Research Article | Open Access
Poornima D. Vijendra, Kavitha M. Huchappa, Roopa Lingappa, Giridhara Basappa, Sathisha G. Jayanna, Vadlapudi Kumar, "Physiological and Biochemical Changes in Moth Bean (Vigna aconitifolia L.) under Cadmium Stress", Journal of Botany, vol. 2016, Article ID 6403938, 13 pages, 2016. https://doi.org/10.1155/2016/6403938
Physiological and Biochemical Changes in Moth Bean (Vigna aconitifolia L.) under Cadmium Stress
Moth bean (Vigna aconitifolia L.), a drought resistant legume, possesses high nutritional value. Cadmium (Cd) is a nonessential and the most toxic heavy metal in plants. The present study was to test the hypothesis of whether moth bean being a drought resistant legume can withstand the cadmium stress. Ten-day-old moth bean seedlings were subjected to cadmium stress and investigated for a period of 15 days every 3-day intervals. Cadmium quantification in moth bean tissues suggests root accumulation and translocation to aerial parts in a concentration dependent manner. Results of physiological and biochemical studies revealed that cadmium has affected the growth parameters like shoot and root lengths and tissue dry weights. Significant alternations in relative water content and cell membrane stability were observed in stressed seedlings. Similarly superoxide radical, lipoxygenase activity, membrane lipid peroxidation products, protein carbonyls, and reduced glutathione and nonprotein thiols were found increased in stressed seedlings compared to controls. However, hydrogen peroxide and ascorbic acid levels were not altered significantly in both stressed and control seedlings. Cadmium translocation ability from roots to aerial parts and elevated levels of nonenzymatic antioxidants in stressed seedlings suggest the cadmium stress withstanding ability of moth bean.
Heavy metals are of great environmental and human health concerns due to their widespread occurrence, persistence in ecosystems, and toxic properties . Agricultural crops differ widely in their tolerance to toxic metals. Tolerance of plants to heavy metals is genetically determined , so that identification of plant genotypes differing in resistance to heavy metals is a promising approach, not only for studying mechanisms protecting plants against toxic metals, but also for the selection of those adapted to production in presence of heavy metals in contaminated soils .
Cadmium (Cd) is highly toxic to all living organisms and is one of the most toxic heavy metals in plants due to its high solubility in water and phytotoxicity [4, 5]. Cadmium is not an essential nutrient in plants and at high concentration inhibits plant growth [6, 7]; even at relatively low concentrations it alters plant metabolism . Detrimental effects of cadmium are manifested in inhibition of photosynthesis and in oxidative stress leading to membrane damage . The intensive application of phosphate fertilizers containing traces of Cd and industrial zinc mining is expanding the soil-Cd contamination rapidly especially in the developing countries [10, 11].
Moth bean (Vigna aconitifolia L.), also called mat bean or matki bean, mout bean, or dew gram or Turkish gram, belongs to Fabaceae. Moth bean is a most popular pulse crop in India. It is cultivated for its immature pods and mature seeds and is consumed by people all around the world, especially in the developing nations [12–14] as it possesses high nutritional value and is a potential source of protein and other nutrients.
Legume crops are less tolerant to Cd toxicity than cereals and grasses and encounter strong inhibition of biomass production due to cadmium . Different degrees of tolerance to cadmium stress were observed among different genotypes of the same species [16–18]. Vigna species can grow under a wide range of climates and environments . Moth bean is a hot weather, drought resistant legume, particularly cultivated in hot, arid to semiarid regions. For optimum production it requires 24°C–32°C of temperature; however, the plant withstands daytime temperature of 45°C and does not tolerate water logging. Some degree of salinity and a wide pH range (3,5–10) are tolerated [15, 20]. Presence of cadmium in the soil decreases the growth of legumes like soybean [21, 22] and chickpea plants . However, in the pertinent literature survey on moth bean and its heavy metal stress responses, few published reports [24, 25] are available to date revealing its cadmium stress responses.
The present study is envisaged to evaluate whether the moth bean is sensitive to cadmium like other legumes or it can withstand the cadmium stress, as this plant can withstand a wide range of environmental stresses. To test this hypothesis the present study has been taken up to evaluate the cadmium sensitivity or tolerance of moth bean plant by subjecting to different concentrations of cadmium for a short duration period of 15 days. The investigations were carried out by determining the changes in physiological growth parameters like fresh and dry mass yield, cell membrane stability, and biochemical parameters including changes in the levels of antioxidant molecules, nonprotein thiols, and antioxidant enzymes and also accumulation of cadmium in plant organs.
2. Materials and Methods
2.1. Plant Material and Cadmium Treatments
Seeds of Vigna aconitifolia (moth bean) cv. PS-16 were procured from local agriculture seed stores of Davangere, Karnataka State, India. The healthy and uniform sized seeds were separated, surface-sterilized with 0.5% sodium hypochlorite (v/v in sterile distilled water) solution imbibed in sterile double distilled water for 48 hrs in the dark, and germinated on Whatman No. 1 filter paper moistened with sterile distilled water. Two-day-old sprouts were sown in germination trays filled with cleaned and dried sand for further growth, watered daily with 1x Hoagland nutrient solution  for 10 days. From the 11th day considered as “0” day, cadmium (Cd) in the form of cadmium chloride (CdCl2) was supplemented in Hoagland nutrient solution at different concentrations, 10 μM, 50 μM, 100 μM, 200 μM, and 500 μM; simultaneously control group of seedlings were also maintained. Stress treatments were continued for 15 days, seedlings were removed from the individual treatment trays every 3-day intervals, and the experiments were conducted according to simple randomized block design. Experiments were conducted in two sets; for each set of experiments 20 healthy seedlings were taken at random, and roots and upper second fully expanded leaves were separated and subjected to physiological and biochemical analysis.
