Supplemental Effects of Biochar and Foliar Application of Ascorbic Acid on Physio-Biochemical Attributes of Barley (Hordeum vulgare L.) under Cadmium-Contaminated Soil

Abstract

Biochar, prepared from organic waste materials, can improve the quality of contaminated soil areas. Biochar can be used as an economic centerpiece over other available resources and can properly utilize large amounts of waste. Soil contaminated with cadmium (Cd) is a worldwide problem that poses potential agricultural and human health hazards. Moreover, Cd toxicity causes serious problems for sustainable food production, especially in food crops like barley. High cadmium concentration in soil is phytotoxic and decreases plant growth and ultimately yields. Biochar and ascorbic acid in ameliorating Cd stress are economically compatible and consistent approaches in agriculture. The present study aimed to evaluate biochar’s and foliar-applied ascorbic acid’s influence on some growth and biochemical characteristics of barley (Hordeum vulgare L.) to Cd stress. The soil was supplemented with biochar 2% w/w and 20 mg Cd kg⁻¹. The foliar application of 30 mM ascorbic acid was done on plants. The results revealed that Cd stress decreased chlorophyll a, chlorophyll b, total chlorophyll, and carotenoids. It also increased oxidative stress indicators, i.e., APX, COD, POD, flavonoids, anthocyanin, phenolics, and electrolyte leakage, in barley with Cd-contamination. A significant enhancement in root and shoot length, gas exchange attributes, and chlorophyll contents validated the effectiveness of Bio + Asa treatments over all other treatments under Cd contamination. In conclusion, the sole applications of biochar and Asa in Cd contamination are also effective, but Bio + Asa is a better amendment for Cd stress alleviation in barley plants.

1. Introduction

The agriculture sector faces many problems, including more food production and mitigation of environmental stress factors to secure future food demands [1]. Soil contamination by trace elements is one of the predominant abiotic stresses that decrease plant productivity. In addition, metal contamination deteriorates food quality and affects human health by entering the food chain. Soil contamination with trace elements is mainly due to anthropogenic activities, including overuse of fertilizers, mining, industrial wastes, and pesticides in agricultural areas. The natural contributors of trace elements to the environment are the weathering of rocks and volcanic activities [2].

Trace element toxicity in plants includes both biologically essential and non-essential elements (Zn, Co, Cu, Mn, Cd, Cr, As, and Pb). Plant uptake all these elements only in trace amounts for normal functions, and they are called micronutrients [3,4,5,6,7,8]. Whenever any of these elements are present at elevated concentrations, it causes metal toxicity [2,9,10]. Soils polluted with metal elements pose severe ecological constraints, as these are not environmentally biodegradable and persistent, ultimately threatening living organisms, especially humans [2,11,12,13,14,15].

Plant uptake trace elements from the soil through roots and their absorbance depend upon their bioavailability in soils. Therefore, plant roots are the first organ that experiences Cd toxicity and enhanced oxygen radicals production and growth inhibition [16]. Cadmium is the 7th one among 20 metals causing toxicity to plants. The availability of Cd depends on other cation concentrations, cation exchange capacity, soil pH, soil texture, and organic matter [17]. By reducing trace elements’ bioavailability, their uptake can be reduced due to the stabilization of trace metals and organic pollutants in the soil. In recent approaches, biochar application as a soil amendment of heavy metal toxicity has proved effective and is widely accepted [18,19,20]. Biochar minimizes the absorption of trace elements by roots, thus lowering its toxicity and increasing soil fertility. Additionally, the addition of biochar to soil increases soil carbon sequestration and biological activities and decreases greenhouse gas emissions [2].

Alternatively, using several hormones, osmo-protectants, minerals, and vitamins would be another promising option under abiotic stresses and could protect several crops such as barley [21]. For example, ascorbic acid (Asa) is a water-soluble vitamin and is well known to scavenge oxidative stress efficiently, and it reduces the number of free radicals that are generated as a consequence of several abiotic stress factors [21]. The foliar-applied ascorbic acid enhances its indigenous production in plant cells, thus mitigating stressful conditions [22,23]. Ascorbic acid is helpful in the maintenance of photosynthesis, cell wall expansion, plant hormone production, regulation of antioxidant systems, ion uptake, biological yield, and harvest index [21,24,25].

