Our earlier work described that, from ten genotypes of rice (Oryza sativa L.) under same levels of Cd toxicity, ‘Lu 9803’ was more resistant to Cd toxicity in hydroponic culture experiment (Afzal et al., 2018). The seeds of Lu 9803 were collected from Hubei Tianmen Di Long Seed Industry Co., Ltd., Hubei, China and the seeds were surface-sterilized with 10% (v/v) commercial bleach for 15 min followed by a thorough washing in distilled water. The seeds were germinated in a small box and placed in growth chamber under white lights (100 W, Guangdong PHILIPS Co., Guangdong, China) with a day/night temperature of 25 ± 2°C and day/night humidity of 80%. After 14 days of seed sowing, uniform sized seedlings were shifted to the pots (6 kg/pot). Uncontaminated soil, obtained from the research fields at Huazhong Agricultural University, was air dried and passed through a 5-mm sieve. The physicochemical properties of the soil used for the pot experiment were as follows: pH 7.12, available potassium 190 mg/kg, organic matter 13.72 g/kg, olsen-P, 17.07 mg/kg, alkaline hydrolysis nitrogen 54.13 mg/kg and total Cd, 0.072 mg/kg. The soil was artificially treated with Cd at various concentrations, i.e., 0 (no Cd), 0.5, and 1 mg/kg using CdCl₂. For iron (Fe) application, stock solution was made from 15 g of FeSO₄.7H₂O and 1 L of distilled water and Fe was supplied at 0 (no Fe), 1.5 and 3 mg/kg which were defined as low, moderate, and high Fe doses, respectively and distilled water was supplied on the control plants. After adding the Cd, the pots were equilibrated for 2 months with four cycles of saturation with distilled water and air drying. A complete randomized design (CRD) with three replications was used. This experiment was conducted in a glass house, under controlled conditions at the experimental station of Huazhong Agricultural University, Wuhan, China (114.20°E, 30.28°N; 50 m above sea level). The plants received natural light with day/night temperatures of 35/30°C and day/night humidity of 70/80%. All pots were monitored daily and weeding and metal-free water were provided as needed and every pot had one hole at the bottom to assimilate moisture. All plants in the glass house territory received natural light, with day/night temperature of 25/30°C and day/night humidity of 60/70%. The total time for plant growth was 146 days (ripening phase). Although, this experiment was conducted in pots, but for the collection of organic acids, two seedlings were transferred to the rhizoboxes which consist of plastic sheet, nylon net and wet soil (Javed et al., 2013). After 48 h, plants were taken from the rhizoboxes and the roots were washed with redistilled water to collect the exudates from root surface. The samples were filtered through a 0.45 µm filter (MillexHA, Millipore) and collected in Eppendorf tubes (Greger and Landberg, 2008). The collected samples were mixed with NaOH (0.01 M) in order to analyze the organic acids. However, the samples used for analysis of oxalic acid were not treated with NaOH (Javed et al., 2013). All chemicals used were of analytical grade and procured from Sinopharm Chemical Reagent Co., Ltd., China.

The plants were harvested in November 2018 (146 days after seed transfer) for the analysis of various morphological parameters. Leaves from each treatment group were picked for chlorophyll, carotenoid, and antioxidant analysis. A fully functional leaf was harvested for the various enzymatic and pigment studies. The leaves were washed with distilled water, placed in liquid nitrogen, and stored at −80°C for further analysis. The plants from each treatment were washed with tap water to remove debris and waste, and then with distilled water. The morphological measurements such as plant height, root length, plant fresh weight and plant dry weight were noticed at three different growth stages i.e., tillering stage (38 days old), vegetative stage (90 days old), and maturity stage (146 days old). Plant height was measured from the top of the leaf tips to the bottom of the roots and root length was also measured in the same way. Plant fresh weight was determined by measuring the weight of plant with a digital weighting balance. The plant samples were oven-dehydrated at 65°C for 72 h for Cd and ions concentration determination and the total plant dry weight was also measured. Yield parameters such as biological yield, 1,000 grain weight, straw yield, grain yield, relative growth rate and harvest index were also measured after the harvesting all the sampled plants at maturity stage (146 days old). Before oven dried, roots immersed in 20 mM Na₂EDTA for 15–20 min to remove Cd adhered to the surface of roots. Then, roots were washed thrice with distilled water and finally once with de-ionized water and dried for further analysis.

