Chickpea (Cicer arietinum L.), a major winter legume in northern South Asia and Australia, frequently encounters low temperatures (0–15 °C) during reproduction, causing substantial yield losses. The present study involved screening two independent sets of 100 genotypes over consecutive winters to identify sources of reproductive-stage cold tolerance and to elucidate the underlying mechanisms. Following outdoor establishment, plants were exposed to controlled cold stress (15/7 °C day/night) during flowering and pod development (15 d) in walk-in growth chambers. Ten representative cold-tolerant (CT) and ten cold-sensitive (CS) genotypes were selected each year based on integrated performance across yield, physiological, biochemical, and reproductive traits for a detailed mechanistic analysis. Cold-sensitive genotypes exhibited severe dysfunction, characterized by high electrolyte leakage (50−59% above CT) and malondialdehyde (39−51% above CT), indicating membrane damage. Reduced chlorophyll content (21−23%), photosystem II efficiency (23−29%), and stomatal conductance (40−43%) impaired photosynthesis. Inadequate cryoprotectants (reduced by 25−58%) and antioxidants (reduced by 38−55%) caused oxidative damage. Reproductive collapse followed, with pollen viability and germination declining by 24−46%, stigma receptivity and ovule viability decreasing by 41−68%, and seed yields falling by 85−95%. Cold-tolerant genotypes-maintained homeostasis through integrated protection in terms of superior membrane stability, enhanced compatible solutes, and elevated antioxidant activities, which sustained photosynthesis and reproductive success, achieving better yields under cold stress. Principal component analysis revealed cold tolerance as an integrated system (PC1:72.6–81.3% variance), clearly separating the CT from the CS genotypes. Membrane stability, photosynthetic efficiency, and pollen viability emerged as diagnostic traits (r = 0.85–0.91 with yield, p < 0.001; heritability 70−99%). Tolerance operated independently of maturity (R² = 0.10–0.18), enabling donor identification across maturity classes. Twenty cold-tolerant genotypes were identified, spanning the early, medium, and late maturity groups, respectively. These findings establish a mechanistic understanding of reproductive-stage cold tolerance, provide vital selection markers, and identify genetic resources for breeding cold-resilient chickpea cultivars.
Funding: This work was supported by the Department of Biotechnology, New Delhi, India (BT/Ag/Network/Chickpea/2019-20 to HN; research fellowships to DP and SK) and Kansas Agricultural Experiment Station, USA (contribution number 26-140-J to PV).
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Citation: Padhiar D, Kaur S, Parida SK, Jha UC, Shama KD, Prasad PVV, et al. (2026) Genotypic variability and trait associations for cold stress tolerance in cultivated chickpea (Cicer arietinum L.) during the reproductive stage. PLoS One 21(2): e0343120. https://doi.org/10.1371/journal.pone.0343120
The specific objectives were as follows: (i) to identify cold-tolerant and cold-sensitive genotypes based on integrated mechanisms across two independent screening experiments; (ii) to characterize the physiological, biochemical, and reproductive trait profiles distinguishing tolerant from sensitive genotypes under reproductive-stage cold stress; (iii) to determine the traits and trait combinations most strongly associated with yield maintenance under cold stress using correlation and multivariate analyses; and (iv) to establish practical, trait-based selection criteria and identify donor genotypes spanning different maturity classes for breeding programs targeting cold-prone production environments.
This study addresses these critical gaps through a systematic multi-trait evaluation of cold tolerance during the reproductive phase. We screened two independent sets of 100 chickpea genotypes (200 unique genotypes in total) over two consecutive winter seasons, combining initial field evaluation with controlled cold stress imposed during flowering and early pod development in walk-in growth chambers. This experimental design enabled the assessment of yield performance under both optimal and cold stress conditions, while simultaneously measuring a comprehensive suite of physiological, biochemical, and reproductive traits under standardized stress conditions.
Previous research has made important contributions to understanding cold tolerance in chickpeas. Early studies identified genetic variations in germplasm collections and wild relatives [ 17 , 18 ], while subsequent studies characterized vegetative-stage responses under controlled conditions [ 16 , 19 ]. Recent investigations have begun to elucidate reproductive-stage responses, demonstrating that cold-tolerant genotypes maintain reproductive organ function through enhanced antioxidant activity and cryoprotectant accumulation [ 6 , 20 ]. However, there are still significant knowledge gaps. Most previous studies have focused on the seedling or vegetative stages, examined small numbers of genotypes (typically <20), or evaluated limited trait sets without integrating physiological, biochemical, and reproductive responses into comprehensive tolerance profiles. Reproductive-stage cold tolerance, a critical determinant of yield stability, remains comparatively underexplored relative to its agronomic importance, and few studies have combined field and controlled environment evaluations to identify potential donor lines suitable for breeding deployment. Furthermore, the relative importance of different tolerance mechanisms, their hierarchical organization, and trait combinations that reliably predict field performance under reproductive-stage cold stress remain poorly understood.
