Impact of Arbuscular Mycorrhizal Symbiosis on Photosynthetic, Antioxidant Enzyme, and Water Flux Parameters in Salt-Stressed Chickpea (Cicer arietinum) Plants

Abstract

Globally, escalating soil salinization poses significant abiotic stress, disproportionately impacting crops like chickpea (Cicer arientinum L.). This legume exhibits high sensitivity to salinity, which disrupts various physiological and metabolic processes, ultimately hindering growth and productivity. AMF (arbuscular mycorrhizal fungus) reduces salt’s detrimental effects on plants’ growth by bolstering the plant’s antioxidant defense system, effectively reducing the damage caused by oxidative stress. In this study, the impact of AMF on salinity stress alleviation in chickpea was investigated in pot-grown experiments. Rhizophagus fasciculatus was used to inoculate the seeds of three different chickpea varieties (HC-3, CSG-8962, and C-235), and the physiological and biochemical changes of the AMF-inoculated and non-inoculated chickpea plants were studied. When exposed to salinity stress, the plants exhibited decreased leaf relative water content (RWC %) (21.13–31.30%), increased leaf relative stress injury, decreased chlorophyll content (45.22–58.24%), photochemical quantum yield, photosynthetic rate, transpiration rate, and stomatal conductance as compared to the control plants, but opposite results were observed in AMF colonized plants. A 9.16% to 14.79% increase in chlorophyll content was reported after AMF colonization. The activities of antioxidant enzymes such as superoxide dismutase (SOD), catalase (CAT), and peroxidase (POX) were increased by salt stress. They were further enhanced by AMF inoculation SOD activity by 20.3% to 23.3%, CAT activity by 65.7% to 78.7%, and POX activity by 32.7% to 39.3%. The findings clearly show that AMF Rhizophagus fasciculatus, via enhancing RWC, photosynthetic parameters, and antioxidant enzymes, can mitigate salinity stress in chickpeas.

1. Introduction

Salinity stress, characterized by an excessive accumulation of salts in the soil, ranks among the most significant abiotic constraints to agricultural productivity worldwide. Recent climatic shifts, marked by declining precipitation, rising temperatures, and increased evapotranspiration, exacerbate this challenge. The risk of salty soil and water is increased in arid and semi-arid regions of the world due to inadequate rainfall, drainage, and management practices [1,2]. A staggering 1.125 billion hectares of land worldwide, roughly the size of Europe, are affected by salinity, with human activity contributing to the degradation of 76 million hectares [3]. Crop production and agricultural operations are highly vulnerable due to this extreme salinization. Many legumes, including faba beans, common beans, lentils, soybeans, mungbeans, and chickpeas, are less productive and grow more slowly in salinity [4]. Salinity inhibits the processes of growth and development of a plant due to reduced relative water content (RWC), chlorophyll content, photochemical quantum yield (PQY), photosynthetic rate (PR), transpiration rate (TR), and stomatal conductance (SC), and also due to cytotoxicity caused by excessive production of reactive oxygen species (ROS) [5,6,7,8]. Increased relative stress injury in chickpea plants was reported by [9] under salt stress conditions. However, plants respond to salt differently and use different adaptive strategies in other situations [10]. Plants accelerate the production of their enzyme antioxidants, such as SOD, CAT, and POX, to lessen the excessive generation of ROS in salty environments [11] or increase the uptake of potassium to prevent excessive water loss and protect the internal structure [12].

Plants exposed to salt can improve their tolerance through mutualistic interactions with rhizosphere microorganisms, in addition to using some internal processes of adaptation [13]. Most terrestrial plants have symbiotic associations with soil microbes, including AMF [14]. According to estimates, AMF in soil is linked to 70–90% of terrestrial plant species [15]. AMF has been demonstrated to improve water uptake and chlorophyll production and increase the enzyme activity of protective systems in host plants, hence boosting salt tolerance [16,17]. It can also compensate for photosynthetically fixed carbon by absorbing water and nutrients from the soil and transferring them to their host plants [18], thus improving photosynthesis and water use efficiency [19]. Arbuscular mycorrhizal (AM) symbiosis in plants boosts the activity of antioxidant enzymes to combat the reactive oxygen species produced by salt [20,21]. This makes the relationship important for agriculture and the environment [22]. The physiological and molecular basis of the AMF action on legumes under salt stress is still uncertain.