Leaves were collected for determination of chlorophyll and carotenoid contents. For chlorophylls, 0.1 g of fresh leaf sample was extracted with 8 mL of 95% acetone for 24 h at 4°C in the dark. The absorbance was measured by a spectrophotometer (UV-2550; Shimadzu, Kyoto, Japan) at 646.6, 663.6, and 450 nm. Chlorophyll content was calculated by the standard method of Arnon (1949).

Gas exchange parameters were also measured during the same days. Net photosynthesis (Pn), leaf stomatal conductance (Gs), transpiration rate (Ts), and intercellular carbon dioxide concentration (Ci) were measured from three different plants in each treatment group. Measurements were conducted between 11:30 and 13:30 on days with clear sky. Rates of leaf Pn, Gs, Ts, and Ci were measured with a LI-COR gas-exchange system (LI-6400; LI-COR Biosciences, Lincoln, NE, USA) with a red-blue LED light source on the leaf chamber. In the LI-COR cuvette, CO₂ concentration was set as 380 mmol/mol and LED light intensity was set at 1000 mmol/m² s, which is the average saturation intensity for photosynthesis in O. sativa (Austin, 1990).

The degree of lipid peroxidation was evaluated as malondialdehyde (MDA) contents. Briefly, 0.1 g of frozen leaves were ground at 4°C in a mortar with 25 mL of 50 mM phosphate buffer solution (pH 7.8) containing 1% polyethene pyrrole. The homogenate was centrifuged at 10,000 × g at 4°C for 15 min. The mixtures were heated at 100°C for 15–30 min and then quickly cooled in an ice bath. The absorbance of the supernatant was recorded by using a spectrophotometer (xMark™ Microplate Absorbance Spectrophotometer; Bio-Rad, USA) at wavelengths of 532, 600, and 450 nm. Lipid peroxidation was expressed as l mol/g by using the formula: 6.45 (A532-A600)-0.56 A450. Lipid peroxidation was measured by using a method previously published by Heath and Packer (1968).

To estimate H₂O₂ content of plant tissues (root and leaf), 3 mL of sample extract was mixed with 1 mL of 0.1% titanium sulfate in 20% (v/v) H₂SO₄ and centrifuged at 6000 × g for 15 min. The yellow color intensity was evaluated at 410 nm. The H₂O₂ level was computed by extinction coefficient of 0.28/mmol cm. The contents of H₂O₂ were measured by the method presented by Jana and Choudhuri (1981).

Stress-induced electrolyte leakage (EL) of uppermost stretched leaves was determined by using methodology of Dionisio-Sese and Tobita (1998). The leaves were cut into minor slices (5 mm length) and placed in test tubes having 8 mL distilled water. These tubes were incubated and transferred into water bath for 2 h prior to measuring the initial electrical conductivity (EC₁). The samples were autoclaved at 121°C for 20 min, and then cooled down to 25°C before measuring the final electrical conductivity (EC₂). Electrolyte leakage was calculated as by the following formula:

EL=(EC1/EC2)×100

To evaluate enzyme activities, fresh leaves (0.5 g) were homogenized in liquid nitrogen and 5 mL of 50 mmol sodium phosphate buffer (pH 7.0) including 0.5 mmol EDTA and 0.15 mol NaCl. The homogenate was centrifuged at 12,000 × g for 10 min at 4°C, and the supernatant was used for measurement of superoxidase dismutase (SOD) and peroxidase (POD) activities. SOD activity was assayed in 3 mL reaction mixture containing 50 mM sodium phosphate buffer (pH 7), 56 mM nitro blue tetrazolium, 1.17 mM riboflavin, 10 mM methionine, and 100 μL enzyme extract. Finally, the sample was measured by using a spectrophotometer (xMark™ Microplate Absorbance Spectrophotometer; Bio-Rad). Enzyme activity was measured by using a method by Chen and Pan (1996) and expressed as U/g FW.