The physiological and biochemical basis of cold tolerance in chickpea is complex and multifaceted. At the cellular level, tolerance depends on maintaining membrane integrity and fluidity, as membrane destabilization disrupts compartmentation and metabolic function [ 9 , 10 ]. Photosynthetic stability, sustaining chlorophyll content, photosystem II efficiency, and stomatal conductance are essential for continued carbon assimilation and energy supply during stress [ 11 ]. Cold stress simultaneously induces osmotic and oxidative stress, requiring the coordinated accumulation of compatible solutes that provide osmotic adjustment, membrane stabilization, and contribute to reactive oxygen species (ROS) scavenging [ 12 , 13 ]. Activation of antioxidant defence systems, including enzymatic components and non-enzymatic antioxidants, mitigates oxidative damage reflected in reduced malondialdehyde (MDA) accumulation and lipid peroxidation [ 14 , 15 ]. These interconnected mechanisms ultimately determine whether reproductive processes can proceed normally, making them critical determinants of yield during cold stress [ 16 ].
Originating from the warm Mediterranean region, chickpea exhibits inherent sensitivity to low temperatures (<20/10 °C day/night), with critical thresholds between 4–6 °C triggering physiological dysfunction [ 4 , 5 ]. Depending on the sowing time and location, crops may encounter chilling stress during vegetative development or, more critically, during the reproductive phase when sensitivity is most pronounced [ 6 , 7 ]. Cold stress during reproduction is particularly harmful. Temperatures below 15 °C during flowering and early pod development disrupt microsporogenesis, impair pollen viability and germination, reduce stigma receptivity and ovule fertility, and trigger widespread flower and pod abortion, collectively resulting in yield losses exceeding 70% in susceptible genotypes [ 5 , 6 , 8 ]. This reproductive vulnerability represents a major constraint on chickpea productivity across South Asia and cool-season Mediterranean environments.
Chickpea (Cicer arietinum L.) is a globally important legume crop valued for its high protein content, essential micronutrients, and biological nitrogen fixation [ 1 , 2 ]. In 2023, the global production of chickpea reached approximately 16.5 million tons, with India contributing nearly 75% [ 3 ]. Despite its importance in food security across South Asia, the Middle East, and the Mediterranean, chickpea production faces persistent challenges from abiotic stresses, particularly cold, heat, and drought, which limit the stability of the yield and restrict its expansion into climatically marginal areas.
2. Materials and methods
2.1. Experimental setup and growth conditions Two independent sets of 100 chickpea genotypes (S1 and S2 Tables in S1 File) were obtained from the International Crops Research Institute for the Semi-Arid Tropics in Hyderabad, India. These two independent sets of 100 genotypes each year (totalling 200 unique genotypes) were screened for cold tolerance using multiple traits. Subsequently, a few contrasting genotypes (cold-tolerant and cold-sensitive) were identified from these two datasets based on their growth, physiological, biochemical, yield, and stability characteristics across two years. Each set, consisting of distinct genotypes, corresponded to a specific study year and was independently assessed over two consecutive years. Field experiments were conducted during the ‘rabi’ (winter) seasons of 2021–22 and 2022–23 at the Department of Botany, Panjab University, Chandigarh, India (30.75° N, 76.78° E). Sowing was carried out on November 1st in both years, a period characterised by gradually declining night temperatures that typically impose cold stress during the reproductive stage. Prior to sowing, seeds were inoculated with Mesorhizobium ciceri (1.95 g kg ⁻ ¹ seed). Plants were grown in 4 kg pots containing a 3:1 sandy loam–to–sand mixture, supplemented with farmyard manure at a 3:1 soil-to-manure ratio. Five seeds were initially sown per pot and subsequently thinned to three uniform seedlings after their emergence. The experiments were conducted using a randomized complete block design (RCBD) with three replicates. Each replicate comprised three pots per genotype and three plants per pot. Plants were maintained at the Department of Botany, Panjab University, Chandigarh, India, and assessed under field and controlled-environment conditions (S3 Table in S1 File). Irrigation was provided when needed. Regular temperature measurements (both maximum and minimum) and relative humidity were recorded from the sowing date to the flowering stage of the plants. To