Chickpea (Cicer arietinum L.), a self-pollinating legume, belongs to the Fabaceae family. It is a diploid plant with eight pairs of chromosomes [23]. It’s considered among the earliest pulses grown and known to exist from prehistoric times in Asia and Europe. It is more commonly grown in arid and semi-arid regions [24]. India produces most of the world’s chickpeas (9.075 million tonnes), making it the biggest producer in the world. Approximately 50% of India’s pulse production comes from chickpea alone. States like Maharashtra (25.97%), Madhya Pradesh (18.59%), Rajasthan (20.65%), Gujarat (10.10%), Uttar Pradesh (5.64%), and Haryana are among the leading producers of chickpeas in India. It is utilized for human consumption and fed to livestock. Chickpea contains 21.1 percent protein, 61.5 percent carbohydrates, and 4.5 percent fat. It possesses high amounts of calcium, iron, niacin, and other mineral nutrients [25]. Despite their nutritional importance, chickpeas face a formidable foe: salinity. This abiotic stress disrupts their growth and productivity, highlighting the need to explore their adaptation mechanisms in salty environments.

Thus, the present investigation was designed to (a) examine the impact of AMF symbiosis on the photosynthetic attributes of chickpeas under conditions of salt stress and (b) explore the effects of AMF on antioxidant enzymes under conditions of salt stress. With the above objectives, this research will clarify the process by which AMF enhances chickpea growth and determine the potential of AMF (Rhizophagus fasciculatus) in the production of chickpeas in saline conditions.

2. Materials and Methods

Three chickpea (Cicer arietinum L.) varieties, HC-3, CSG-8962, and C-235, were grown in dune sand-filled pots [saturation percentage 39%, pH 8.075 ± 0.005, EC 0.13 ± 0.01 dSm⁻¹, Nitrogen 10.4 ± 0.1 mg kg⁻¹ (Kjeldahl 1883), Phosphorus 2.55 ± 0.05 mg kg⁻¹ (Olsen 1954), Potassium 162.5 ± 0.5 mg kg⁻¹] in a screen house of the Department of Botany and Plant Physiology, CCS HAU, Hisar-125 004, India. HC-3 exhibits moderate tolerance to salinity, CSG-8962 is known for its resilience to salt conditions, and C-235 is recognized for its sensitivity to abiotic stresses. These contrasting traits make them suitable candidates for comparative analysis under salinity stress. Rhizophagus fasciculatus obtained from the Plant Pathology Department, CCS HAU, Hisar, was used for mycorrhizal colonization. Rhizophagus fasciculatus was chosen because it has demonstrated potential to improve nutrient absorption and oxidative stress resilience, thereby enhancing the salt tolerance in legumes. Before sowing, four chloride-dominant salinity levels (control, 2.0, 3.0, and 4.0 dS m⁻¹) were established to simulate varying salt stress conditions. A combination of several salts, including NaCl, MgCl₂, MgSO₄, and CaCl₂, was used to generate the chloride (Cl⁻) dominated saltwater. The ratios of Na⁺:Ca²⁺ + Mg²⁺ and Ca²⁺:Mg²⁺ were 1:1 and 1:3, respectively, while the Cl⁻:SO₄²⁻ was 7:3 on a meq basis. Seeds were germinated in saline conditions by introducing salinity treatments before sowing mimic-existing salinity stress. Tap water was used for irrigation, and soil moisture was consistently maintained at 70% field capacity. Gravimetric methods were employed to monitor and adjust soil moisture. To promote root colonization, the inoculum was introduced to the pot 3–4 cm deep from the soil surface before the sowing of seeds. Sporocarps were isolated from the roots of bajra plants cultivated under sterile conditions. The sporocarps were then quantified microscopically, and 450–500 sporocarps were evenly added to each pot to ensure they were adequately inoculated. An identical amount of autoclaved inoculum was added to pots that were not AM-treated to create the same soil texture. There were nine pots for each treatment. Nitrogen-free nutrient solution was provided to the crop every 15 days. Data was collected 80 days after the sowing date (DAS). Plants were at the flowering stage (80 DAS), a critical phase for evaluating the physiological and biochemical impacts of salinity and mycorrhizal inoculation. The experiment was a factorial experiment based on a completely randomized design.

