C in splitless mode for the first 3 min. The oven program was 40 °C for 3 min, ramped at 5 °C min -1 to 250 °C, and held for 5 min. The MS transfer line temperature was set at 280 °C. Ionization was carried out by electron impact (70 eV), and mass spectra were acquired in the range of m/z 35–350 at a scan rate of 3.2 scans s -1 . Compound identification was achieved by comparing spectra with the NIST 17 library and verifying experimental retention indices calculated from a homologous series of C8–C20 n-alkanes (Supelco). Soluble metabolite analysis Methanolic extracts of C. quitensis were prepared and adjusted to a concentration equivalent to 10 mg mL-1 dry extract. Before to the analysis, extracts were filtered through 0.45 µm PVDF membranes. Chromatographic separation was performed using an Agilent 1260 Series HPLC system coupled to an Agilent 6410 triple quadrupole mass spectrometer (Agilent Technologies, Santa Clara, CA, USA), equipped with a Zorbax Eclipse C18 reversed-phase column (150 × 4.6 mm, 5.0 µm) and a C18 guard column. The column temperature was maintained at 25 °C. Mobile phase (A) was 10 mM ammonium acetate (pH 4.0) in water, while mobile phase (B) was 10 mM ammonium acetate (pH 4.0) in 95% (v/v) acetonitrile. The elution gradient was programmed as follows: from 0 to 5 min, 0–50% B; from 5 to 10 min, 50–52% B; from 10 to 20 min, 52–100% B; from 20 to 35 min, 100–65% B; and from 35 to 40 min, 65–0% B. The flow rate was maintained at 0.6 mL min⁻¹, and the injection volume was set at 20 µL. Detection was carried out by electrospray ionization (ESI) in negative and positive modes, with a capillary voltage of 4000 V, nebulizer pressure of 310 kPa, drying gas (nitrogen) flow of 9 L min -1 , and gas temperature of 330 °C. Data acquisition was performed in scan mode, covering a mass range of m/z 50–1000. Compound identification was achieved by comparing MS/MS spectra with the METLIN database and, when available, with authentic reference standards. Quantification was performed using calibration curves prepared with structurally related standards, and results were expressed as nmol g -1 DW. Gene expression analysis by RT-qPCR Total RNA was extracted from fresh C. quitensis leaves using the Spectrum™ Plant Total RNA Kit (Sigma-Aldrich, USA), following the manufacturer’s instructions and including on-column DNase I digestion (On-Column DNase I Digest Set, Sigma-Aldrich) to remove genomic DNA. 100 mg of tissue were ground in liquid nitrogen and homogenized in the lysis solution supplemented with 0.8% (v/v) 2-mercaptoethanol. RNA concentration and purity were assessed spectrophotometrically using a NanoQuant Plate (Tecan, Austria), based on A 260 /A 280 and A 260 /A 230 ratios, accepting only samples within the range of 1.8–2.1. RNA integrity was further verified by denaturing agarose gel electrophoresis (1% w/v) containing 2.2 M formaldehyde in 1× MOPS buffer, confirming sharp and well-defined 25S and 18S rRNA bands. First-strand cDNA was synthesized from 1 µg of total RNA using the AffinityScript qPCR cDNA Synthesis Kit (Agilent Technologies) and oligo(dT) primers in a final volume of 20 µL. Reactions were incubated at 25 °C for 5 min (primer annealing), 42 °C for 15 min (reverse transcription), and 95 °C for 5 min (inactivation). The resulting cDNA was stored at –20 °C until further use. Quantitative PCR was carried out on an Agilent Stratagene Mx3005P thermocycler with Brilliant II SYBR® Green qPCR Master Mix (Agilent Technologies). Each reaction had a final volume of 20 µL and contained 10 µL of 2× master mix, 0.3 µM of each primer, and 1 µL of cDNA diluted 1:10. Cycling conditions were: initial denaturation at 95 °C for 10 min, followed by 40 cycles of 95 °C for 15 s, 60 °C for 30 s, and 72 °C for 30 s. Amplicon specificity was confirmed by melting curve analysis from 55 to 95 °C. The 18S rRNA gene (forward: CCTTACGGCGTTCCCTTACA; reverse: CGGGAAACCATCTTCCTCCC) was used as the internal reference. This marker had been previously validated in C. quitensis as stable under heat stress and other abiotic conditions (Contreras et al., 2019). Relative expression levels were calculated using the 2 ⁻ΔΔCT