2.2. Plant Growth Measurements
Plant growth was assessed by measuring the shoot and root lengths and fresh (FW) and dry weights (DW) of the plants from individual treatment group. For the dry weight determination, samples were oven-dried at 80°C for 15 min and then vacuum-dried at 40°C to constant weight and then dry weights (DW) were recorded.
2.3. Cadmium (Cd) Uptake
Amount of cadmium (Cd) in 15-day stressed plant tissues was quantified by atomic absorption spectrometer analysis. Briefly, leaves and roots were separated from the plants, washed in deionized water for 2 min, and air-dried and further drying was carried out in a microwave station at 80°C for 2 days. Dried plant tissue was ground into fine powder; 1 g of tissue powder was digested into 3 parts of di-acid mixture containing 1 M HNO3 and 1 M HCl (3 : 1 ratio) at 60°C. Cadmium content was determined by atomic absorption spectrometer (Chemito-AA201) analysis using cadmium standard (Sigma-Aldrich, St. Louis).
2.4. Relative Water Content (RWC)
Relative water content (RWC) was determined in fresh leaf discs that were weighed and immediately floated on double distilled water in petri dishes to saturate for the next 24 h, in the dark. The adhering water was blotted and turgor weights were recorded. Subsequently dry mass was obtained after dehydrating at 70°C for 48 h. Relative water content was calculated using the following formula:
2.5. Cell Membrane Stability (CMS)
Cell membrane stability (% of injury) was determined by electrical conductivity measurements  in control and cadmium stressed leaf discs using a conductivity meter (Systronics, India). Subsequently, the leaf tissues were killed by autoclaving at 121°C and 1.06 kg cm−2 pressure for 15 min and electrical conductivities were measured again to determine the total electrolyte concentrations.
The CMS index (% of injury) of the leaf tissues was determined using the following equation:where and represent the electrolyte concentrations measured after incubating at 10°C for 24 h and and represent the total electrolyte concentration measured after autoclaving the leaf tissues of both treatments and controls, respectively.
2.6. Estimation of Active Oxygen Species (AOS) Levels
The amount of superoxide radicals () was estimated by determining the amount of nitrite formed from hydroxylamine  in the fresh leaves and root extracts prepared separately into 65 mM phosphate buffer (pH 7.8) and supernatants were used for the estimation of superoxide. The reaction mixture contained 0.9 mL of 65 mM phosphate buffer (pH 7.8) and 10 mM hydroxylamine hydrochloride and plant tissue extract. After incubation at room temperature 17 mM sulphanilamide and 7 mM α-naphthol were added; after incubation for 5–10 min at room temperature, diethyl ether was added and absorbance of supernatants was recorded at 530 nm. Amount of nitrite formed was determined with the help of a standard curve established with standard source of and expressed as μM g−1 FW of tissue. The amount of total hydrogen peroxide (H2O2) was estimated in the fresh leaves and roots by ferrithiocyanate method . Tissue homogenates were prepared into 5% trichloroacetic acid; supernatants were used immediately for the total peroxide estimation. The reaction mixture contained tissue extract, 50% trichloroacetic acid, and 10 mM ferrous ammonium sulphate, and colour was developed with 2.5 M potassium thiocyanate. Absorbance of reaction mixture was measured at 480 nm and amount of total H2O2 was calculated using the extinction coefficient 0.28 μmol−1 cm−1.
2.7. Assay of Lipoxygenase (LOX) Activity (EC 22.214.171.124)
Lipoxygenase (EC 126.96.36.199) assay was carried out in the fresh leaf and root extracts (500 mg) as described by . Tissue homogenates were prepared separately into 3 mL of 0.2 M borate buffer (pH 8.0) and supernatants used for the lipoxygenase assay. The assay mixture contained 0.2 M boric acid buffer (pH 10.0) and plant extract and linoleic acid as a substrate in a final volume of 3 mL. The reaction was carried out at 30°C for 4 min and absorption of reaction mixture was measured at 234 nm. Molar absorption coefficient of 25000 M−1 cm−1 was used for determining the concentration of the hydroperoxides produced and lipoxygenase activity was expressed as absorbance increase (DA234) mg−1 of protein min−1.
