Open Access

Phytoremediation of cyanophos insecticide by Plantago major L. in water

Journal of Environmental Health Science and Engineering201412:38

DOI: 10.1186/2052-336X-12-38

Received: 4 October 2012

Accepted: 14 January 2014

Published: 21 January 2014

Abstract

Cyanophos is commonly used in Egypt to control various agricultural and horticultural pests. It is not easily hydrolyzed and thus they are highly persistent and accumulate in various aquatic compartments such as rivers and lakes. Such issues may be solved by phytoremediation, which is the use of plants for the cleanup of pollutants. Here, we tested Plantago major L. to clean water polluted with cyanophos insecticide under laboratory conditions.The biosorption capacity (KF) of cyanophos were 76.91, 26.18 and 21.09 μg/g for dry roots, fruit (seeds with shells) and leaves of the Plantago major L., respectively. Viable Plantago major L. in water significantly reduced cyanophos by 11.0% & 94.7% during 2 hours & 9 days of exposure as compared with 0.8% & 36.9% in water without the plantain. In water with plantain, cyanophos significantly accumulated in plantain roots and leaves to reach maximum levels after two and four hours of treatment, respectively. After 1 day, the concentration of cyanophos decreased in roots and shoots until the end of testing. Three major degradation products were detected at roots and leaf samples. Here we demonstrate that plantago major L. removes efficiently cyanophos residue in water and has a potential activity for pesticide phytoremediation.

Keywords

Phytoremediation Plantago major L Water Cyanophos insecticide

Background

Cyanophos (O, O-dimethyl O-4-cyanophenyl phosphorothioate) is an organophosphorus insecticide with a commercial name of Cyanox [1]. Cyanox is commonly used in Egypt to control various agricultural and horticultural insect pests such as Hemiptera of Aphididae, Coccidae, Diaspididae, Lepidoptera, etc. in various fruits and vegetables [2]. Cyanophos used in Africa to control quelea and other granivorous species that are considered pests of cereal crops [3]. Mobile ground spraying with cyanophos control quelea, as routinely practiced in Senegal during the 1995/1996 cropping season, was found to be hazardous to the environment [1]. The toxicological effect of cyanophos is the inhibition of acetylcholine esterase activity [2]. Cyanophos is not easily hydrolyzed and highly persistent and accumulate in various aquatic compartments such as rivers and lakes [4]. Desmethyl-cyanophos, 4-Cyanophenol and desmethyl-cyanophos oxon are degradation products of cyanophos in soil [5]. All conventional methods for the removal of pesticides are found to be either uneconomical or insufficient [6]. Therefore, it becomes essential to search for effective and economical alternative method to overcome the constraints of convention methods. Biological method such as biosorption is an attractive and promising alternative which accumulate organic and inorganic matter including metal, dyes, phenols and pesticides and offers potential advantages such as low operating cost, minimization of chemical or biological sludge [7]. Several researchers reported on biosorption uptake of phenols, dyes and pesticides by biosorption [8].

Phytoremediation is an accumulation of plant-associated processes which include biotransformation, phytoaccumulation, phytoextraction, phytovolatilization, and rhizodegradation from enhanced microbial activity in plant rhizospheres [9] and plant transformation, conjugation, and sequestration are vital tools in waste management [10]. There have been several studies focused on the phytoremediation of pesticides [11, 12]. Plant remediation of soils, sediments, and water is a cost-effective and resource-conservative approach for clean-up of contaminated sites [9]. Biosorption is one of the effective alternative methods for the removal of pesticides in contaminated water samples. Plants can accumulate or metabolize a variety of organic compounds, including, imidacloprid [13], triazophos [14], chlorpyrifos [12, 15], methyl parathion [16], and atrazine [17].