method (Livak & Schmittgen, 2001), considering three biological and two technical replicates per treatment. Specific primers were designed for genes encoding antioxidant enzymes ( CAT1 , APX1 , SOD1 , POD1 ), stress-related transcription factors ( DREB1A , DREB2A , NAC2 , ZAT12 , HSFA2 ), heat shock proteins ( HSP101 , HSP90 , HSP70 ), late embryogenesis and dehydration-responsive proteins ( RD22 , RD29A , COR15A ), enzymes of phenylpropanoid, flavonoid, and phytohormone biosynthesis ( AOS , LOX2 , ICS1 , PAL1 ), and pathogenesis-related proteins ( PR1 , PR3 , PR5 ). Primer sequences were: CAT1 (F: GGAGCCAGTGCTAAGGGTTT; R: GGATGCCTGGAACCACAAGA), APX1 (F: CTCATGGAGCCAACAGTGGT; R: AAACAGCCATGACTCTCGGG), SOD1 (F: TCCATGAGTTCGGTGACACG; R: TCAGGCCAACAACACCACAT), POD1 (F: ATATTTTGCAGCCGCTTCGC; R: CCTCCAGGTTCTTGCCCATT), DREB1A (F: ACAGAGGAGTTCGTCGGAGA; R: ACGGACAGTGCTCTCTTGTG), DREB2A (F: TTTGGCTCGGTACTTTCCCC; R: CAAAGCAGGACCGCTCATTG), NAC2 (F: TTGCTCCTGGGTTCCGTTTT; R: CTTGTGGCACACCAGCTTTC), ZAT12 (F: CAAGTCGACGGTGGATGTCA; R: ACAAAGCGCGTGTAACCAAC), HSFA2 (F: AACAGCTTTGTGGTGTGGGA; R: ACCTCAGCTACAAGCACACC), HSP101 (F: ATAAGCGGGTTGTGGGACAG; R: GGGGTCGAACACCACAATCT), HSP70 (F: TTGATGCTGCAAACAAGGCG; R: GGTTACGTGTTGCACATCCC), HSP90 (F: TAGCGTTGAGGGACAGCTTG; R: GCCGAAGTGTGAAAGCGAAG), RD22 (F: GAGAACGGGATGCGAGCTAA; R: ACACAACATGAGTCTCCGGG), RD29A (F: TTTGGTGACGAGTCAGGAGC; R: CCGTCAGATTCCGGCGAATA), COR15A (F: GTCGTTGATCTACGCCGCTA; R: TGGCATCCTTAGCCTCTCCT), AOS (F: ACCGTACGATCAAAGCCGTT; R: CGGTAGCCTCCGGTTAGTTC), LOX2 (F: CACATTGAGGCCCTTAGCCA; R: CCATAGCTGGTCGTAGGCAG), ICS1 (F: ATCGGAAACACGCCTGAGAG; R: TTCTGCTGGAAGCCCACAAA), PAL1 (F: CGAAATGGATCCGCTCCAGA; R: TTACGCGCAGAGATGAGTCC), PR1 (F: CCTCGTACATTCTCATGGTCAAT; R: CCATTGTTACACTGAACCCTAGC), PR3 (F: TGTTTTGTTGTGTGTTTTTCCTG; R: AGCAATACCCTCCTGTAAATGGT), PR5 (F: CCGAGGTAATTGTGAGACTGGAG; R: CCTGATTGGGTTGATTAAGTGCA). Statistical Analysis Data for each physiological, biochemical, and molecular parameter were normalized according to the variable type. Pigment concentrations were expressed on a dry mass basis, calculated from the relative water content of each sample, and metabolite content according to the dry extract normalized by original tissue mass. Enzyme activities were expressed relative to total protein content, which was determined with the Bradford assay. Cycle threshold (CT) values were adjusted for gene expression using the 2 ⁻ΔΔCT method, with 18S rRNA as the reference gene. Data were transformed to visualize relative expression as log 2 (x + 1). All statistical analyses were performed using analysis of variance (ANOVA), with either one- or two-way designs depending on the experimental setup. When significant effects were detected, post hoc comparisons were conducted using Tukey multiple comparison tests (HSD). Heatmaps were generated from z-score normalized data, and curve fitting was performed according to the distribution patterns of the data. All analyses and graphical representations were done with GraphPad Prism 10.5 (GraphPad Software, San Diego, CA, USA). RESULTS Experimental design and LT 50 determination Plants of C. quitensis collected in the field were brought to Chile, where they were first established under in vitro conditions. They were propagated and later acclimated to hydroponic culture to generate uniform experimental material. Thermal lethality was assessed on intact healthy plantlets to progressively increasing temperature regimes, and membrane stability was evaluated through electrolyte leakage (conductivity). The LT 50 , defined as the temperature at which 50% damage occurred, was estimated at 58.4 °C after 3 h of exposure (Fig. 1a). This value was subsequently used as the reference for the priming experiment. Four experimental treatments were established: (i) control plants maintained at 12 °C, (ii) plants exposed directly to LT 50 , (iii) volatile receivers exposed to signals from control plants, and (iv) volatile receivers exposed to signals from LT 50 -stressed plants (Fig. 1b). This setup allowed us to test whether airborne cues released during lethal heat stress could prime neighboring conspecifics for improved thermotolerance. Pigments