Barley (Hordeum vulgare L., Gramineae) is the fourth major cereal grain among cereal crops worldwide [1,21]. However, its cultivation in Pakistan is continuously decreasing because of insufficient compost use, poor soil health, and allied abiotic stresses [26]. In 2014, Pakistan produced 67 thousand tonnes, which now in 2019 and 2020 has become 63 thousand tonnes [27]. It is consumed for humans and used as fodder for livestock and in the malting and brewing process [21,28]. In the current experimental study, the effect of biochar and ascorbic acid application was studied using a pot experiment on barley subjected to Cd stress. Their performance was assessed in terms of growth, photosynthetic pigments, antioxidant activities, and ion accumulation. It was hypothesized that plants with ascorbic acid and biochar might perform better than untreated barley plants under Cd stress. In the current study, barley was selected as an essential nutritional crop, as it is economically important and among the most widely consumed plants.

2. Materials and Methods

2.1. Seed Sterilization

Seeds of barley (Hordeum vulgare L. Genotype B-14011) were obtained from Ayub Agriculture Research Institute, Faisalabad, Pakistan. All seeds were disinfected using 95% ethanol for 1 min; then, a 70% sodium hypochlorite solution (NaOHCl) for 10 min. The seeds were washed with distilled water six times.

2.2. Experimental Design

In 2020, a pot experiment was performed in the greenhouse of the Old Botanical Garden, University of Agriculture Faisalabad, Faisalabad, Pakistan. The treatments consisted of (a) control, (b) biochar, (c) biochar + ascorbic acid = Bio + Asa, (d) ascorbic acid spray = Asa, arranged into two groups: (Cd-contaminated and non-contaminated) and 3 replications in a completely randomized design (CRD).

2.3. Seeds Sowing and Pot Preparation

Barley seeds were sown in plastic pots filled with 5 kg of soil. The sowing season was January 2020. The seedlings were irrigated thrice a week until the termination of the experiment. Each 5 kg plastic pot was provided the desired dose of biochar (non-biochar and 2% w/w of pot soil). The biochar was prepared from pyrolysis of air-dried vegetable waste and then was powdered and sieved through a 2 mm sieve.

2.4. Cadmium (Cd) Contamination

Cd treatment was prepared by spraying the reagent into the soil. Cadmium nitrate (Cd(NO₃)₂·4H₂O) was mixed to attain the homogenized concentration of 20 mg Cd kg^–1 of dry soil, keeping in mind the threshold limit in the plant (0.2–1.5 ppm) and soil (3–6 mg kg⁻¹ soil) suggested by the Commission Regulation (EU) [29], the Indian standard [30], and the WHO/FAO [31].

2.5. Ascorbic Acid (Asa)

The solution of Asa was prepared with a 30 mM strength containing 0.1% of Tween 20 (Polysorbate 20) as a surfactant. Controlled barley plants were only sprayed with distilled water. The ascorbic acid foliar application was provided four weeks after sowing. The plants were harvested after 45 days of sowing. The concentrations of Asa, Cd, and biochar in the present study were considered from the literature.

2.6. Growth Attributes

The lengths of soot and roots were determined using a measuring tape. Immediately after harvesting, the fresh weight was estimated by using a digital weighing balance. The samples were preserved at −30 °C for a more fresh analysis. Sample plants from individual treatments were oven-dehydrated at 65 °C for 3 days.

2.7. Measurement of Chlorophyll Contents and Gas Exchange Characteristics

Gaseous exchange parameters: the transpiration rate, the stomatal conductance, and the photosynthetic rate were measured on leaves of three plants. Each replicate used the infrared gas analyzer (Cl-340 Handheld Photosynthesis System, Washington, DC, USA). All these measurements were recorded between 11 a.m. and 2 p.m. with a photosynthetic photon flux density (PPFD) not lower than 1800 μmol m⁻² s⁻¹. The total chlorophyll was estimated following the Lichtenthaler and Wellburn [27] method. In addition, chlorophyll a, chlorophyll b, and carotenoid contents were measured as stated by the standard Arnon [28] protocol. For this, fresh leaves (0.1 g) were homogenized in 8 mL of acetone (95%) at 4 °C for 1 day in the dark. The optical density was recorded at 646, 663, and 450 nm by a spectrophotometer (UV-2550; Shimadzu, Kyoto, Japan).