2.1. Measurements

2.1.1. Mycorrhizal Status

The study analyzed AMF colonization in Cicer arietinum roots, revealing variable fungal structures: spores, arbuscules for nutrient exchange, hyphae for resource transport, and vesicles for storage. Colonization ranged from dense networks with abundant structures to sparse patterns with isolated hyphae. The quantitative assessment showed differences in colonization efficiency among plant varieties. Microscopy images (Figure 1A–F) confirmed the presence of these structures. There were differences in the AMF colonization patterns found in Cicer arietinum roots, highlighting AMF’s role in enhancing nutrient uptake and stress resilience under salinity.

2.1.2. Relative Water Content

Leaf RWC was determined following the methodology of [26]. Freshly collected leaf samples were weighed to obtain their initial weight. Each leaf was then immersed in a separate petri dish filled with distilled water for complete hydration. After retrieval, excess water was gently blotted from the leaves, and their turgid weight was recorded. Finally, the samples were dried in an oven at 85 °C for 72 h and weighed again to achieve dry weight. RWC percentage was subsequently anticipated using the provided formula:

RWC (%) = (Fresh weight − Dry weight/Turgid weight − Dry weight) × 100

2.1.3. Relative Stress Injury

Relative stress injury was estimated using the method of [27]. Leaf samples (200 mg) were incubated in 10 mL of distilled water at room temperature for 5 h in 20 mL test tubes. The solution’s initial electrical conductivity (EC1) was measured using an EC meter. Subsequently, the samples were subjected to heat stress by boiling in a water bath for 50 min, inducing complete tissue disruption. After cooling, the final EC (EC2) was measured. RSI was then calculated using the following formula:

RSI (%) = (EC1/EC2) × 100

2.1.4. Total Chlorophyll Content

Following the protocol of [28], leaves were gently washed and blotted dry before being cut into 200 mg discs. Each disc was then submerged in a test tube containing 5 mL of dimethyl sulfoxide (DMSO) for overnight incubation. Subsequently, the absorbance of the extracted chlorophyll in the DMSO solution was recorded at 663 nm and 645 nm using a spectrophotometer. Chlorophyll content was then calculated in mg g⁻¹ FW using the formula provided by [29]:

Chl “a” = 12.3 A 663 − 0.86 A 645 × V/a × W

Chl “b” = 19.3 A 645 − 3.6 A 663 × V/a × W

Total chlorophyll = Chl ‘a’ + Chl ‘b’

Here, V = Volume of DMSO, a = Path length, W = Tissue weight taken (mg).

2.1.5. Photochemical Quantum Yield (Fv/Fm)

Chlorophyll fluorescence measurements were conducted on intact plants at midday using a chlorophyll fluorometer (OS-30p, Opti-Science, Inc., Hudson, NY, USA). A fully expanded leaf was attached to a clip and acclimated to darkness for at least two minutes. The dark-adapted leaf was then illuminated for one second by a constant light pulse (1500 mol m⁻² s⁻¹) emitted by three LEDs within the sensor. Variable fluorescence (Fv) was calculated as the difference between the initial fluorescence (F0) and the maximum fluorescence (Fm). Subsequently, the PQY was determined as the Fv/Fm ratio.

2.1.6. Photosynthetic Rate, Transpiration Rate, and Stomatal Conductance

Photosynthetic rate (µmole CO₂ m⁻² s⁻¹), transpiration rate (mmole H₂O m⁻² s⁻¹), and stomatal conductance (mmole H₂O m⁻² s⁻¹) were determined on the third fully expanded leaf from the top of plants from three pots using a portable Infra Red Gas Analyzer (IRGA, LCi-SD, ADC Bioscience, Hoddesdon, UK). Only clean, dry leaves devoid of visible damage or disease were selected. The leaf chamber positioned the leaf with its photosystem, receiving direct sunlight, and data was recorded upon value stabilization. The system automatically calculated the photosynthetic rate based on pre-loaded flow and leaf area data. All measurements were performed between 11:00 AM and 12:30 PM.