2.8. Membrane Lipid Peroxidation and Protein Oxidation Products
Membrane lipid peroxidation in leaves and roots was determined as malondialdehyde (MDA) equivalent products that react with thiobarbituric acid . Absorbance of reaction mixture was measured at 532 nm spectrophotometrically and concentration of MDA was calculated using extinction coefficient of 155 mM−1 cm−1 and expressed as μM−1 g fresh weight of tissue. Protein oxidation was measured in tissue extracts as the total carbonyl group content by reaction with 2,4-dinitrophenylhydrazine (DNPH) . Tissue extracts were prepared into 5 mL of 50 mM potassium phosphate buffer (pH 7.4) containing 120 mM KCl and 0.1 g PVP; aliquots of supernatants containing at least 0.5 mg protein were incubated with 0.03% Triton X-100 and 1% streptomycin sulphate for 15 min to remove the nucleic acids. After centrifugation, equal volumes of supernatant and 10 mM DNPH in 2 M HCl were mixed with each other, after incubation for 1 h, proteins were precipitated with 20% TCA (w/v), precipitate was washed three times with ethanol : ethylacetate (1 : 1) and dissolved into 6 M guanidine hydrochloride in 20 mM potassium phosphate buffer, pH was adjusted to 2.3 with trifluoroacetic acid, and absorption was measured at 380 nm. Carbonyl group content was calculated using a molar absorption coefficient of 22,000 M−1 cm−1 and expressed as μM g−1 FW.
2.9. Estimation of Nonenzymic Antioxidants
Total ascorbate was determined according to the modified procedure of  in fresh leaf and root tissue homogenates prepared into 5% metaphosphoric acid and dithiothreitol (for reducing dehydroascorbate to ASC) using 5% (w/v) -ethylmaleimide and 4% (w/v) 2,2′-bipyridyl. Absorbance of the reaction was measured spectrophotometrically at 525 nm and using standard ascorbate calibration, tissue total ascorbate was determined and expressed as mg−1 g FW. Reduced glutathione content was determined using 5,5′-dithiobis-2-nitro benzoic acid (DTNB) reagent in fresh leaf and root extracts prepared separately into 5% sulfosalicylic acid . Absorbance of the reaction was measured spectrophotometrically at 412 nm and the amount of glutathione was determined using a standard calibration curve constructed with reduced glutathione (GSH). Similarly, nonprotein thiols (NPSH) were determined using DTNB reagent  in fresh leaf and root tissue homogenates prepared separately into 1 M HCl containing 1 mM EDTA. Absorbance of reaction mixture was measured at 412 nm and the amount of nonprotein thiols was quantified using a reduced glutathione (GSH) calibration curve. Free cysteine content was measured using acetic acid-ninhydrin reagent in the fresh leaf and root tissue extracts prepared into 5% perchloric acid . Absorbance was read at 560 nm and the amount of cysteine was quantified using a calibration curve prepared with cysteine standard.
2.10. Statistical Analysis of the Data
Data from 3 trials of each experiment with at least 10 plants (tissue samples from same group) for each trial were statistically analyzed (one-way ANOVA) using GraphPad Prism software (version 3.0). Unless otherwise stated elsewhere, average values (mean) and standard errors (SE) were determined, means were compared with Dunnett’s test (at 5% level of significance), and data were represented as mean ± SE.
3.1. Seed Germination and Seedling Growth
Seed germination was achieved within 2-3 days and the germination percentage was determined to be between 97 and 99%. Seedlings attained nearly 6.5–8.2 cms of height in 10 days on 1x Hoagland’s nutrient solution.
3.2. Plant Growth Measurements
Shoot lengths and root lengths of the Cd stressed seedlings were found to be slightly affected but no significant changes were observed with the increase in stress intensity (concentration of Cd and duration of exposure) and duration compared to respective control group of plants (Figure 1). Similarly, a slight reduction in the dry weights of tissue mass in cadmium stressed plants was observed at all the concentrations of cadmium and duration (Figure 2).
3.3. Cadmium Accumulation in Tissues
Cadmium content was found to be significantly increased in both leaves and roots of stressed plants compared to respective controls. Tissue accumulation was in a concentration dependent manner supplemented in the medium. Higher amounts of cadmium were determined in leaves than in roots at 10 μM concentration supplemented in the medium, higher amounts in roots than in leaves at 200 μM concentration supplemented in the medium, and similar amounts in both leaves and roots at 500 μM concentration supplemented in the medium (Figure 3).
3.4. Relative Water Content (RWC)
Relative water content (RWC) was determined in fresh leaves and roots. Significant differences in relative water content were observed between the controls and cadmium stressed plants after 15 days of cadmium exposure. In the cadmium stressed plants nearly 67% decrease in RWC was observed in leaves, while in roots the decrease in RWC was nearly 56% (Figure 4) compared to control group of plants.
3.5. Cell Membrane Stability
Cell membrane stability or membrane injury (%) was assessed by an indirect measurement of electrolyte leakage of the leaf discs. Significant decrease in cell membrane stability was observed in stressed plant leaves compared to control group leaves. There was an inverse relationship observed between the intensity of cadmium stress and cell membrane stability. Greater than or equal to 100% injury was observed only at 500 μM cadmium stress, while at other concentrations of cadmium, the % of injury was observed to be less than 100% (Figure 5).
3.6. Changes in Active Oxygen Species (AOS) Levels
Cadmium stress induced an increase in superoxide radical () production in both leaf and root tissues, compared to controls with higher levels noticed in the stressed tissues. The levels of superoxide radical were found to be up to 4-fold higher in roots than in leaves. Cadmium at 500 μM concentration induced about 4.0-fold increase in production in leaves and about 6.2-fold increase in roots at the same concentration (Figures 6(a) and 6(b)). However, a consistent increase in hydrogen peroxide production was observed in both leaves and roots of stressed plants compared to controls, with higher amounts determined in leaves than in roots (Figures 6(c) and 6(d)).