POD activity in the leaves was estimated by using the method of Sakharov and Ardila (1999) by using guaiacol as the substrate. A reaction mixture (3 mL) containing 0.05 mL of enzyme extract, 2.75 mL of 50 mM phosphate buffer (pH 7.0), 0.1 mL of 1% H₂O₂, and 0.1 mL of 4% guaiacol solution was prepared. Increases in the absorbance at 470 nm because of guaiacol oxidation was recorded for 2 min. One unit of enzyme activity was defined as the amount of the enzyme.

Catalase (CAT) activity was analyzed according to Aebi (1984). The assay mixture (3.0 mL) was comprised of 100 μL enzyme extract, 100 μL H₂O₂ (300 mM) and 2.8 mL 50 mM phosphate buffer with 2 mM ETDA (pH 7.0). The CAT activity was measured from the decline in absorbance at 240 nm as a result of H₂O₂ loss (ε = 39.4 mM/cm).

Ascorbate peroxidase (APX) activity was measured according to Nakano and Asada (1981). The mixture containing 100 μL enzyme extract, 100 μL ascorbate (7.5 mM), 100 μL H₂O₂ (300 mM), and 2.7 mL 25 mM potassium phosphate buffer with 2 mM EDTA (pH 7.0) was used for measuring APX activity. The oxidation pattern of ascorbate was estimated from the variations in wavelength at 290 nm (ε = 2.8 mM/cm).

Plant ethanol extracts were prepared for the determination of non-enzymatic antioxidants and some key osmolytes. For this purpose, 50 mg of plant dry material was homogenized with 10 mL ethanol (80%) and filtered through Whatman No. 41 filter paper. The residue was re-extracted with ethanol and the two extracts were pooled together to a final volume of 20 mL. The determination of flavonoids (Pękal and Pyrzynska, 2014), phenolics (Bray and Thorpe, 1954), ascorbic acid (Azuma et al., 1999), anthocyanin (Lewis et al., 1998) and total sugars (Dubois et al., 1956) was performed from the extracts.

Fresh leaf material (0.1 g) was mixed thoroughly in 5 mL aqueous sulfosalicylic acid (3%). The mixture was centrifuged at 10000 × g for 15 min and aliquot (1 mL) was poured into a test tube having 1 mL acidic ninhydrin and 1 mL glacial acetic acid. The reaction mixture was first heated at 100°C for 10 min and then cooled in an ice bath. The reaction mixture was extracted with 4 mL toluene and test tubes are vortexed for 20 s and cooled. Thereafter, the light absorbance at 520 nm was measured by using UV–VIS spectrophotometer (Hitachi U-2910, Tokyo, Japan). The free proline content was determined on the basis of standard curve at 520 nm absorbance and expressed as µmol/g FW (Bates et al., 1973).

For nutrient analysis, plant roots and shoots were washed twice in redistilled water, dipped in 20 mM EDTA for 3 s and then, again washed with deionized water twice for the removal of adsorbed metal on plant surface. The washed samples were then oven dried for 24 h at 105°C. The dried roots and shoots were digested by using wet digestion method in HNO₃:HClO₄ (7:3 V/V) until clear samples were obtained. Each sample was filtered and diluted with redistilled water up to 50 mL. The root and shoot contents of Fe, Mg, and K and were analyzed by using Atomic Absorption Spectrophotometer (AAS) model Agilent 240FS-AA.

In order to determine the concentration of organic acids, freeze dried exudates were mixed with ethanol (80%) and 20 μL of the solutions was injected into C18 column (Brownlee Analytical C-183 µm; length 150 mm × 4.6 mm², USA). Quantitative analysis of organic acids in root exudates was executed with high performance liquid chromatography (HPLC) having a Flexer FX-10 UHPLC isocratic pump (PerkinElmer, MA, USA). The mobile phase used in HPLC was comprised of acidic solution of aceto-nitrile containing aceto-nitrile: H₂SO₄:acetic acid in ratios of 15:4:1 respectively, and pH of 4.9. The samples were analyzed at a flow rate of 1.0 mL/min for a time period of 10 min. The inner temperature of the column was fixed at 45°C and quantification of organic acids was carried out at 214 nm wavelength with the help of a detector (UV–VIS Series 200, USA) as described by Uddin et al. (2015). Freeze-dried samples were dissolved in redistilled water and the pH of the exudates was recorded with LL micro-pH glass electrode by using a pH meter (ISTEK Model 4005–08007 Seoul, South Korea).