2.1.7. Antioxidant Enzyme Activity

Preparation of Enzyme Extract: Five hundred milligrams of leaf samples were thoroughly rinsed with ice-cold distilled water. Each sample was then individually homogenized in 5 mL of pre-chilled extraction buffer using a mortar and pestle. The extraction buffer consisted of 0.1 M phosphate solution adjusted to a pH of 7. Following homogenization, the extracts were centrifuged at 15,000 rpm for 20 min at 4 °C. The resulting supernatants were then used to examine the following enzymes.

Superoxide Dismutase (SOD): SOD activity was determined with a slight modification of the method described by [30]. Each reaction mixture contained 1 mL of enzyme extract, 0.5 mL each of methionine, nitroblue tetrazolium (NBT), EDTA, Na₂CO₃ solution, and buffer to reach a final volume of 4 mL. The pH of the mixture was adjusted to 10.2, and 0.5 mL of riboflavin was added after buffer adjustment. Reaction tubes were shaken and positioned 30 cm away from a light source (8 × 20 W fluorescent lamp). After 10 min of incubation, the light source was turned off, and a black cloth was quickly placed over the tubes. Absorbance at 560 nm was measured using a spectrophotometer. A non-irradiated control mixture, incapable of developing color, was included. The SOD present in the sample stopped the photochemical reaction, and the degree of inhibition was used to estimate enzyme activity. A plot of log A₅₆₀ versus enzyme extract volume was generated. One unit of SOD activity was defined as the enzyme extract amount required for 50% inhibition of the photochemical reaction and was expressed as mg⁻¹ protein min⁻¹.

Catalase (CAT): CAT activity was determined following the method of [31]. The reaction mixture consisted of 500 μL of enzyme extract, 0.2 mL of 0.1 M hydrogen peroxide (H₂O₂), and 1.5 mL of 50 mM potassium phosphate buffer. Absorbance measurements were initiated upon adding the enzyme sample, and the mixture was incubated for 3 min. The change in absorbance at 240 nm was then recorded at 15-s intervals for 1.5 min. The specific activity of the enzyme was estimated in units of mg⁻¹ protein min⁻¹ based on the initial rate of H₂O₂ decomposition.

Peroxidase (POX): POX activity was determined based on the method described by [32]. Each reaction mixture (3 mL) contained 0.1 M phosphate buffer (pH 7.0), 0.1 mM guaiacol, and cell-free extract (100 µL). The reaction was started by adding 0.1 mM hydrogen peroxide (H₂O₂), and the subsequent increase in absorbance at 470 nm was monitored for 2 min. The 26.6 mM⁻¹ cm⁻¹ extinction coefficient for guaiacol was employed to calculate the activity.

2.2. Statistical Analysis

Data were subjected to ANOVA using OPSTAT, CCS HAU, Hisar, Haryana, India to determine the statistical significance of observed differences. Tukey’s range test was performed to compare means at p < 0.05. A complete randomized design (CRD) was used to analyze data statistically, and each observation was replicated thrice. Origin Pro (2018) was used to make PCA (Principal component analysis) biplot.

3. Results

3.1. Relative Water Content

With increasing salinity, the RWC content was significantly reduced. Maximum reduction was noted in C-235 (31.30%), whereas HC-3 (21.13%) and CSG-8962 (22.07%) showed a lesser decline at 4 dSm⁻¹. There was a noticeable increment upon colonization with Rhizophagus fasciculatus. RWC was increased by 5.95% in variety HC-3, 5.01% in CSG-8962, and 16.11% in C-235 at 4 dSm⁻¹ (Figure 2).

3.2. Relative Stress Injury

All the studied varieties experienced an increment in relative stress injury. A more significant increase was recorded in C-235 (37.03%) than in HC-3 (36.76%) and CSG-8962 (34.2%) at 4 dSm⁻¹ as compared to their respective control. The mycorrhizal symbiosis led to a slight decrease in RSI in variety HC-3 (12.00%), CSG-8962 (7.48%), and C-235 (12.70%) at 4 dSm⁻¹. Varieties with less RSI % showed better tolerance to high salinity (Figure 3).

3.3. Total Chlorophyll Content

With increasing salinity, the total chlorophyll content was significantly reduced. Maximum reduction was recorded in C-235 (58.24%), whereas HC-3 (45.22%) and CSG-8962 (45.65%) showed a lesser decline at 4 dSm⁻¹ as compared to control. There was a noticeable increment upon colonization with Rhizophagus fasciculatus. An increment of 10.25% in variety HC-3, 14.79% in CSG-8962, and 9.16% in C-235 at 4 dSm⁻¹ was noted (Figure 4).