2.8. Electrolyte Leakage

Plant electrolyte leakage (EL) was estimated by incubating leaves at 23 °C in distilled water for 24 h in the dark. Next, the samples were vortexed, and initial electrical conductivity was recorded by using a conductivity meter. Then, at 60 °C, samples were autoclaved for 15 min. Then final conductivity of samples was measured after cooling at room temperature. The following equation was used to calculate the electrolyte leakage [32]:

$$\mathrm{Electrolyte}\mathrm{leakage}\left(\mathrm{EL}\%\right)=\left(\frac{\mathrm{Initial}\mathrm{Electrical}\mathrm{Conductivity}}{\mathrm{Final}\mathrm{Electrical}\mathrm{Conductivity}}\right)\times 100$$

(1)

2.9. Oxidative Stress Indicators

Malondialdehyde (MDA) contents were estimated by grinding 0.1 g fresh samples at 4 °C in a 50 mM concentration of 25 mL phosphate buffer with 7.8 pH and 1% concentrated polyethene pyrrole solution. The homogenous reaction mixture was then centrifuged at 10,000× g for 15 min at 4 °C. After this, the solution was heated at 100 °C for 20 min and immediately cooled in ice-cold water. The color intensity was recorded at 450, 532, and 600 nm wavelengths. Lipid peroxidation was indicated as 1 mol g^–1 following the protocol provided by Heath and Packer [33].

$$\mathrm{MDA}({\mathsf{\mu }\mathrm{mol}\mathrm{g}}^{-1})=6.45\left(\mathrm{A}532-\mathrm{A}600\right)-(0.56\times \mathrm{A}450)$$

(2)

The determination of H₂O₂ levels of barley plant samples was done by homogenously mixing 3 mL of leaf extract, 1 mL of 0.1% titanium sulfate, and 20% H₂SO₄ v/v. The solution was centrifuged at 6000× g for about 15 min. The colour absorbance was recorded at 410 nm by using a spectrophotometer. H₂O₂ contents were measured by a 0.28 mmol^–1 cm^–1 extinction coefficient, followed by Jana and Choudhuri [34].

2.10. Determination of Antioxidant Enzymatic Activities

Peroxidase (POD) enzyme activity in the leaves was found as reported by the Sakharov and Ardila [32] method by using a guaiacol catalyst. Three mL of the reaction mixture was prepared with 0.05 mL of enzyme concentrate, 2.75 mL of phosphate buffer with a 50 mM strength and 7.0 pH, 0.1 mL of 1% H₂O₂, and 0.1 mL of 4% guaiacol solution. Absorbance was noted at a 470 nm wavelength. The unit enzyme activity was accounted for as the contents of peroxidase enzyme present.

Catalase (CAT) concentration in cells was assayed by following the Aebi [33] method. For this, 3.0 mL of a reaction mixture comprised 100 L of enzyme extract, 100 L of 300 mM concentrated H₂O₂, 2.8 mL of 50 mM phosphate buffer, along with 2 mM of ETDA with a pH = 7.0. The activity of the CAT enzyme was measured at 240 by a decline in absorbance as by H₂O₂ loss. Finally, the superoxide dismutase (SOD) activity was estimated by following the Beauchamp and Fridovich method [35].

2.11. Estimation of Proline, Sugars, and Non-Enzymatic Antioxidants

To estimate proline and soluble sugars and several non-enzymatic antioxidants, ethanol extracts of leaf samples were prepared using 50 mg of dried leaf material and were homogenized in 10 mL of 80% ethanol. Then, this solution was filtered, followed by re-extraction in ethanol. The 20 mL of the final volume was maintained by mixing both sample extracts, and this mixture was used to determine contents like total soluble proteins [36], anthocyanin [37], phenolics [38], flavonoids [39], total sugars [40], and ascorbic acid [41] contents.

For the estimation of proline, 0.1 g of fresh leaves were extracted in a 5 mL sulfosalicylic acid (3%) and then centrifuged (at 10,000× g) for 15 min. After this, a 1 mL aliquot was taken in a test tube containing 1 mL of glacial acetic acid and 1 mL of acidic ninhydrin mixture. It was then boiled for 10 min at 100 °C and immediately cooled down in an ice bath and then vortexed for 20 s and cooled down at room temperature. Absorbance at a 520 nm wavelength was recorded using a spectrophotometer [42].