Finely ground samples were digested with pure HNO₃ at 190°C for 45 min (10 min pre-heating, 15 min heating, 20 min cooling) in a microwave oven (Mars 6, CEM Corporation, USA) with the settings described in details by Jezek et al. (2015). Samples were diluted with 2% HNO₃ and determined by atomic absorption spectrophotometer (AAS) model Agilent 240FS-AA.

For TEM, leaf samples were collected and placed in liquid nitrogen. Small sections of the leaves (1–3 mm in length) were fixed in 4% glutaraldehyde (v/v) in 0.2-mol/L SPB (sodium phosphate buffer, pH 7.2) for 6–8 h and post-fixed in 1% OsO₄ for 1 h, then in 0.2-mol/L SPB (pH 7.2) for 1–2 h. Samples were dehydrated in a graded ethanol series (50%, 60%, 70%, 80%, 90%, 95% and 100%) followed by acetone, filtered and embedded in Spurr resin. Ultra-thin sections (80 nm) were prepared and mounted on copper grids for observation under a transmission electron microscope (JEOL TEM-1200EX) at an accelerating voltage of 60.0 kV or 80.0 kV.

Statistical analysis of data was performed with analysis of variance (ANOVA) by using a statistical program Co-Stat version 6.2, Cohorts Software, 2003, Monterey, CA, USA. All the data obtained was tested by two-way analysis of variance (ANOVA). Thus, the differences between treatments were determined by using ANOVA, and the least significant difference test (P < 0.05) was used for multiple comparisons between treatment means. Logarithmic or inverse transformations were performed for data normalization, where necessary, prior to analysis. Pearson’s correlation analysis was performed to quantify relationships between various analyzed variables. The graphical presentation was carried out by using Origin-Pro 2017. The Pearson correlation (heat-map) coefficients between the measured variables of O. sativa were also calculated using the RStudio software.

Heavy metal contamination of the environment through anthropogenic activities and/or natural processes is a widespread and serious problem (Saleem et al., 2020d; Saleem et al., 2019a; Saleem et al., 2020f). Heavy metals occur in various forms in soil, which differ greatly with respect to their solubility/bioavailability. The geochemical behavior of heavy metals in soil, their uptake by plants, and effect on crop productivity is affected by various physicochemical properties of soil (Hashem et al., 2020; Saleem et al., 2020i; Saleem et al., 2020k). Although, the plants grown in soil containing high levels of Cd show visible symptoms of injury reflected in terms of chlorosis, growth inhibition, browning of root tips and finally death (Afzal et al., 2018; Imran et al., 2020a). The inhibiting effect of Cd on fresh and dry mass accumulation, height, root length, leaf area, and other biometric parameters of plants are reported in almost all investigations. Differences in the degree of expressed phytotoxicity due to various Cd-concentrations applied to the root medium, the duration of treatment, as well as the characteristics of species and cultivars were established (Madhu and Sadagopan, 2020; He et al., 2017). Cd inhibited growth of most plant species (El-Esawi et al., 2020; Imran et al., 2020a; Javed et al., 2020). In the present study, we also noticed the same pattern, i.e., plant height, root length, plant fresh weight, plant dry weight at different growth stages (Fig. 1) (tillering, vegetative and maturity stage) and various yield parameters i.e., biological yield, 1,000 grain weight, straw yield, grain yield, relative growth rate and harvest index (Fig. 2) were also decreased with the addition of Cd in the soil. The maximum growth and yield parameters were observed in the plants which were grown in the control treatment, while increasing Cd concentration in the soil, induced a significant decreased in various growth, biomass, and yield parameters. Although, growth and yield-related attributes in O. sativa can be increased the exogenous application of FeSO₄, which non-significantly (P < 0.05) increased all these parameters, compared to those plants which were grown without the application of FeSO₄ (Figs. 1 and 2). This is because of Fe is required by plants for their normal growth parameters and it has been found beneficial regarding its role as reducing heavy metals toxicity in various plants (Danish et al., 2019a; Kobayashi et al., 2019; Zaheer et al., 2020b). This is also because Fe enhances protein properties (metabolic function and stock of amino acid function in plants), increases the photosynthetic processes for creating a healthy plant and provides a substantial growth in a short time in the stress condition or even in the plants grown in a normal soil (Ghasemi et al., 2014; Zaheer et al., 2020c).