3.7. Changes in Membrane Lipid and Protein Oxidation Products
Cadmium stress caused a significant increase in the amount of lipid peroxidation in leaves and roots at all the concentrations, as measured in terms of thiobarbituric acid reactive oxygen species (TBARS) with highest amount of TBARS (MDA equivalent product); concentration was recorded in 500 μM cadmium exposed plant tissues. Compared to controls about 3-fold increase was observed in leaves and roots by day 15 (Figures 7(a) and 7(b)). Similarly, cadmium stress significantly increased carbonyl groups content in leaves and roots compared to control group of plants, with higher values recorded in leaves than in roots (Figures 7(c) and 7(d)).
3.8. Changes in Lipoxygenase (LOX) Activity (EC 188.8.131.52)
Lipoxygenase activity was found to be increased in both leaves and roots of stressed plants compared to respective controls. The increase was recorded to be consistent with the intensity of cadmium stress and duration, with higher levels in leaves than in roots (Figure 8).
3.9. Changes in Nonenzymatic Antioxidants
Ascorbic acid (AsA) content was found to be slightly decreased in leaves of cadmium stressed plants compared to respective controls. However, a significant increase in ascorbate content was observed in roots, with higher levels recorded in 500 μM cadmium stressed plants on day 15 (Figures 9(a) and 9(c)). Glutathione (reduced) content was found to be significantly increased in leaves and roots of cadmium stressed plants, compared to respective controls, with higher levels recorded in leaves than in roots at all the concentrations of cadmium and duration (Figures 9(b) and 9(d)). Nonprotein thiols (NPSH) and cysteine content were found to be increased in leaves and roots of cadmium stressed plants, compared to respective controls at all the concentrations of cadmium and duration (Figures 10(a), 10(b), 10(c), and 10(d)).
Cadmium is one of the nonessential heavy metal pollutants, which is highly toxic to plants, because of its relatively high mobility in the soil-plant system [37, 38]. Cadmium stress impairs the physiological and biochemical processes in plants such as photosynthesis, water relations, and mineral uptake [39, 40] and also causes disruption of membrane composition and function [41, 42]. Results of the present study in cadmium stressed seedlings of V. aconitifolia revealed that cadmium stress affected shoot and root growth (Figure 1) and tissue dry matter accumulation (Figure 2). The results of cadmium quantitative analysis revealed that cadmium has accumulated in both leaves and roots on a concentration dependent manner supplemented in the medium (Figure 3), suggesting the possible translocation of cadmium from roots to aerial parts (leaves) at all the concentrations used to impose stress. Cadmium reaches the aerial parts through the xylem by easily penetrating root system of xylem [43, 44]. Cadmium quantification (Figure 3) in moth bean plants clearly suggests that leaves are also affected by cadmium stress, which might be due to the translocation of this trace metal from roots to aerial parts like shoots and leaves in moth bean plant. Cadmium affects moth bean growth and leaf pigment composition [24, 25]. The difference in cadmium uptake and tissue distribution may act as the first defense line by reducing the toxic metal level in organs or cells .
Leaf relative water content (RWC) and cell membrane stability were affected in the stressed seedlings compared to nonstressed control seedlings (Figures 4 and 5). Higher loss in RWC was observed in leaves (67%) compared to roots (56%); further this loss of RWC was observed to be dependent on severity of stress and duration of exposure of seedlings to the stress Figures 4(a) and 4(b). Similar findings were also reported in lettuce  and radish seedlings  that cadmium exposure reduced the relative water contents. Reference  demonstrated that cadmium decreases leaf conductance and interacts with stomatal regulation in a concentration dependent manner which is a manifestation of cadmium toxicity in plants that is not influenced by abscisic acid. Plant water status reflects the metabolic activity that is measured as RWC in leaves . The altered leaf RWC and cell membrane damage (% injury) confirms the manifestation of cadmium toxicity in stressed V. aconitifolia seedlings, which might be due to the elevated levels of ROS in both superoxide radicals and hydrogen peroxide (Figure 6). Published data from other plant species suggests that cadmium alters the cellular redox state, by increasing the production of reactive oxygen species (ROS) like H2O2 (hydrogen peroxide), (superoxide), and (hydroxyl radicals) [37, 50]. Accumulation of ROS such as H2O2 and in cell invariably damages the components such as DNA, protein, and lipids. Growing evidence suggests that a correlation exists between the rate of increase in H2O2 content and level of membrane lipid peroxidation as well as electrolyte leakage that are often used as biochemical markers to assess the extent of oxidative damage in plants under stress [51–53].
To this perspective, results of the present study strongly indicate the onset of cadmium-induced oxidative stress manifested as oxidative damage to membranes in the V. aconitifolia seedlings, as observed by the increase in amounts of TBARS (malondialdehyde), carbonyl content in leaves and roots, which are the products of membrane lipid peroxidation and protein oxidation, respectively (Figures 7(a), 7(b), 7(c), and 7(d)). Increase in TBARS content is assumed to be a common symptom of heavy metal stress [54–56]. Protein oxidation in stressed tissues is the most commonly occurring oxidative modification; accumulation of oxidized proteins reflects not only the rate of protein oxidation but also the rate of oxidized protein degradation [32, 57]. Increase in TBARS content could be correlated with the elevated levels of lipoxygenase activity (Figure 8). Lipoxygenases are a family of enzymes that catalyze oxygenation of polyunsaturated fatty acids (PUFAs) into lipid hydroperoxides (LOOHs) involved in stress responses . Elevation in lipoxygenase activity during altered cellular redox state was reported in other plants also [58, 59].