2.12. Analysis of Cadmium Contents

The leaf samples were immersed in HNO₃-HClO₄ (3:1, v/v) overnight [43]. Then, 5.0 mL of HNO₃ and digesting samples on the hot plate were added until a clear solution was obtained. Cd contents were evaluated using an atomic absorption spectrophotometer. Cd concentration measurements were estimated from the working curve after calibrating the instrument with the standards of known concentrations [44].

2.13. Statistical Analysis

A two-way analysis of variance (ANOVA) was carried out for data evaluation, and the difference in treatments was determined. A mean comparison test between treatments was made using the least significant difference test (p < 0.05) [45]. Logarithmic transformations for data normalization were performed where necessary before analysis. To compute associations among various analyzed variables, we implemented Pearson’s correlation analysis. The graphical demonstration of data was carried out by Origin 2021 [46].

3. Results

3.1. Root and Shoot Length

The effect of applied treatments was significant on root and shoot length. The results showed that Bio + Asa were significantly different from control-treated plants for improving the root length in Cd-contaminated and non-contaminated soils. In Cd-contaminated soils, biochar and Asa were statistically alike with Bio + Asa for root length. However, Bio + Asa were significantly better over biochar and Asa under non-contamination for root length (Figure 1A). For shoot length, no significant change was noted between Bio + Asa and biochar under Cd contamination. However, Bio + Asa and biochar significantly improved shoot length over Asa and the control under Cd contamination. Bio + Asa differed significantly for an increase in shoot length in non-contaminated soils. Application of biochar and Asa remained non-significant with one another but differed significantly over the control for an increase in shoot length under non-contaminated conditions (Figure 1B).

3.2. Chlorophyll Contents

Application of treatments significantly affects the chlorophyll a, b, and total chlorophyll contents under Cd-contaminated and non-contaminated soils. Chlorophyll a was significantly decreased in Cd contamination over non-Cd contamination without amendments. A significant increase in chlorophyll a was observed where Bio + Asa was applied compared to the control under Cd-contaminated soil. Biochar also differed significantly for improvement in chlorophyll a over Asa and the control in Cd-contaminated soil. No significant change was noticed among biochar and Bio + Asa for chlorophyll a under non-Cd-contaminated soils. However, biochar, Bio + Asa, and Asa significantly enhanced chlorophyll a in non-Cd-contaminated soils (Figure 2A).

For chlorophyll b, Bio + Asa, Asa, and biochar remained significantly different from the control under Cd-contaminated soils. Application of Bio + Asa significantly increased chlorophyll b among all the treatments in non-Cd-contaminated soils. However, Asa caused a significant decrease in chlorophyll b over biochar and the control in non-Cd-contaminated soils. Biochar was statistically alike with the control for chlorophyll b in non-Cd-contaminated soil (Figure 2B).

The Bio + Asa treatment was significantly best for improvement in total chlorophyll compared to control plants in Cd-contaminated soil. Sole application of biochar and Asa also remained significantly better for enhancing the total chlorophyll over the control under Cd contamination. However, the biochar application was significantly better than Asa for enhancement in total chlorophyll in Cd contamination. In non-Cd-contaminated soil, Bio + Asa was significantly best for total chlorophyll over all the treatments. Biochar also differed significantly over the control and Asa for enhancement in total chlorophyll in non-Cd-contaminated soil. However, no significant change was observed between Asa and the control for total chlorophyll in non-Cd-contaminated soil (Figure 1C).

A significant increase in carotenoids validated the effectiveness of Bio + Asa overall under treatments in Cd-contaminated and non-contaminated soils. The results also showed that the sole application of biochar also performed significantly better for enhancing carotenoids from the control under Cd contamination and non-contaminated conditions. Application of Asa significantly increased carotenoids in Cd-contaminated soil but caused a significant decrease in non-Cd-contaminated soil (Figure 2D).