Photochemical efficiency, chlorophyll content, and photosynthetic intensity are considered as sensitive indicators for the growth status of plants under Cd stress (Abbas et al., 2020; Rizwan et al., 2016a). Studies have shown that Cd is an effective inhibitor of photosynthesis (Bashir et al., 2018; He et al., 2017). Cd, in fact, strongly inhibits the synthesis of chlorophylls and their stable binding to proteins, thereby damaging the photosynthetic apparatus, in particular decreasing light-harvesting complex II and photosystems I (PSI) and II (PSII), and inhibiting some of the enzymes of the Calvin cycle (Hoseini and Zargari, 2013; Rizwan et al., 2019b). Cd can bind competitively to the essential Ca-binding sites in PSII during the photoactivation of the water-splitting system, and direct inhibition of oxygen evolution is also possible (Ali et al., 2015; Chen et al., 2011). Ferrous sulfate (FeSO₄) also known as copperas or black alum, common iron fertilizer for flowers and trees, it is easy to dissolve in water, can promote the chlorophyll formation, make color Nonglv and full of luster, effectively prevent the plant yellows due to iron deficiency, but also to regulate the soil acidity regulator, long-term use can change soil acidity and salinity, ensure that keep the soil acidity (Di Palma et al., 2015). In the present study, we have noticed that chlorophyll contents, carotenoid contents, Net photosynthesis (Pn), leaf stomatal conductance (Gs) and transpiration rate (Ts) were decreased with the increasing levels of Cd concentration in the soil, compared to the control treatment. Although, intercellular carbon dioxide concentration (Ci) showed non-significant results at all levels of Cd and FeSO₄ (Fig. 3). The photosynthetic pigments and gas exchange attributes in Cd-stressed can be increased by the application of FeSO₄, which non-non-significantly (P < 0.05) increased photosynthetic pigments and gas exchange attributes compared to those plants which were grown without the application of FeSO₄ (Fig. 3). These findings might be explained by the fact that Fe plays a critical role in metabolic reactions in organelles, such as pigment synthesis and chains of electron transport in photosynthesis (Wang et al., 2020). In addition, these findings revealed that photosynthetic capacity in response to Fe fertilizers depended on Fe types. This difference may be explained by the fact that Fe may hamper leaf penetration and movement in the apoplast because of the negatively charged plant cuticles and cell walls (Ghasemi et al., 2014; Wang et al., 2020; Zaheer et al., 2020b). A previous study has demonstrated that Fe can increase photosynthetic pigments and gas exchange attributes under Cd stress (Bashir et al., 2018).