Nonenzymatic mechanisms of ROS detoxification can also operate during plant stress, and the main nonenzymatic antioxidants include ascorbate and glutathione (GSH), nonprotein thiols in metal stress [60–62]. After application of heavy metals, the ascorbate-glutathione cycle seems to be a mechanism of great importance in controlling the cellular redox status . It was observed during the present study that the amount of ascorbic acid (Figures 9(a) and 9(c)) remained more or less stable during the entire period of study, while glutathione (GSH) (Figures 9(b) and 9(d)), other nonprotein thiols, and cysteine concentration (Figures 10(a), 10(b), 10(c), and 10(d)) were increased in cadmium stressed V. aconitifolia seedlings with increase in Cd stress intensity. Ascorbate is a primary as well as a secondary antioxidant [58, 62, 63] that may bind metals, thereby affecting their movement across biological membranes, or may act as a reducing agent, protecting the oxidation of the mercapto (-SH) groups by contributing electron or reducing power for photosystem II . Similarly GSH is a well characterized antioxidant that plays a prominent role in defense system of plants  and plays a central role in the regeneration of ascorbate and functions as an antioxidant scavenging radical, and also GSH is the direct substrate for the synthesis of phytochelatins (PC), which chelate the metals. The nonprotein thiols (NPSH) and cysteine were measured during the present study because of their importance in polythiol synthesis (phytochelatin) and metal sequestration [65, 66].
H2O2 appears to be a central signal component of plant adaptation to both biotic and abiotic stresses . Previous studies in other plants have given several lines of evidence indicating that cadmium exposure increased the production of H2O2 [57, 67, 68]. The consistent increase of H2O2 (Figures 6(a) and 6(b)) and ascorbate contents in leaves and roots (Figures 9(a) and 9(c)) and also elevated levels of reduced glutathione (Figures 9(b) and 9(d)) and nonprotein thiols in the leaves and roots (Figures 10(a), 10(b), 10(c), and 10(d)) of V. aconitifolia seedlings with increasing intensity of cadmium stress in the present study suggest that H2O2 might have induced signal transduction cascade seedlings to the imposed cadmium stress. With the increased nonenzymatic antioxidants nonprotein thiols, and translocation of cadmium from roots to leaves, these mechanisms probably confer the cadmium stress tolerance of V. aconitifolia seedlings.
Cadmium is one of heavy metals that accumulates in the agriculture soil at toxic levels due to anthropogenic activity. In the present study it has been observed that cadmium affected the physiological and biochemical parameters in both leaves and roots by altering redox status, increased lipoxygenase activity, and affected RWC and cell membrane stability in moth bean (V. aconitifolia) seedlings. In spite of these effects of cadmium, moth bean could maintain its growth vigour by minimizing the cadmium toxicity not only by translocation of cadmium from rhizosphere to aerial parts, but also by inducing the H2O2 mediated signaling, maintaining the active operation of ascorbate-glutathione cycle and increasing the concentration of nonprotein thiols and cysteine content. In the present study, it has been demonstrated that, unlike other cadmium-sensitive legumes, moth bean is tolerant to cadmium toxicity at the levels studied. Further studies are to be carried out at cellular and molecular level to confirm the possible cadmium withstanding potential of V. aconitifolia.
The authors declare that there are no competing interests regarding the publication of this paper.
- Y. J. Cui, Y.-G. Zhu, R. H. Zhai, Y. Huang, Y. Qiu, and J. Z. Liang, “Exposure to metal mixtures and human health impacts in a contaminated area in Nanning, China,” Environment International, vol. 31, no. 6, pp. 784–790, 2005.
- L. Sanità Di Toppi and R. Gabbrielli, “Response to cadmium in higher plants,” Environmental and Experimental Botany, vol. 41, no. 2, pp. 105–130, 1999.
- F. Rivera-Becerril, C. Calantzis, K. Turnau et al., “Cadmium accumulation and buffering of cadmium-induced stress by arbuscular mycorrhiza in three Pisum sativum L. genotypes,” Journal of Experimental Botany, vol. 53, no. 371, pp. 1177–1185, 2002.
- S. Clemens, “Toxic metal accumulation, responses to exposure and mechanisms of tolerance in plants,” Biochimie, vol. 88, no. 11, pp. 1707–1719, 2006.
- L. Zhao, Y.-L. Sun, S.-X. Cui et al., “Cd-induced changes in leaf proteome of the hyperaccumulator plant Phytolacca americana,” Chemosphere, vol. 85, no. 1, pp. 56–66, 2011.
- R. Anita, L. K. Chugh, V. Sawhney, and S. K. Sawhney, “Effect of cadmium and chromium on seed germination and growth of peas,” in Proceedings of Environmental Pollution, S. K. Arora, M. Singh, and R. P. Aggarwal, Eds., pp. 66–77, Haryana Agricultural University Press, Hisar, India, 1990.
- N. C. Aery and D. K. Rana, “Growth and cadmium uptake in barley under cadmium stress,” Journal of Environmental Biology, vol. 24, no. 2, pp. 117–123, 2003.
- F. Van Assche and H. Clijsters, “Effects of metals on enzyme activity in plants,” Plant, Cell & Environment, vol. 13, no. 3, pp. 195–206, 1990.