3.3. Gas Exchange Attributes

The influence of treatments was significant on stomatal conductance, net photosynthesis, and the transpiration rate under non-contaminated and Cd contaminated conditions. No significant change among all the treatments was noted in Cd-contaminated soil for net photosynthesis. However, in non-contaminated soil, biochar and Bio + Asa remained significantly better than the control for the improvement in net photosynthesis. The Asa treatment remained non-significant for net photosynthesis over the control in non-contaminated soil (Figure 3A). In Cd-contaminated soils, all treatments were statistically alike for stomatal conductance. However, Bio + Asa and Asa significantly decreased stomatal conductance over biochar and the control, for stomatal conductance under non-Cd-contaminated soil (Figure 3B). For the transpiration rate, Bio + Asa differed significantly over the control in Cd-contaminated soil. All other treatments were statistically alike for a transpiration rate under Cd-contamination. In non-Cd-contaminated soil, Bio + Asa and biochar were significantly different over the control for the transpiration rate (Figure 3C).

3.4. SOD, POD, and APX

For SOD (Figure 4A) and POD (Figure 4B), a significant decrease was noted where Bio + Asa, Asa, and biochar were applied under Cd contamination and non-contamination. Bio + Asa and Asa significantly reduced APX over biochar and the control under Cd contamination and non-contamination. No significant change in all the treatments was noted between biochar and the control for APX under Cd-contamination and non-contamination (Figure 4C).

3.5. Catalase, Phenolics, Flavonoids, and Anthocyanin

Bio + Asa significantly differed from the control in decreasing catalase (Figure 5A), phenolics (Figure 5B), flavonoids (Figure 5C), and anthocyanin (Figure 5D) in both Cd-contaminated and non-contaminated plants. In Cd-contaminated soils, biochar and Asa differed significantly for catalase, phenolics, flavonoids, and anthocyanin. However, biochar and Asa were significantly better over control for phenolics, flavonoids, and anthocyanin under non-Cd-contaminated soil. For catalase, biochar remained non-significant over the control under non-Cd contamination.

3.6. Ascorbic Acid, MDA, H₂O₂, and Electrolyte Leakage

Treatments significantly affect ascorbic acid, MDA, H₂O₂ and electrolyte leakage under Cd-contaminated and non-Cd-contaminated soil. Bio + Asa, biochar, and Asa significantly decreased ascorbic acid in Cd-contaminated conditions. In non-Cd contamination, Bio + Asa and Asa differed significantly, but biochar remained non-significant over the control for ascorbic acid. In MDA (Figure 6A) and H₂O₂ (Figure 6B), Bio + Asa, biochar, and Asa caused a significant decline compared to the control under Cd-contamination and non-Cd contamination. For electrolyte leakage (Figure 6C), biochar and Bio + Asa caused a significant decline, but Asa was non-significant over the control in the Cd-contamination condition. In non-Cd contamination, biochar, Asa, and Bio + Asa significantly decreased electrolyte leakage from the control.

3.7. Proline, Total Soluble Proteins, Root Cd Concentration, and Shoot Cd Concentration

The results showed that the effect of the provided treatments was significant on proline, total soluble sugar, and Cd concentration in roots and shoots. Bio + Asa were significantly better over biochar and Asa for decreased proline (Figure 7A) in Cd-contaminated and non-Cd-contaminated conditions. A significant increase was noted in Bio + Asa for total soluble proteins, but the no-significant change was noticed among biochar and Asa over the control under Cd contamination. Bio + Asa and Asa significantly increased soluble sugars over the control under non-Cd contamination (Figure 7B). Bio + Asa, biochar, and Asa differed significantly over the control for the decrease in the root (Figure 7C) and shoot (Figure 7D) cadmium concentration under Cd-contaminated conditions. In non-contaminated soil, biochar and Bio + Asa were significant, but Asa was non-significant over the control for the decrease in the Cd concentration of the roots. However, Asa, biochar, and Bio + Asa remained significant for the decrease in shoot Cd concentration over the control in non-Cd-contaminated soil.

3.8. Pearson Correlation

Pearson correlation showed that shoot and root Cd concentration was significantly negative in correlation with shoot length, chlorophyll a, chlorophyll b, total chlorophyll contents, stomatal conductance, the transpirational rate, and total soluble sugars in Cd contamination. Attributes, i.e., SOD, POD, APX, catalyse, phenolics, flavonoids, anthocyanin, ascorbic acid, MDA, H₂O₂, electrolyte leakage, and proline, were significantly positive in correlation with shoot and root Cd concentration under the Cd-contaminated condition (Figure 8a). In non-Cd-contaminated conditions, a parallel trend was observed. High Cd in shoots and roots enhanced the antioxidants and decreased the growth attributes, chlorophyll contents, and gas exchange attributes (Figure 8b).