In normal plant growth, reactive oxygen species (ROS) are produced during respiration and photosynthesis as a by-product and act as a signaling molecule that controls various processes in plants such as programmed cell death (Alam et al., 2020; Rana et al., 2020a). Although, stress conditions can disturb the dynamic equilibrium of production and elimination under normal growth in plants (Rana et al., 2020c; Saleem et al., 2019b), which promotes ROS accumulation, membrane lipid peroxidation, and disrupt the structure and function of cell membrane system (Ali et al., 2020a; Kamran et al., 2019; Mohamed et al., 2020; Rehman et al., 2019a). Numerous reports demonstrated that Cd stress increased the ROS generation in plants, which may cause oxidative damage in plants (Adrees et al., 2020; Madhu and Sadagopan, 2020; Wen et al., 2020). Moreover, Cd also caused cellular damage in plants which can be examined by estimating the level of lipid peroxidation in plants by disturbing the stability of membranes (Kong et al., 2020; Shahid et al., 2019). Environmental stress can trigger ROS production in plants, which results in oxidative damage. However, plants possess an efficient antioxidative defense system to detoxify the ROS generation (Ali et al., 2020b; Rana et al., 2020b; Rehman et al., 2020b). Antioxidants are involved in ROS metabolism by dismutation of superoxide anion to hydrogen peroxide (H₂O₂) and molecular oxygen (O₂) thus maintains superoxide radicals in a steady state (Yaseen et al., 2020; Zaheer et al., 2020a). Our results reported an increase in lipid peroxidation and oxidative stress in the roots and leaves of O. sativa (Fig. 4), This increase in lipid peroxidation and oxidative stress might be due to the progressive increase in soil Cd concentration, which is a stress factor triggering physiological responses in plants (Rizwan et al., 2016b; He et al., 2017). In fact, oxidative stress indicators showed a dose-dependent increase with the concentration of Cd in the soil (Fig. 4). In this study, enzymatic and non-enzymatic antioxidants were increased up to the level of 0.5 mg Cd/kg soil, which were decreased with further increase in Cd stress, i.e., 1 mg Cd/kg soil, compared to those plants which were grown in 0 mg Cd/kg soil (Figs. 5 and 6). The variable antioxidant response under different Cd stress conditions might be due to changes in gene expression and function of the protein in various plant tissues (El-Esawi et al., 2020; Hoseini and Zargari, 2013; He et al., 2017). In the present study, we also noticed the contents of sugar also decrease due to Cd exposure in the soil while proline contents were increased with the increase in Cd concentration in the soil (Fig. 6). This is because of proline accumulation in plant tissue/organs is a response to Cd toxicity, which might be associated with signal transduction and prevents membrane distortion (Abbas et al., 2020; Javed et al., 2020). Although, oxidative stress indicators, i.e., MDA contents, H₂O₂ initiation and EL (%) decreased in Cd-stressed O. sativa by increasing enzymatic (SOD, POD, CAT and APX) and non-enzymatic (phenolic, flavonoid, ascorbic acid, and anthocyanin) contents under the application of FeSO₄ compared to those plants which were grown without the application of FeSO₄. Increasing the activities of various antioxidants suggests that the plant has a better defense system, which can scavenge ROS generation (Rizwan et al., 2019b; Zafar-Ul-Hye et al., 2020). It is well known that the exogenous supplementation of Fe decreases oxidative stress in plants (Bashir et al., 2018; Ghasemi et al., 2014).

Essential nutrients are required for normal growth of plants. Numerous reports demonstrated that the uptake and translocation of essential elements in plants were restricted under Cd stress (Rizwan et al., 2016b; Rizwan et al., 2019a; Rizwan et al., 2019b). Excess Cd decreased the Fe, Mg and P contents in numerous plant species which may cause ions deficiency in plants. With increasing concentrations of Cd (0.5 and 1 mg/kg) in growth medium, the contents of Fe, Mg and P in the roots and shoots of O. sativa were decreased significantly (P < 0.05) when compared to those plants grown without Cd addition (Fig. 7). Similarly, Cd concentration in the roots and shoots of plants increased significantly (P < 0.05) by increasing the Cd concentration in the soil, compared to those plants which were grown in the without Cd addition (Fig. 8). According to the results, the contents of essential minerals (Fe, Mg and P) were slightly higher in shoots as compared to roots (Fig. 7). This might be attributed to small acropetal translocation of these elements possibly due to reduction in mineral ions accumulation as Cd and these ions compete for ZIP transporters at uptake sites (Abbas et al., 2020; Javed et al., 2017). Moreover, the decrease in essential ions accumulation in different organs of O. sativa seedlings under varying Cd concentrations might also be due to the alteration in the physiological processes such as the failure of biosynthesis of chlorophyll and carotenoid contents (Afzal et al., 2018; Imran et al., 2020a; Madhu and Sadagopan, 2020). The contents of Fe, Mg and P in the roots and shoots of O. sativa can be increased by the application of FeSO₄, which significantly (P < 0.05) increased the contents of Fe, Mg and P in the roots and shoots of O. sativa, compared to those plants which were grown without the application of FeSO₄ (Fig. 7). However, application of FeSO₄ decreased the contents of Cd in the roots and shoots of O. sativa compared to those plants which were grown without the application of FeSO₄ (Fig. 8). The reduced Cd concentration in all organs of O. sativa might be due to the higher Fe contents and other nutrients in the plants, as Cd and some other nutrients showed antagonistic effects reported previously by Danish et al. (Danish et al., 2019b). Excess Cd reduced the Fe and other nutrients concentration and its root to shoot translocation in O. sativa (Rizwan et al., 2016b) while exogenous Fe supply reduced the Cd concentrations in O. sativa compared to the plants without Fe supply (Bashir et al., 2018). It was also reported that both Fe and Cd share the same entry routs within O. sativa and the expression of Fe transport system may be induced under Cd stress (Afzal et al., 2018; Bashir et al., 2018).