3.4. Photochemical Quantum Yield

In general, photochemical quantum yield at 80 DAS decreased with the increasing salinity levels. Values decreased from control to 4 dSm⁻¹, i.e., 0.796 to 0.636 in HC-3, 0.782 to 0.620 in CSG-8962, and 0.771 to 0.620 in C-235. Application of mycorrhiza resulted in a progressive increase in photochemical quantum yield at each salt level and in control in all three varieties (Figure 5). However, the interaction among all parameters was not statistically significant.

3.5. Photosynthetic Rate, Transpiration Rate, and Stomatal Conductance

The photosynthetic rate decreased significantly from control to 4 dSm⁻¹ in all three varieties. A decrease from 20.72 to 10.18, 18.35 to 6.50, and 17.56 to 5.49 was observed in varieties HC-3, CSG-8962, and C-235, respectively, from control to 4 dSm⁻¹ at 80 DAS. Mycorrhizal inoculation increased the photosynthetic rate in all three varieties. HC-3 showed maximum photosynthetic rate (Figure 6). In addition to the reduction in photosynthetic rate, salt treatment significantly decreased the transpiration rate (TR) and stomatal conductance (SC) parameters (Figure 7 and Figure 8). A decline from 4.05 to 2.84 in HC-3, 3.84 to 2.52 in CSG-8962, and 3.36 to 2.25 in C-235 was recorded in TR. Similarly, a decrease from 0.62 to 0.36, 0.51 to 0.26, and 0.46 to 0.23 was observed in varieties HC-3, CSG-8962, and C-235, respectively, from control to 4 dSm⁻¹ in SC at 80 DAS. Mycorrhiza significantly improved these parameters in all three varieties.

3.6. Superoxide Dismutase (SOD)

Under salt stress conditions, the specific activity of SOD was observed to increase in all the varieties. The specific activity of SOD increased by 56.6%, 40.8%, and 29.1% in varieties HC-3, CSG-8962, and C-235, respectively (Figure 9). The activity of SOD was higher in plants colonized with mycorrhiza in control as well as salt-stressed plants. The activity increased by 23.3% in HC-3, 22.7% in CSG-8962, and 20.3% in C-235 at 4 dSm⁻¹ at 80 DAS after mycorrhizal colonization.

3.7. Catalase

The specific activity of CAT was increased with increasing salinity levels. An increase of 92.3% in HC-3, 92.1% in CSG-8962, and 91.5% in C-235 was noted at 80 DAS (Figure 10) from control to 4 dSm⁻¹. Mycorrhizal colonization further increased specific activity by 78.7%, 67.8%, and 65.7% at 4 dSm⁻¹ in varieties HC-3, CSG-8962, and C-235, respectively.

3.8. Peroxidase

An increase in specific POX activity was observed with increasing salinity levels. The particular activity of POX increased by 88.0% in HC-3, 86.0% in CSG-8962, and 81.6% in C-235 at 80 DAS (Figure 11) from control to 4 dSm⁻¹. After mycorrhizal colonization, specific activity increased by 39.3%, 33.9%, and 32.7% at 4 dSm⁻¹ in varieties HC-3, CSG-8962, and C-235, respectively. The maximum significant increment was recorded in HC-3, both with and without mycorrhiza.

3.9. Principal Component Analysis

The Principal Component Analysis (PCA) biplot provides insights into the performance and responses of treatments (T1–T8) and genotypes (HC-3, CSG-8962, and C-235) under different conditions, such as unstressed and salt-stressed environments (Figure 12). Treatments are as follows: T1 (control), T2 (2 dSm⁻¹), T3 (3 dSm⁻¹), T4 (4 dSm⁻¹), T5 (control + AM), T6 (2 dSm⁻¹ + AM), T7 (3 dSm⁻¹ + AM), T8 (4 dSm⁻¹ + AM).