and photosynthetic performance Exposure to LT 50 caused a sharp decrease in photosynthetic pigments, with total chlorophyll and carotenoid contents reduced by up to 55% relative to the control (Fig. 2a–c). Clear impairments matched the decrease in pigments in PSII performance. Reductions of more than 40% were observed in the maximum quantum yield (F v /F m ), the initial photosynthetic efficiency (α), and the integrated area under the light–response curve (AUC) (Fig. 2d–e). These results show that acute heat stress severely disrupts the photosynthetic apparatus. Plant receivers of VOCs, exposed to signals from LT 50 -treated plants, consistently displayed intermediate pigment contents and photosynthetic parameters between control and directly stressed groups. This pattern shows that airborne signals partially alleviated the photosynthetic collapse caused by lethal stress, supporting a priming effect at the physiological level. Oxidative stress and antioxidant activity Exposure to LT 50 significantly increased oxidative burst in C. quitensis , with ROS and TBARS levels increasing more than threefold over control levels (Fig. 3a–b). This response was accompanied by an activation of major antioxidant enzymes, including SOD, CAT, and APX (Fig. 3c–e), consistent with the rapid deployment of enzymatic defenses against oxidative stress. In addition, POD and GR also increased their activities (Fig. 3f–g), supporting both peroxidative detoxification and the glutathione redox cycle. The increase in PAL activity (Fig. 3h) further suggested that phenylpropanoid metabolism contributed an additional layer of protection. VOCs receiver plants exposed to LT 50 donors exhibited a distinct, intermediate phenotype. Although antioxidant enzyme activities were moderately enhanced compared with controls, ROS and TBARS accumulation remained significantly lower than in LT 50 -treated plants. These results suggest that volatile signals preconditioned receivers for a more balanced redox adjustment, reducing membrane peroxidation and linking oxidative homeostasis with secondary metabolic defenses. Hormonal profile LT 50 exposure triggered a hormonal reprogramming in C. quitensis (Fig. 4). Stress-associated phytohormones were strongly induced, with ABA and JA contents increasing up to 4.5- and 3.2-fold relative to controls, accompanied by higher levels of their conjugates and precursors (PA, JA-Ile, MeJA). In contrast, growth-promoting regulators such as GA 4 and trans-zeatin declined significantly, as did SA. These results suggest a shift from developmental and basal defense signals toward stress-responsive pathways. BRs (24-epibrassinolide) and the ethylene precursor ACC showed only moderate changes, suggesting a more selective adjustment within the hormonal network. Receiver plants exposed to LT 50 -stressed VOCs donors, showed anticipatory shifts in hormonal balance. Basal levels of ABA and JA were elevated, whereas GA 4 and SA decrease, even in the absence of direct heat exposure. This primed condition points to a reconfiguration of hormonal homeostasis toward a defensive and alert state, enabling plants to activate protective responses more rapidly and effectively when exposed to subsequent stress. Volatile and non-volatile metabolites GC–MS analysis revealed that LT 50 exposure generates a reconfiguration in the VOCs profile in C. quitensis , with characteristic stress-responsive compounds such as 1-octen-3-ol, hexanal, β-ionone, and 2-pentylfuran accumulating up to sixfold relative to controls (Fig. 5a). These VOCs, carried by the continuous airflow between chambers, are thus the same compounds reaching the receiver plants, supporting their potential role as airborne signals. On the other hand, LC–MS profiling revealed a significant reprogramming of soluble secondary metabolites (Fig. 5b). LT 50 treatment led to increased accumulation of phenolic acids and flavonoid glycosides (including schaftoside, vitexin, and saponarin), together with derivatives of α-linolenic acid and other lipid-associated metabolites. These changes may indicate a coordinated activation of antioxidant, structural, and defense-related metabolic