- M. N. V. Prasad, “Cadmium toxicity and tolerance in vascular plants,” Environmental and Experimental Botany, vol. 35, no. 4, pp. 525–545, 1995.
- F. Pinot, S. E. Kreps, M. Bachelet, P. Hainaut, M. Bakonyi, and B. S. Polla, “Cadmium in the environment: sources, mechanisms of biotoxicity, and biomarkers,” Reviews on Environmental Health, vol. 15, no. 3, pp. 299–323, 2000.
- K. Peng, X. Li, C. Luo, and Z. Shen, “Vegetation composition and heavy metal uptake by wild plants at three contaminated sites in Xiangxi area, China,” Journal of Environmental Science and Health-Part A Toxic/Hazardous Substances and Environmental Engineering, vol. 41, no. 1, pp. 65–76, 2006.
- S. S. Kadam and D. K. Salunkhe, “Nutritional composition, processing, and utilization of horse gram and moth bean,” Critical Reviews in Food Science and Nutrition, vol. 22, no. 1, pp. 1–26, 1985.
- R. Adsule, “Moth bean (Vigna aconitifolia (Jacq.) Marechal),” in Food and Feed from Legumes and Oil Seeds, E. Nwokolo and J. Smart, Eds., pp. 203–205, Chapman & Hall, London, UK, 1996.
- L. Bravo, P. Siddhuraju, and F. Saura-Calixto, “Effect of various processing methods on the in vitro starch digestibility and resistant starch content of indian pulses,” Journal of Agricultural and Food Chemistry, vol. 46, no. 11, pp. 4667–4674, 1998.
- M. Inouhe, S. Ninomiya, H. Tohoyama, M. Joho, and T. Murayama, “Different characteristics of roots in the cadmium-tolerance and Cd-binding complex formation between mono- and dicotyledonous plants,” Journal of Plant Research, vol. 107, no. 3, pp. 201–207, 1994.
- Y.-M. Li, R. L. Chaney, A. A. Schneiter, J. F. Miller, E. M. Elias, and J. J. Hammond, “Screening for low grain cadmium phenotypes in sunflower, durum wheat and flax,” Euphytica, vol. 94, no. 1, pp. 23–30, 1997.
- D. Ci, D. Jiang, T. Dai, Q. Jing, and W. Cao, “Variation in cadmium tolerance and accumulation and their relationship in wheat recombinant inbred lines at seedling stage,” Biological Trace Element Research, vol. 142, no. 3, pp. 807–818, 2011.
- G. Saeidi, M. Rickauer, and L. Gentzbittel, “Tolerance for cadmium pollution in a core-collection of the model legume, Medicago truncatula L. at seedling stage,” Australian Journal of Crop Science, vol. 6, no. 4, pp. 641–648, 2012.
- R. P. S. Kharb, V. P. Singh, and Y. S. Tomar, “Moth bean (Vigna aconitifolia Jacq. (Mareehal)). A review,” Forage Research Journal, vol. 13, pp. 113–132, 1987.
- A. M. A. Mazen, “Assessment of heavy metal accumulation and performance of some physiological parameters in Zea mays L. and Vicia faba L. grown on soil amended by sewage sludge resulting from sewage water treatment in the state of Qatar,” Qatar University Science Journal, vol. 15, pp. 353–359, 1995.
- D. A. Cataldo, T. R. Garland, and R. E. Wildung, “Cadmium uptake kinetics in intact soybean plants,” Plant Physiology, vol. 73, no. 3, pp. 844–848, 1983.
- R. H. Dowdy and G. E. Ham, “Soybean growth and elemental content as influenced by soil amendments of sewage sludge and heavy metals: seedling studies,” Agronomy Journal, vol. 69, no. 2, pp. 300–303, 1977.
- S. A. Hasan, B. Ali, S. Hayat, and A. Ahmad, “Cadmium-induced changes in the growth and carbonic anhydrase activity of chickpea,” Turkish Journal of Biology, vol. 31, no. 3, pp. 137–140, 2007.
- K. K. Bora, S. R. Mathur, and M. Lal, “Relative physiological and biochemical tolerance in moth bean cultivars to cadmium stress,” Current Agriculture, vol. 27, pp. 81–84, 2003.
- S. R. Mathur, K. B. Shukla, and M. K. Sharma, “Effect of cadmium on seedling growth, lipid peroxidation and photosynthetic pigments of mothbean cultivar,” International Journal of Plant Sciences, vol. 1, no. 2, pp. 200–201, 2006.
- D. R. Hoagland and D. I. Arnon, The Water Culture Method for Growing Plants without Soil, Circular 347, California Agricultural Experimental Station, University of California, Berkeley, Berkeley, Calif, USA, 1950.
- Z. Wang, Y. Zhang, Z. Huang, and L. Huang, “Antioxidative response of metal-accumulator and non-accumulator plants under cadmium stress,” Plant and Soil, vol. 310, no. 1-2, pp. 137–149, 2008.
- S. Choudhury and S. K. Panda, “Toxic effects, oxidative stress and ultrastructural changes in moss Taxithelium nepalense (Schwaegr.) broth. under chromium and lead phytotoxicity,” Water, Air, and Soil Pollution, vol. 167, no. 1–4, pp. 73–90, 2005.