3.9.1. Division of the Biplot by PC1

The lower right quadrant of the biplot, associated with PC1, contains treatments under unstressed conditions/control (T1). This group is characterized by relatively higher values of parameters such as Relative Water Content (RWC), Chlorophyll (CHL), Photosynthetic Quantum Yield (PQY), Photosynthetic Rate (PR), Transpiration Rate (TR), and Stomatal Conductance (SC). These parameters indicate optimal physiological functioning under favorable conditions. Each genotype (HC-3, CSG-8962, and C-235) shows distinct clustering under T1, reflecting inherent variability in their performance even in unstressed environments.

The lower left quadrant of the biplot corresponds to salinity treatments of T3 and T4, which represent stressed conditions. The reduced RWC, CHL, PQY, PR, TR, and SC values indicate significant stress-induced physiological impacts.

The upper right quadrant of the biplot includes treatments T5, T6, and T7, which demonstrate tolerance. The HC-3 genotype in this region displays significantly higher levels of RWC, CHL, PQY, PR, TR, and SC, distinguishing it as more tolerant to salt stress than other genotypes. This positioning highlights the adaptability of HC-3 under these conditions, which may be due to superior physiological or stress-adaptive mechanisms.

3.9.2. Divison of Biplot by PC2

Under salt stress treatments, PC2 separates the three genotypes (HC-3, CSG-8962, and C-235). The three genotypes occupy the upper left quadrant region under severe salt stress treatments T7 and T8. Elevated levels of antioxidant enzymes, such as SOD, CAT, and POX, characterize these treatments. The increased activity of these enzymes suggests an enhanced oxidative stress response, a critical mechanism for coping with salt-induced reactive oxygen species (ROS).

The clustering of CSG-8962 and C-235 under treatments T3 and T4 in the lower left quadrant region reflects higher RSI, indicating their vulnerability to salt stress. This susceptibility is likely associated with reduced antioxidant activity and physiological parameters, making these genotypes less efficient in mitigating salt-induced damage.

4. Discussion

High salt levels in the soil create a double problem for plants. Firstly, it acts like a drought, making it difficult for them to absorb water. Secondly, it disrupts the uptake of essential nutrients. This combination of water shortage and nutrient deficiency weakens plants and reduces their growth and productivity. Mycorrhizal inoculation significantly improved plant height, biomass, and pod production across all varieties, underscoring its protective effects against salinity-induced stress. This arbuscular mycorrhizal association also helps plants mitigate salinity stress by improving water status, photosynthesis, and antioxidant production to scavenge ROS. In plants, RWC is a crucial indicator of their hydration status. It reflects the balance between water uptake and water loss through transpiration. Relative water content decreased in plants that were subjected to salt stress. Numerous investigations verified a reduced RWC in plants under salt stress [33,34,35]. The current study’s reduced relative water content of leaves may have resulted from their decreased absorption of water at increased levels of salt [36]. It might also be due to the reduced root hydraulic conductivity. However, mycorrhiza inoculation improves the relative water content of stressed plants, as observed by [37] in poplar and [38] in Capsicum annum. In our study, the PCA biplot (Figure 12) indicating increased RWC under treatments T5, T6, and T7 placed on the right side of the biplot confirms the importance of mycorrhizal inoculum in improving RWC. Increased RSI % in our study, as indicated by the placement of treatments T2, T3, and T4 for all three genotypes in the lower left corner of the PCA biplot under salt stress, is by the observations of [10] for chickpea and [39] for triticale. This is due to the production of harmful molecules called ROS. These ROS act like microscopic vandals, attacking the unsaturated fats within cell membranes, causing them to peroxidize and lose their integrity. This leads to leaky membranes, allowing precious water to escape and ultimately resulting in desiccation and stunted growth. Decreased RSI in Azadirachta indica under water stress after mycorrhizal inoculation was recorded by [40]. In this study, AMF also effectively shielded membrane lipids from oxidative damage, reducing RSI.