4. Discussion

The current study was designed to investigate the effect of biochar and ascorbic acid in Cd-contaminated soil. Barley was selected because of its major contribution to food consumption and, because of Cd accumulation via the food chain, it would be harmful to health. Biochar can sorb contaminants like Cd on its surface, thus reducing metal toxicity in plants [47,48]. Additionally, exogenously applied, several osmoprotectants like vitamins, minerals, and other micronutrients reduce environmental stress and improve plant vigor [21]. Therefore, in this study, the possible amendments of barley plants to Cd stress through foliar-sprayed ascorbic acid, biochar, and their combined application was examined by investigating its various physio-biochemical attributes. In this study, the application of treatment Bio + Asa proved effective in increasing plant length and photosynthetic pigments in cadmium stress compared to non-contaminated plants. In the literature, the increase in plant biomass, height, and photosynthetic pigment contents is linked with the antioxidant effect of ascorbic acid and biochar addition to soil [2,21].

Ascorbic acid directly reacts with several reactive oxygen species and improves plant redox status [49]. Our findings are in accordance with other ascorbic acid foliar sprays related to the plant protection mechanisms from oxidative stress, and the current study is supported by other scientific reports by Barzegar et al. [49] and Noreen et al. [21], in which foliar application with ascorbic acid spray enhanced biomass in sweet pepper [21,50].

Biochar application in Cd-polluted soils decreased Cd accumulation in plant tissues and grains [2]. Trace elements’ removal from the rhizosphere occurs by their sorption due to the carbonaceous properties of biochar and thus decreases their availability to plants [2,51]. Similarly, Puga et al. [52] reported that the nutritional contents increased in Mucuna aterrima and Jack bean shoots with biochar application [52]. Biochar increased mineral concentrations in rice grown in paddy soil contaminated with trace elements [53]. The current experiment validates this result. An applied treatment mitigates metal stress symptoms in barley and improves plants’ vigor. Their foliar application up regulates activities of non-enzymatic antioxidants like ascorbic acid under stressful conditions [21]. Kamal et al. [54] reported that ascorbic acid biosynthesis was enhanced by foliar application on cotton under heat stress [54]. Ascorbic acid is generally produced endogenously in plants during abiotic stress; however, exogenously sprayed application may further enhance its synthesis under different stress factors like soil metal contamination [55]. In the current study, a significant positive effect of Bio + Asa treatment was observed in increasing phenolic, ascorbic acid, and flavonoid concentration in barley during cadmium stress. Ascorbic acid foliar spray elevated anthocyanin contents.

Several primary and secondary metabolites, i.e., soluble sugars and proline, play an important role in cell osmotic adjustments. Proline accumulates in high concentrations during stressful conditions, which is reported in many articles [21,56,57,58]. Our results justify these findings. Biochar and ascorbic acid effectively increased proline and soluble sugar contents in barley in combined treatment and separate applications in Cd stress.

This is an undisputed fact that ROS are ubiquitously involved in cell signalling and that they regulate several physiological and developmental processes only in small quantities. Their concentrations become high during abiotic stress that disrupts plant metabolic activities. Antioxidants are well-known ROS scavengers [16]. In agreement with these results, enzymatic antioxidants like SOD, POD, APX activities, and catalase contents of barley plants were increased in Cd stress, and the application of Bio + Asa’s combined application had significant positive effects.

Antioxidant enzymes reduce the hydrogen peroxide levels and lipid hydro peroxides, which cause membrane peroxidation, thus maintaining normal cell functions despite having high levels of ROS accumulated in root cells under Cd stress [16,59]. Our results justify these facts. In our findings, MDA levels were higher in cadmium stress, and H₂O₂ was produced in lesser amounts in ascorbic-acid-treated plants in both contaminated and non-contaminated groups.

5. Conclusions

In conclusion, the application of ascorbic acid and biochar for the remediation of cadmium toxicity positively affected barley growth and physiological and yield attributes. Plant physiological parameters like shoot length and root length significantly improved with the application of Asa and biochar in mitigating Cd stress. Among all applied treatments, Bio + Asa for barley had more efficacious effects than all other treatments to ameliorate metal toxicity. Further investigation is suggested at the field level in different climatic zones to validate the current results of biochar application as an organic matter substitute with ascorbic acid for mitigation of cadmium stress in plants.