pathways. VOCs receivers consistently displayed intermediate levels of these metabolites, falling between control and LT 50 plants, suggesting that airborne cues are sufficient to trigger partial metabolic reprogramming. This graded response supports the idea that VOC-mediated signaling can shape the biochemical profile of unstressed neighbors toward a primed state, enhancing their readiness to withstand subsequent thermal stress. Gene expression LT 50 treatment induced heat stress–responsive gene expression, such as HSP70 , HSP101 , and HSFA2 , increasing up to 30-fold relative to controls (Fig. 6). Genes associated with jasmonate biosynthesis ( AOS , OPR3 ), abiotic stress regulators ( DREB1A , DREB2A ), and antioxidant enzymes ( APX1 , CAT1 ) were also significantly upregulated. This coordinated transcriptional reprogramming suggests the activation of protective modules spanning chaperone activity, hormonal signaling, and redox homeostasis under lethal heat exposure. Receiver plants exposed to VOCs from LT 50 -stressed donors showed elevated basal expression of many of these same genes despite the absence of direct stress. The anticipatory activation suggests that VOC molecules modulate the transcriptional profile of neighboring plants, preconditioning them for a faster and more effective activation of stress-adaptive pathways during thermal challenge. DISCUSSION Our results provide evidence that C. quitensis perceives and responds to VOCs emitted by conspecifics under acute thermal stress, establishing a primed physiological and molecular state that enhances responses to subsequent heat exposure by integrating physiological, biochemical, hormonal, metabolite profiling, and RT-qPCR datasets. The results indicate that VOC exposure elicits a coordinated reprogramming that limits oxidative damage and sustains photosynthetic performance, even in the absence of direct stress. It is worth noting that the LT 50 applied in this study (58.4 °C, 3 h) represents an extreme form of stress that plants are unlikely to encounter under natural Antarctic conditions. We chose this level of treatment deliberately, as it provided a reproducible lethal threshold that made it possible to uncover defense mechanisms that would be difficult to detect under milder conditions. Thus, the responses observed should be interpreted as simulated trajectories of stress adaptation, providing mechanistic insights rather than direct predictions of field behavior. Direct exposure to LT 50 caused severe loss of photosynthetic pigments, destabilization of PSII, oxidative damage, and canonical activation of antioxidant enzymes. This response is consistent with the expected response in other plant species, where severe stress manifests as a general destabilization of metabolism (Kerchev & Van Breusegem, 2022). On the other hand, VOC-receiving plants maintained intermediate levels of pigments and PSII efficiency, accumulated less MDA, and showed moderate activation of antioxidant defenses. This decrease did not reach a stress threshold but rather reflected a priming adjustment, allowing metabolic reorganization in preparation for the upcoming stress. Overall, this response is consistent with the concept of VOC-mediated priming, which favors pigment retention and PSII efficiency to maintain an active and energetically favorable metabolism (Arimura & Uemura, 2025; Brambilla et al., 2022). Hormonal profiling revealed increased ABA and JA and reduced GA 4 and SA, indicating a reallocation of resources away from growth and SA-dependent defenses toward ABA- and JA-dependent stress pathways (Brosset & Blande, 2022; Peleg & Blumwald, 2011). These changes were reflected at the phytochemical level through the accumulation of glycosylated flavonoids (such as schaftoside, vitexin, saponarin) and phenolic acids, compounds with recognized antioxidant and hormone-modulatory functions (Mitra et al., 2021; Ramel et al., 2012). The VOCs released under LT 50 included carotenoid-derived apocarotenoids (β-ionone, β-cyclocitral) and oxylipin-related compounds (hexanal, 1-octen-3-ol, 2-pentylfuran), carried by the continuous airflow that also reached the receiver plants (Faizan et al., 2022; Felemban et al., 2024; Morote et al., 2023). This composition is derived to two convergent signaling pathways: a carotenoid-dependent one mediated by carotenoid-cleaving dioxygenases (CCDs), which generate retrograde signals for nuclear gene regulation (Shi et al., 2020), and oxylipin biosynthesis mediated by lipoxygenase (LOX), allene oxide synthase (AOS), and 12-oxophytodienoate reductase 3 (OPR3), which includes green leaf VOCs such as (Z)-3-hexenal, known to enhance thermotolerance and antioxidant capacity (Copolovici et al., 2012). A further methodological aspect is that VOCs were collected using SPME in the emitter chamber, which was connected to the receivers by a continuous stream of filtered air. In this way, the sampled headspace corresponded to the same VOCs composition that reached the neighboring plants. By contrast, in natural Antarctic settings, the fate of airborne signals would be primarily shaped by wind, subtle changes in terrain, and boundary layer effects, all of which can strongly alter both the strength and the spatial footprint of VOC dispersion (Ninkovic et al., 2021). Field-based measurements of VOC fluxes will therefore be necessary to determine how effectively such cues propagate across plant communities under realistic conditions. Targeted gene expression analysis supported these pathways, showing induction of HSFA2 , DREB1A and DREB2A , heat shock proteins ( HSPs ), and antioxidant genes in VOC receivers, suggesting that volatile cues activate gene expression networks that reduce response latency under stress. These mechanisms are ecologically relevant in maritime Antarctica, where C. quitensis coexists with Deschampsia antarctica , mosses, lichens, and microbial communities under conditions of limited wind flow and close spatial proximity that may favor volatile exchange (Pascual-Díaz et al., 2020; Vera et al., 2013). Plant–microbe interactions are also likely to modulate the qualitative and quantitative composition of VOCs, shaping signal strength and perception in situ (Bergman et al., 2025; Ninkovic et al., 2021). Given the increasing frequency of summer heat extremes in the Antarctic Peninsula (Turner et al., 2021), VOC-mediated priming may provide a significant adaptive advantage, aligning individual physiology with the demands of a rapidly warming polar environment. However, C. quitensis populations have shown an increase in distribution and ground cover (Cannone et al., 2022), a phenomenon that probably reflects multiple mechanisms, including phenotypic plasticity, associations with microbiota, and potentially volatile-mediated priming. This perspective opens an innovative line of research that links the results obtained here with broader ecological dynamics. An open question is how these mechanisms might play out under gradual, sublethal warming instead of sudden thermal shocks. It is possible that in such progressive scenarios, the release and perception of VOCs could work as signals at the community level, moving through neighboring plants and gradually shaping their collective resilience. Testing this hypothesis in long-term warming experiments will be crucial to determine whether VOC-mediated priming scales from individual responses to ecosystem-level adaptation. Overall, we propose a conceptual model in which acute thermal stress elicits the release of VOCs through carotenoid oxidation, lipid peroxidation, and the activation of secondary metabolism. Neighboring plants integrate these airborne cues into hormonal and transcriptional pathways, pre-activating defenses that limit oxidative stress and help sustain photosynthetic function. We further suggest that volatile-mediated priming forms part of the stress memory of Antarctic plants, linking redox homeostasis with population-level resilience. Recognizing volatile signaling as an element of retrograde communication will be important for predicting the adaptive trajectories of polar ecosystems under continued warming. Concluding remarks Our study supports the idea that, even under extreme conditions such as Antarctica, C. quitensis releases carotenoid- and oxylipin-derived VOCs that