Our results showed that salinity stress triggered a dramatic reduction in chlorophyll pigments. This aligns with previous [41,42,43] findings for Avena sativa, Zea mays, and Jatropha curcas. A decline in chlorophyll in Moringa oleifera with increasing salinity was noted by [11] and [44], who reported similar findings in spinach exposed to saline irrigation. These reductions are attributed to increased chlorophyllase activity, leading to pigment degradation and ultimately reduced photosynthesis and growth. Salinity also hinders the synthesis of new proteins and disrupts the function of pigment–protein complexes [45,46]. Our research, however, indicates that AMF inoculation can effectively counteract these adverse effects. Compared to stressed plants without AMF, our AMF-inoculated plants displayed significantly higher chlorophyll content, as shown in the PCA biplot under treatments T5, T6, and T7. This finding aligns with similar observations in Cinnamomum camphora [47], common wheat [48], and Gleditsia sinensis [49]. This enhancement in chlorophyll content is likely due to improved mineral uptake facilitated by AMF, particularly magnesium, a crucial component of the chlorophyll molecule [47]. Consequently, AMF-inoculated plants exhibit greater photosynthetic activity, leading to better overall growth under saline conditions. Salinity stress reduced our study’s photochemical quantum yield (Fv/Fm). Salinity prevented the transfer of electrons from the primary acceptor (QA) to the secondary acceptor (QB) in PSII, causing a reduction in the Fv/Fm ratio [50,51]. In response to salt, a decrease in the maximum quantum yield of PSII was also reported by [52,53] in cotton and peas, respectively. Mycorrhizal inoculation improved photochemical quantum yield in all three varieties. The increment is due to the increased photosynthetic activity of PSII of mycorrhizal plants. The results are consistent with [54] for baby corn plants.

Due to salt stress, most crops experience reduced photosynthetic activity throughout their growth and development. In the present study, salt stress significantly reduced photosynthetic parameters like PR, TR, and SC in chickpea plants. Under salt stress, plants close their stomata to conserve water and prevent further salt uptake. This closure reduces PR (reducing the raw material needed for photosynthesis), TR, and SC [55]. To conserve water under high salinity, plants can reduce their SC and TR [56]. Studies have shown that the PR, TR, and SC of Zoysia matrella were significantly reduced under high salinity [57]. A similar decline in PR and TR is reported in sorghum [58]. Research has indicated that mycorrhizal colonization reduced the detrimental effects of salinity stress on maize and significantly improved its photosynthetic performance [59,60]. Enhanced transpiration rates are indicative of improved water uptake efficiency. This mitigated the risk of water deficits even under conditions of elevated osmotic pressure, leading to improved overall plant growth. Our PCA biplot indicates similar results.

Under stress conditions, various antioxidant enzymes become upregulated. They act as scavengers, neutralizing harmful ROS and mitigating the damaging impacts of oxidative stress on vulnerable substances such as lipids, proteins, and nucleic acids. Our findings on increased activity of SOD, CAT, and POX enzymes under salinity stress align with similar observations in rice [61], pea [62], and Azolla species [63]. This elevation in antioxidant activity helps plants rapidly scavenge ROS and maintain their levels below harmful thresholds. Specifically, SOD converts superoxide radicals into water and hydrogen peroxide, while CAT further breaks down hydrogen peroxide into water and oxygen [64]. In our study, AMF colonization further enhanced the activity of these antioxidant enzymes, facilitating even faster ROS scavenging and minimizing disruption to metabolic processes. PCA biplot clearly shows the highest activity of antioxidant enzymes (SOD, CAT, and POX) under treatment T8 for all three genotypes. This aligns with previous reports for basil [65] and wheat [66], where AMF application boosted antioxidant enzyme activity under stress. Similar benefits were observed in maize [67], lettuce [68], and Xanthoceras sorbifolium [69]. These findings collectively recommend that AMF can significantly improve plant performance under salt stress by supporting their antioxidant defense systems.

5. Conclusions

Salinity stress significantly impairs chickpea (Cicer arietinum) growth by disrupting key physiological and biochemical processes, including membrane integrity, water retention, and photosynthetic activity. This study demonstrated that inoculation with arbuscular mycorrhizal fungi (AMF) effectively alleviates these adverse effects by enhancing the antioxidant defense system and maintaining photosynthetic efficiency. The results underscore the potential of AMF as a sustainable strategy to improve chickpea resilience and productivity in saline environments.