evaluate the effects of cold stress, plants were initially raised under outdoor conditions with a light intensity of 1,300–1,500 µmol m ⁻ ² s ⁻ ¹ and 60–70% relative humidity, where average maximum temperatures gradually declined from 30–32 °C at sowing to approximately 15–18 °C, and minimum temperatures decreased from 14–16 °C to nearly 5–7 °C until the completion of the vegetative stage (S1a Fig in S1 File). The average temperature followed the same trend, indicating that the plants experienced a slight natural decrease in temperature without an abrupt cold exposure. At 78 days after sowing in the 1st and 2nd years, when all genotypes had initiated flowering, the plants were transferred to walk-in growth chambers for temperature treatments (days to flowering: S4 and S5 Tables in S1 File). The temperature regime in the growth chamber during the flowering-to-maturity phase followed a clearly defined sequence. During the acclimation phase, temperatures were gradually lowered (1 °C per day) from a 12 h day/12 h night regime of 25/15 °C to 15/7 °C, with a 12-h photoperiod (600 µmol m ⁻ ² s ⁻ ¹) and 65–70% relative humidity to impose controlled reproductive-stage cold stress. Cold stress (15/7 °C) was maintained for 15 d, whereas the control set of plants was grown at 25/15 °C for 15 d under similar light and RH conditions. Following stress, recovery was initiated by increasing the temperature by 2 °C per day until it reached 30/25 °C (day/night, 12-h light/12-h dark) in both the control and cold-stressed plants. The plants were then allowed to fully mature at this temperature. This contrast highlights that outdoor conditions imposed gradual cooling, whereas growth chamber treatments ensured precise cold stress at critical stages of plant development.
2.2. Trait measurement Each year, 100 distinct genotypes were evaluated. Growth and yield traits were assessed at physiological maturity under both control and cold stress conditions, whereas physiological, biochemical, and reproductive traits were measured immediately after the 15-day cold stress treatment (on the 16th day in the control and cold-stressed environments). The stress duration was kept constant for all genotypes by initiating cold exposure at the same reproductive stage and maintaining identical chamber conditions for each set. In total, 31 traits were measured and grouped into six categories: Growth traits: plant height, biomass, harvest index Phenological traits included days to flowering (DTF), days to podding (DTP), and days to maturity (DTM). Yield traits: pod number, seed number, total seed weight per plant (seed yield), and 10-seed weight (seed size). Reproductive traits included pollen viability, pollen germination, stigma receptivity, and ovule viability. Physiological traits included membrane stability (electrolyte leakage), chlorophyll fluorescence (Fv/Fm), chlorophyll content (SPAD), relative leaf water content, stomatal conductance, and nodulation ability. Biochemical traits included malondialdehyde (MDA), proline, trehalose, total soluble sugars, and antioxidant enzymes (superoxide dismutase (SOD), catalase (CAT), ascorbate peroxidase (APX), glutathione reductase (GR), ascorbic acid (AsA), and reduced glutathione (GSH).
2.3. Predicted yield (Ŷ s ) and cold tolerance indexes The relationship between seed yield under cold stress (Ys) and seed yield under control conditions (Yc) was analysed using linear regression for each year separately. The predicted seed yield under cold stress (Ŷs) was estimated from the regression equation derived between Yc and Ys. The slope (a) and intercept (b) of the regression were 0.94 and 2.35 in the first year, and 0.40 and 0.52 in the second year, respectively. Cold-tolerance residuals (R) were calculated as the difference between observed and predicted seed yield under cold stress (R = Ys − Ŷs), following the framework described by [21]. These residuals represent yield variation under cold stress that is independent of yield potential under control conditions. After calculating the residuals, they were used as dependent variables in regression analyses against multiple explanatory traits to identify parameters associated with cold tolerance or traits that are easier to assess. (i) ratio of seed yield under cold to control conditions (Ys/Yc); (ii) ratio of total biomass; (iii) ratio of seed number per plant; (iv) ratio of 10-seed weight (seed size); (v) ratio of harvest index; (vi) ratio of plant height; (vii) days to flowering; (viii) ratio of days to podding; (ix) ratio of days to maturity; and (x) ratio of physiological, biochemical, and reproductive traits. Polynomial (Type II) regression models were applied where appropriate.