contribute to thermotolerance in neighboring plants. This aerial signaling represents an overlooked dimension of resilience that operates intraspecifically, and given the non-exclusive composition of C. quitensis , it could also represent interspecific signaling. By framing volatile communication as part of the adaptive capacity of vascular plants in polar ecosystems, our findings highlight the central role of chemical signaling in shaping stress memory and growth–defense trade-offs, and in influencing community dynamics under the pressures of rapid climate warming. Acknowledgements We thank Dr. Paulo Câmara (Universidade de Brasília, Brasília, DF, Brazil) and the Brazilian Antarctic Program (PROANTAR), including the personnel of Comandante Ferraz Station and the Instituto Antártico Brasileiro, for logistical support. We are especially grateful to Dr. Júlia Viegas Mundim (Universidade de Brasília) and Dr. Marisol Pizarro (Universidad de Santiago de Chile) for assistance in the field, and to Antônio Calvo for logistical and safety support. We also acknowledge Dr. Rocío Santander for advice on initial GC–MS analyses and Héctor Henríquez for technical assistance during GC–MS work. Author contributions Rodrigo A. Contreras: Conceptualization; Methodology; Investigation (fieldwork); Formal analysis; Writing – original draft; Writing – review & editing; Supervision. Gustavo E. Zúñiga: Investigation (experimental development); Formal analysis; Writing – original draft; Writing – review & editing. Both authors contributed equally to study design and interpretation, approved the final version of the manuscript, and share responsibility as corresponding authors. ORCiD Rodrigo A. Contreras: 0000-0001-9970-3125 Gustavo E. 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LT 50 determination and experimental setup for volatile-mediated thermotolerance in C. quitensis . (a) Whole plantlets acclimated to hydroponic culture were exposed to a temperature gradient, and electrolyte leakage was measured to assess membrane damage. A four-parameter logistic regression was fitted to the data (R² = 0.97), with LT 50 estimated at 58.4 °C (95% CI: 58.0–58.8 °C). The red dot represents the LT 50 value evaluated independently after its determination by interpolation (mean ± SE, n = 6). (b) Experimental setup for volatile-mediated priming assays. (1) Native plant collection under Antarctic conditions, (2) in vitro propagation, (3) transfer to hydroponic culture, and (4) exposure in connected glass chambers with unidirectional, charcoal-filtered airflow. In the control setup both plants remained unstressed, whereas in the experimental setup (5) the donor plant was subjected to LT 50 heat stress and (6) headspace volatiles were carried by the airflow to (7) an unstressed receiver plant, with volatile sampling by SPME. Figure 2. Pigment composition and photosynthetic performance in C. quitensis under heat stress and volatile priming. (a) Chlorophyll a and b contents, (b) total carotenoid content, and (c) pigment ratios across experimental treatments. (d) Maximum quantum yield of PSII (Fv/Fm) and (e) light response curves of electron transport rate (ETR) as a function of photosynthetically active radiation (PAR). Treatments included field-grown control, in vitro (pre-hydroponic), hydroponic-acclimated (pre-stress), untreated experimental control, LT 50 -treated (3 h at 58.4 °C), volatile receiver from control plants, and volatile receiver primed from LT 50 donors. Data are means ± SE (n = 6). Different lowercase letters indicate significant differences among treatments (two-way ANOVA for chlorophyll parameters; one-way ANOVA for carotenoids and Fv/Fm), followed by Tukey’s HSD test (P < 0.05). Figure 3. Reactive oxygen species, lipid peroxidation, and antioxidant enzyme activities in C. quitensis under experimental treatments. (a) Total ROS, (b) TBARS, (c) SOD activity, (d) CAT activity, (e) APX activity, (f) POD activity, (g) GR activity, and (h) PAL activity. Treatments included field-grown control, in vitro (pre-hydroponic), hydroponic-acclimated (pre-stress), untreated experimental control, LT 50 -treated (3 h at 58.4 °C), volatile