- S. Sugisaka, “The occurrence of peroxide in a perennial plant,” Populs Gelrica, vol. 57, pp. 308–309, 1976.
- P. E. Skorzynska, M. Drazkiewicz, and Z. Krupa, “The activity of the antioxidative system in cadmium-treated Arabidopsis thaliana,” Biologia Plantarum, vol. 47, no. 1, pp. 71–78, 2004.
- R. L. Heath and L. Packer, “Photoperoxidation in isolated chloroplasts. I. Kinetics and stoichiometry of fatty acid peroxidation,” Archives of Biochemistry and Biophysics, vol. 125, no. 1, pp. 189–198, 1968.
- L. B. Pena, L. A. Pasquini, M. L. Tomaro, and S. M. Gallego, “Proteolytic system in sunflower (Helianthus annuus L.) leaves under cadmium stress,” Plant Science, vol. 171, no. 4, pp. 531–537, 2006.
- M. Y. Law, S. A. Charles, and B. Halliwell, “Glutathione and ascorbic acid in spinach (Spinacia oleracea) chloroplasts. The effect of hydrogen peroxide and of Paraquat,” Biochemical Journal, vol. 210, no. 3, pp. 899–903, 1983.
- O. W. Griffith, “Determination of glutathione and glutathione disulfide using glutathione reductase and 2-vinylpyridine,” Analytical Biochemistry, vol. 106, no. 1, pp. 207–212, 1980.
- S. S. Sharma, S. Kaul, A. Metwally, K. C. Goyal, I. Finkemeier, and K.-J. Dietz, “Cadmium toxicity to barley (Hordeum vulgare) as affected by varying Fe nutritional status,” Plant Science, vol. 166, no. 5, pp. 1287–1295, 2004.
- M. K. Gaitonde, “A spectrophotometric method for the direct determination of cysteine in the presence of other naturally occurring amino acids,” Biochemical Journal, vol. 104, no. 2, pp. 627–633, 1967.
- M. P. Benavides, S. M. Gallego, and M. L. Tomaro, “Cadmium toxicity in plants,” Brazilian Journal of Plant Physiology, vol. 17, no. 1, pp. 21–34, 2005.
- M. D. Groppa, M. P. Ianuzzo, E. P. Rosales, S. C. Vázquez, and M. P. Benavides, “Cadmium modulates NADPH oxidase activity and expression in sunflower leaves,” Biologia Plantarum, vol. 56, no. 1, pp. 167–171, 2012.
- U. J. López-Chuken and S. D. Young, “Modelling sulphate-enhanced cadmium uptake by Zea mays from nutrient solution under conditions of constant free Cd2+ ion activity,” Journal of Environmental Sciences, vol. 22, no. 7, pp. 1080–1085, 2010.
- S. S. Gill, N. A. Khan, and N. Tuteja, “Cadmium at high dose perturbs growth, photosynthesis and nitrogen metabolism while at low dose it up regulates sulfur assimilation and antioxidant machinery in garden cress (Lepidium sativum L.),” Plant Science, vol. 182, no. 1, pp. 112–120, 2012.
- P. L. Gratão, C. C. Monteiro, M. L. Rossi et al., “Differential ultrastructural changes in tomato hormonal mutants exposed to cadmium,” Environmental and Experimental Botany, vol. 67, no. 2, pp. 387–394, 2009.
- S. M. Gallego, L. B. Pena, R. A. Barcia et al., “Unravelling cadmium toxicity and tolerance in plants: insight into regulatory mechanisms,” Environmental and Experimental Botany, vol. 83, pp. 33–46, 2012.
- D. E. Salt and W. E. Rauser, “MgATP-dependent transport of phytochelatins across the tonoplast of oat roots,” Plant Physiology, vol. 107, no. 4, pp. 1293–1301, 1995.
- M. G. Yang, X. Y. Lin, and X. E. Yang, “Impact of Cd on growth and nutrient accumulation of different plant species,” Chinese Journal of Applied Ecology, vol. 9, no. 1, pp. 89–94, 1998.
- F.-B. Wu, J. Dong, Q. Q. Qiong, and G.-P. Zhang, “Subcellular distribution and chemical form of Cd and Cd-Zn interaction in different barley genotypes,” Chemosphere, vol. 60, no. 10, pp. 1437–1446, 2005.
- G. Costa, J. Michaut, and J. Morel, “Influence of cadmium on water relations and gas exchanges, in phosphorus deficient Lupinus albus L.,” Plant Physiology and Biochemistry, vol. 32, no. 1, pp. 105–114, 1994.
- G. Costa and J. L. Morel, “Water relations, gas exchange and amino acid content in Cd-treated lettuce,” Plant Physiology and Biochemistry, vol. 32, no. 4, pp. 561–570, 1994.
- L. Perfus-Barbeoch, N. Leonhardt, A. Vavasseur, and C. Forestier, “Heavy metal toxicity: cadmium permeates through calcium channels and disturbs the plant water status,” The Plant Journal, vol. 32, no. 4, pp. 539–548, 2002.
- D. J. Flower and M. M. Ludlow, “Contribution of osmotic adjustment to the dehydration tolerance of water-stressed pigeon pea (Cajanus cajan (L.) millsp.) leaves,” Plant, Cell and Environment, vol. 9, no. 1, pp. 33–40, 1986.