Author Contributions

Conceptualization, P.P., S.D., and S.T.; Data curation, S.D., R.P., and J.M.M.; Formal analysis, Y.K.A., N.S., and N.L.; Funding acquisition, J.M.M. and K.M.E.; Investigation, S.D. and J.M.M.; Methodology, S.D., S.T., Y.K.A., N.S., and R.P.; Resources, S.D., Y.K.A., and K.M.E.; Software, P.P., S.D., N.S., P.K., and N.L.; Validation, S.D., R.P., and J.M.M.; Writing—original draft, S.D. and S.T.; Writing—review & editing, S.D., Y.K.A., R.P., and J.M.M. All authors have read and agreed to the published version of the manuscript.

Data Availability Statement

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

Acknowledgments

The authors extend their deep appreciation to Researchers Supporting Project number (RSPD2025R952), King Saud University, Riyadh, Saudi Arabia.

Conflicts of Interest

The authors declare no conflicts of interest.

Footnotes

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Figures

Figure 1. AMF (Rhizophagus fasciculatus) colonization in C. arietinum roots: (A) Heavily colonized root section of chickpea; (B) Spores (S), arbuscles (A), and hyphae (H) formed in roots; (C) Hyphae (H) and vesicles (V) in chickpea roots; (D) AMF spores (S) and hyphae (H) in the colonized roots; (E) Arbuscles (A), vesicles (V), spores (S), and hyphae (H) of AMF in the colonized roots; (F) Vesicle in the colonized chickpea root.

Figure 2. Impact of mycorrhiza on RWC of chickpea varieties under the influence of salt treatment. Bars are the means of three replicas (n = 3) ± S.E. Asterisks show significant differences at * p [less than] 0.05, ** p [less than] 0.01, respectively.

Figure 3. Impact of mycorrhiza on chickpea varieties’ RSI under salt treatment. Bars are the means of three replicas (n = 3) ± S.E. Asterisks show significant differences at * p [less than] 0.05, ** p [less than] 0.01 levels, respectively.

Figure 4. Impact of mycorrhiza on chickpea varieties’ total chlorophyll content under salt treatment. Bars are the means of three replicas (n = 3) ± S.E. Asterisks show significant differences at * p [less than] 0.05, ** p [less than] 0.01 levels, respectively.

Figure 5. Impact of mycorrhiza on the photochemical quantum yield of chickpea varieties under the influence of salt treatment. Bars are the means of three replicas (n = 3) ± S.E. Asterisks show significant differences at * p [less than] 0.05, ** p [less than] 0.01 levels, respectively.

Figure 6. Impact of mycorrhiza on the photosynthetic rate of chickpea varieties under the influence of salt treatment. Bars are the means of three replicas (n = 3) ± S.E. Asterisks show significant differences at * p [less than] 0.05, ** p [less than] 0.01, and *** p [less than] 0.001 levels, respectively.

Figure 7. Impact of mycorrhiza on the transpiration rate of chickpea varieties under the influence of salt treatment. Bars are the means of three replicas (n = 3) ± S.E. Asterisks show significant differences at * p [less than] 0.05, ** p [less than] 0.01 levels, respectively.

Figure 8. Impact of mycorrhiza on chickpea varieties’ stomatal conductance under salt treatment. Bars are the means of three replicas (n = 3) ± S.E. Asterisks show significant differences at * p [less than] 0.05, ** p [less than] 0.01 levels, respectively.

Figure 9. Impact of mycorrhiza on SOD activity of chickpea varieties under the influence of salt treatment. Bars are the means of three replicas (n = 3) ± S.E. Asterisks show significant differences at * p [less than] 0.05, ** p [less than] 0.01 levels, respectively.

Figure 10. Impact of mycorrhiza on chickpea varieties’ CAT activity under salt treatment. Bars are the means of three replicas (n = 3) ± S.E. Asterisks show significant differences at * p [less than] 0.05, ** p [less than] 0.01, and *** p [less than] 0.001 levels, respectively.

Figure 11. Impact of mycorrhiza on POX activity of chickpea varieties under the influence of salt treatment. Bars are the means of three replicas (n = 3) ± S.E. Asterisks show significant differences at * p [less than] 0.05, ** p [less than] 0.01, and *** p [less than] 0.001 levels, respectively.

Figure 12. PCA biplot for several chickpea genotype parameters under treatments T1 (control), T2 (2 dSm−1), T3 (3 dSm−1), T4 (4 dSm−1), T5 (control + AM), T6 (2 dSm−1 + AM), T7 (3 dSm−1 + AM), T8 (4 dSm−1 + AM).

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