receiver from control plants, and volatile receiver primed from LT 50 donors. Values are means ± SE (n = 6). Different lowercase letters indicate significant differences among treatments (one-way ANOVA, Tukey’s HSD, P < 0.05). Figure 4. Relative phytohormone profiles in C. quitensis across experimental treatments. (a) Growth-related hormones: indole-3-acetic acid (IAA), trans -zeatin, gibberellin A 4 (GA 4 ), and 24-epibrassinolide. (b) Stress-related hormones: abscisic acid (ABA), phaseic acid (PA), dihydrophaseic acid (DPA), and 1-aminocyclopropane-1-carboxylic acid (ACC). (c) Defense-related hormones: salicylic acid (SA), methyl salicylate (MeSA), jasmonic acid (JA), jasmonoyl-isoleucine (JA-Ile), and methyl jasmonate (MeJA). Bars represent means ± SE (n = 6) expressed relative to hydroponic-acclimated controls (100%). Chemical structures are shown with their corresponding color codes. Letters in the summary table denote significant differences among treatments (one-way ANOVA, Tukey’s HSD, P < 0.05). Figure 5. Integrative metabolomic profiling of C. quitensis under heat stress and volatile priming. (a) GC–MS volatile profiles represented as Z‐score heatmaps for control (CEU) and LT 50 -treated plants, with representative chemical structures shown. Corresponding chromatograms display relative abundances (% base peak) of detected volatiles. (b) LC–MS non-volatile metabolite profiles shown as Z‐score heatmaps for CEU, LT 50 , volatile receiver control (VRC), and volatile receiver primed (VRP) plants, with representative metabolite structures illustrated. These profiles reveal distinct metabolic reprogramming of C. quitensis in response to thermal stress and volatile signaling. Figure 6. Relative expression of stress-related genes in C. quitensis under experimental treatments. Gene expression was quantified by RT–qPCR in control (CEU), LT 50 -treated (3 h at 58.4 °C), volatile receiver control (VRC), and volatile receiver primed (VRP) plants. Values are log₂(2 ⁻ΔΔCT +1) means (n = 6). Color scale indicates relative transcript abundance, with warmer colors representing higher expression and cooler colors lower expression. Figure 7. Proposed model of volatile-mediated thermotolerance priming in C. quitensis . Exposure of emitter plants to heat stress at LT 50 induces ROS overproduction, leading to carotenoid degradation through CCDs and oxidation, and lipid peroxidation via the LOX pathway. These processes generate apocarotenoids (e.g. β-ionone, β-cyclocitral) and oxylipins/green leaf volatiles (GLVs, e.g. jasmonic acid, C6 aldehydes), which are emitted as VOCs. VOC transfer to neighboring receiver plants induces a primed physiological state characterized by enhanced expression of heat shock proteins (HSPs), increased antioxidant activity, higher basal levels of ABA and JA, and elevated flavonoid accumulation. Supplementary Material File (image6.emf) Download 3.89 MB Information & Authors Information Version history V1 Version 1 11 September 2025 Peer review timeline Published Plant, Cell & Environment Version of Record 28 Dec 2025 Published Copyright This work is licensed under a Non Exclusive No Reuse License. Keywords antarctica colobanthus quitensis hormones secondary metabolism thermotolerance volatile organic compounds Authors Affiliations Rodrigo Contreras 0000-0001-9970-3125 [email protected] Universidad de Santiago de Chile Facultad de Quimica y Biologia View all articles by this author Gustavo Zuñiga Universidad de Santiago de Chile Facultad de Quimica y Biologia View all articles by this author Metrics & Citations Metrics Article Usage 173 views 133 downloads .FvxKWukQNSOunydq8rnd { width: 100px; } Citations Download citation Rodrigo Contreras, Gustavo Zuñiga. Intraspecific priming of thermotolerance by heat-induced volatiles in Antarctic Colobanthus quitensis. Authorea . 11 September 2025. DOI: https://doi.org/10.22541/au.175760850.09198149/v1 If you have the appropriate software installed, you can download article citation data to the citation manager of your choice. Simply select your manager software from the list below and click Download. 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