2.4. Physiological traits Physiological traits were measured according to previously established protocols [6,22,23]. Membrane injury was quantified via electrolyte leakage (EL) by measuring the conductivity before and after heat treatment [22]. Lipid peroxidation was estimated as malondialdehyde (MDA) content using the thiobarbituric acid method, with absorbance measured at 532 nm, and the concentration calculated using an extinction coefficient of 155 mM ⁻ ¹ cm ⁻ ¹ [24]. The relative leaf water content (RLWC) was calculated from the fresh, turgid, and dry weights [25]. Stomatal conductance (gS) was measured using an SC-1 portable leaf porometer (Decagon Devices, Pullman, WA, USA) on leaves from the second or third node below the apex at 11:00 a.m. [23]. The SPAD chlorophyll index was estimated using a SPAD chlorophyll meter (Apogee Instruments, Logan, UT, USA) on marked leaves between 10:00 and 11:00 h [23]. Chlorophyll fluorescence (ChlF) was measured at 11:00 h using dark-adapted measurements with a modulated fluorometer (Model OS1-FL, Opti-Sciences, Tyngsboro, MA, USA) [23]. Carotenoids were extracted using 80% acetone and quantified spectrophotometrically at 440, 645, and 663 nm [26]. Nodulation ability was assessed by manual counting of nodules per plant [27].
2.5. Reproductive traits Reproductive traits were assessed using previously established methods [6,20,22]. All reproductive samples were collected from plants grown under controlled chamber conditions one day before anthesis (stigma and ovule) or at anthesis (pollen). Pollen viability was determined using 0.5% acetocarmine staining, with approximately 200 grains observed per sample, and viable grains were identified by their uniform size, shape, and strong red staining [22]. Pollen germination was assessed in vitro on nutrient medium (10% sucrose, 1,640 mM boric acid, 990 mM potassium nitrate, 812 mM magnesium sulfate, 1,269 mM calcium nitrate, pH 6.5), and the germination percentage was calculated as the proportion of grains with elongated pollen tubes [22,28]. Stigma receptivity was evaluated using an esterase-mediated chromogenic assay with α-naphthaleneacetic acid and fast blue B salt, scored on a 1–5 scale based on staining intensity [29]. Ovule viability was assessed using the triphenyl tetrazolium chloride (TTC) reduction assay, with viability scored on a 1–5 scale based on red pigmentation intensity [22].
2.6. Antioxidants 2.6.1. Enzymatic antioxidants. Antioxidant enzyme activity was determined using standard protocols [6,20,30–32]. Fresh tissue (~500 mg) was homogenized in ice-cold 50 mM phosphate buffer (pH 7.0–7.8) and centrifuged at 3,360–15,000 g for 5–15 min at 4 °C. The supernatants were used as enzyme extracts in all assays. Superoxide dismutase (SOD) activity was assayed by monitoring the inhibition of nitro blue tetrazolium (NBT) photoreduction [33]. The reaction mixture contained enzyme extract, phosphate buffer (pH 7.8), methionine, NBT, EDTA, and riboflavin, and the absorbance was measured at 560 nm after 10 min of light exposure. The activity was expressed as units per mg of protein. Catalase (CAT) activity was determined by monitoring the decline in absorbance at 410 nm due to H₂O₂ decomposition [30]. The activity was calculated using an extinction coefficient of 40 mM ⁻ ¹ cm ⁻ ¹ and expressed as mmol H₂O₂ decomposed per mg of protein. Ascorbate peroxidase (APX) activity was measured by monitoring ascorbate oxidation at 290 nm [31]. Activity was calculated using an extinction coefficient of 2.8 mM ⁻ ¹ cm ⁻ ¹ and expressed as mmol of ascorbate oxidized per min per mg of protein. Glutathione reductase (GR) activity was determined by following NADPH oxidation at 340 nm [32]. The activity was expressed as mmol GSSG reduced per min per mg of protein. 2.6.2. Non-enzymatic antioxidants. Ascorbate (AsA) content was determined following extraction in 6% TCA and reaction with 2% dinitrophenylhydrazine (DNPH) and thiourea, with absorbance measured at 530 nm after heating and sulfuric acid addition [34]. Reduced glutathione (GSH) was estimated using the DTNB method, with tissue homogenized in metaphosphoric acid and absorbance measured at 412 nm [35]. Both were quantified against standard curves and expressed as mg g ⁻ ¹ DW (AsA) or nmol g ⁻ ¹ DW (GSH).
2.7. Osmolytes The osmolyte content was determined using standard colourimetric methods [6,20,36–39]. Proline was extracted using 3% sulfosalicylic acid, reacted with acidified ninhydrin, and the chromophore was extracted into toluene. The absorbance was measured at 520 nm, and the proline content was calculated using a standard curve [36]. Trehalose was extracted using 80% hot ethanol and quantified using the anthrone-TCA method [37,38]. Total soluble sugars were extracted in 80% ethanol through repeated extractions, reacted with anthrone reagent, and quantified at 625 nm using a glucose standard curve [39]. All osmolyte concentrations were expressed as nmol g ⁻ ¹ DW (proline and trehalose) or mg g ⁻ ¹ FW (total sugars).