Since Fe supply reduced Cd accumulation in O. sativa in this study without affecting tolerance mechanism and Cd acropetal translocation, we speculated that an avoidance mechanism plays a pivotal role in Fe-mediated reduction of Cd uptake (Fig. 8). Root exudates, especially fumaric acid, formic acid, acetic acid, citric acid, malic acid, and oxalic acid are suggested to be efficient ligands to alleviate Cd toxicity in plants as they can reduce the bioavailability of Cd (Greger and Landberg, 2008; Konate et al., 2017; Ullah et al., 2016). Organic acids are exuded as anions by anion channels, and their release should be balanced by cations/proton efflux as reported for poplar roots under metal stress (Javed et al., 2019; Shahid et al., 2019). O. sativa, metal stress impaired the pumping activities of H⁺ ATPase, which caused a decreased efflux of protons. This could explain the application of FeSO₄ in the present study at low Cd treatment, besides a direct binding of Cd to organic anions (Fig. 8). Addition of organic acids to soil promotes ammonification which produces protons and leads to a pH decrease. Therefore, acidosis of root exudates might also depend on nitrogen conversion triggered by decomposition of the exuded organic acids (Abbas et al., 2020; Javed et al., 2020).

Addition of Cd can change the anatomic and structural features of cells, which is considered as the worst effect of Cd. Shah and Dubey (1995) observed low mitotic index, cell division, cell proliferation, and chromosomal aberration in various plant species under Cd stress. Moreover, increased nucleolus and vacuole numbers, condensed cytoplasm, reduced mitochondria number, plasmolysis, enlarged vacuoles, disorganized chloroplasts, and ruptured nuclear envelope were found in the root and leaf cells of different crops under Cd stress. Ali et al. (2013) observed cracked cell walls, undeveloped mitochondria, plasmolysis, and endoplasmic reticulum absence in the root tip cells of Brassica napus grown under a high concentration of Cd (500 mmol/L). The ability to detect significant genetic alterations in plants at different Cd exposure levels, prior to the onset of physiological effects, might serve as a useful molecular biomarker for the early detection of Cd exposure and indicator of related biological effects (Ali et al., 2013; He et al., 2017). Similar results, we also observed in the present study, that addition of Cd in the soil affected most of the cellular organelles of the cell especially ultra-structure of chloroplast (Fig. 9). Although, the ultra-structure of double membranous bounded organelles can be improved by the application of FeSO₄, and little improvement in chloroplast structure occurred with the application of different levels of Fe in the soil (Fig. 9). The concentration of Fe in the cells of the plants is crucial because it can regulate many important amino acids, cell division, and cell construction (Ghasemi et al., 2014).

A heatmap-histogram analysis was also constructed to explore the relationship between the different growth, nutrient uptake and organic acids exudation and Cd uptake attributes (Fig. 10). The significant differences were observed in the plant growth parameters, photosynthetic pigments, gas exchange parameters, nutrient uptake and sugar contents in the treatments which were not spiked artificially with Cd (comprised with application of FeSO₄). While rest of the heat-map showing non-significant results with all other parameters with the treatments of Cd in the nutrient solution. Although, blue color is showing non-significant differences within the treatments, while black color is depicting a significant different in the histogram study. This histogram study is showing a clear differences of Cd toxicity on ecophysiology of O. sativa under the application of FeSO₄ in the soil.