- M. C. Romero-Puertas, M. Rodríguez-serrano, F. J. Corpas, M. Gómez, L. A. del Río, and L. M. Sandalio, “Cadmium-induced subcellular accumulation of O2·− and H2O2 in pea leaves,” Plant, Cell & Environment, vol. 27, no. 9, pp. 1122–1134, 2004.
- E. Bandeoğlu, F. Eyidoğan, M. Yücel, and H. A. Öktem, “Antioxidant responses of shoots and roots of lentil to NaCl-salinity stress,” Plant Growth Regulation, vol. 42, no. 1, pp. 69–77, 2004.
- E. López, C. Arce, M. J. Oset-Gasque, S. Cañadas, and M. P. González, “Cadmium induces reactive oxygen species generation and lipid peroxidation in cortical neurons in culture,” Free Radical Biology and Medicine, vol. 40, no. 6, pp. 940–951, 2006.
- S. K. Tian, L. L. Lu, X. E. Yang, H. G. Huang, K. Wang, and P. H. Brown, “Root adaptations to cadmium-induced oxidative stress contribute to Cd tolerance in the hyperaccumulator Sedum alfredii,” Biologia Plantarum, vol. 56, no. 2, pp. 344–350, 2012.
- N. Mallick, “Copper-induced oxidative stress in the chlorophycean microalga Chlorella vulgaris: response of the antioxidant system,” Journal of Plant Physiology, vol. 161, no. 5, pp. 591–597, 2004.
- A. M. Reddy, S. G. Kumar, G. Jyothsnakumari, S. Thimmanaik, and C. Sudhakar, “Lead induced changes in antioxidant metabolism of horsegram (Macrotyloma uniflorum (Lam.) Verdc.) and bengalgram (Cicer arietinum L.),” Chemosphere, vol. 60, no. 1, pp. 97–104, 2005.
- S. Singh, S. Eapen, and S. F. D'Souza, “Cadmium accumulation and its influence on lipid peroxidation and antioxidative system in an aquatic plant, Bacopa monnieri L.,” Chemosphere, vol. 62, no. 2, pp. 233–246, 2006.
- L. Garnier, F. Simon-Plas, P. Thuleau et al., “Cadmium affects tobacco cells by a series of three waves of reactive oxygen species that contribute to cytotoxicity,” Plant, Cell and Environment, vol. 29, no. 10, pp. 1956–1969, 2006.
- G. Spiteller, “The relationship between changes in the cell wall, lipid peroxidation, proliferation, senescence and cell death,” Physiologia Plantarum, vol. 119, no. 1, pp. 5–18, 2003.
- E. Skórzyńska-Polit, B. Pawlikowska-Pawlęga, E. Szczuka, M. Drążkiewicz, and Z. Krupa, “The activity and localization of lipoxygenases in Arabidopsis thaliana under cadmium and copper stresses,” Plant Growth Regulation, vol. 48, no. 1, pp. 29–39, 2006.
- C. H. Foyer, “Ascorbic acid,” in Antioxidants in Higher Plants, R. G. Alscher and J. L. Hess, Eds., pp. 31–58, CRC Press, Boca Raton, Fla, USA, 1993.
- C. H. Foyer and G. Noctor, “Redox regulation in photosynthetic organisms: signaling, acclimation, and practical implications,” Antioxidants and Redox Signaling, vol. 11, no. 4, pp. 861–905, 2009.
- P. Ahmad, C. A. Jaleel, M. A. Salem, G. Nabi, and S. Sharma, “Roles of enzymatic and nonenzymatic antioxidants in plants during abiotic stress,” Critical Reviews in Biotechnology, vol. 30, no. 3, pp. 161–175, 2010.
- A. Cuypers, J. Vangronsveld, and H. Clijsters, “Peroxidases in roots and primary leaves of Phaseolus vulgaris Copper and Zinc Phytotoxicity: a comparison,” Journal of Plant Physiology, vol. 159, no. 8, pp. 869–876, 2002.
- K. P. Gowrinatha and V. N. R. Rao, “Reversal of heavy metal toxicity by ascorbic acid in microalgae,” Journal of Swap-Bot, vol. 9, pp. 27–29, 1992.
- D. D. Perrin and A. E. Watt, “Complex formation of zinc and cadmium with glutathione,” Biochem Biophys Acta (BBA)—General Subjects, vol. 230, no. 1, pp. 96–104, 1971.
- M. Oven, J. E. Page, M. H. Zenk, and T. M. Kutchan, “Molecular characterization of the homo-phytochelatin synthase of soybean Glycine max: relation to phytochelatin synthase,” The Journal of Biological Chemistry, vol. 277, no. 7, pp. 4747–4754, 2002.
- Y. T. Hsu and C. H. Kao, “Toxicity in leaves of rice exposed to cadmium is due to hydrogen peroxide accumulation,” Plant and Soil, vol. 298, no. 1-2, pp. 231–241, 2007.
- K. Verma, G. S. Shekhawat, A. Sharma, S. K. Mehta, and V. Sharma, “Cadmium induced oxidative stress and changes in soluble and ionically bound cell wall peroxidase activities in roots of seedling and 3-4 leaf stage plants of Brassica juncea (L.) czern,” Plant Cell Reports, vol. 27, no. 7, pp. 1261–1269, 2008.
Copyright © 2016 Poornima D. Vijendra et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.