The common broad leaved plantain (Plantago major L.) is a very familiar perennial weed found anywhere by roadsides, and in meadows, cultivated fields, waste areas, and canal water. The seed and husks are in fiber expanding to become highly gelatinous when soaked in water. The methanol, ethanol and aqueous extract of Plantago major L. contained antibacterial activity against some gram negative and positive bacteria besides a weak anti-narcotic activity [18]. The encouraging results of previous studies regarding phytoremediation gained the attention of researcher to continue studies in this field. Therefore, the objective of this work was to evaluate phytoremediation by living broadleaf plantain (Plantago major L.) and non- living material from plants as cleanup methods for water contaminated with the insecticide cyanophos.

Methods

Pesticide and plant

Cyanophos (Cyanox 50% EC) 0,0-dimethyl 0-(4-cyanophenyl) phosphorothioate was obtained from the Central Agriculture Pesticide Laboratory, Agriculture Research Center, Dokki, Gaiza, Egypt.

The common broadleaf plantain (Plantago major L.) used as seedlings in Phytoremediation experimentals and adult plants in biosorption assays from meadow-land in Zagazig University, Zagazig, Sharkia governorate, Egypt.

Biosorption assays

Raw agricultural solid wastes have been used as adsorbents. These materials are available in large quantities and may have potential as adsorbents due to their physico-chemical characteristics and low-cost [19]. So, Low cost materials (leaves, roots and fruits of Plantago major L.) have been tested for their ability to quickly sorb cyanophos. Adults Plantago major L. were collected with the help of fine jet of water causing minimum damage to the roots washed thoroughly with distilled water and blotted dry. Different plant parts separated manually to leaves, roots and fruits (seeds plus shells). The plantain leaves, roots and fruits (seeds plus shells) dried naturally on laboratory benches at room temperature (28–30°C) for 5 days until crisp. Sorption was measured using 0.5 g of (powder) leaves, roots and fruits (seeds plus shells), each of the broad-leaved plantain in centrifuge tube was shaken with 10 ml of the aqueous adsorbate for four hours (equilibrium concentration). Five initial concentrations (CB) were used in each case, ranging 1, 5, 10, 20 and 40 μg/mL plus water blank. After centrifugation at 2000 r.p.m. for 15 minutes, the concentrations of cyanophos in the supernatant (Ce) were determined. Aliquot (4 mL) of the supernatant was analyzed. All adsorption studies were conducted at room temperature 30°C ± 2°C and three replicated were used. The amount adsorbed (μg/g) calculated [20]. Author aimed at plotting the adsorption isotherms due to which it is possible to compare the sorption capacity of cyanophos on different adsorbents (leaves, roots and fruits of Plantago major L.). Freundlich sorption isotherm assumes that the uptake of sorbate occurs on a heterogeneous surface by multilayer sorption and can be described by the following equation: Y = KF Ce (n-1) where, KF is a Freundlich constant related to the adsorption capacity (μg/g), and n-1 is the intensity of adsorption. The values of KF and n-1can be determined from the intercept and slope, respectively of the linear plot of log y versus log Ce. The empirical Freundlich isotherm often satisfactory model of experimental data [21].

Phytoremediation assay

Whole Plantago major L. uptake experiment was performed in nutrient solution in Erlenmeyer flasks during test period from 2 h to 9 days. A whole Plantago major L. were grown in 250 ml of Hogland solution [22], containing 10 μg/ml of cyanophs in each 18 Erlenmeyer (6-periods × 3-replicates) flask 500 mL. The same number of flasks with pesticide only solution (10 μg/mL) was prepared. Three flasks were prepared as a control with a plant alone. After 2 and 4 hours, 1, 3, 6, and 9 days, three exposed and three control plants were collected. The experiment was studied at the room temperature (30 ± 2°C). Plant roots were rinsed in running tap water for 2 minutes and were blotted dry. The plants dissected into individual leaves and roots then 4 g of leaves and 2 g of roots were analyzed for the pesticide.

Residue analysis

Water samples were extracted with methylene chloride without clean up using a continuous liquid-liquid extraction [23]. Cyanophos was extracted from the root and leaf samples with acetone or water–acetone and then extracting with petroleum ether and dichloromethane. The organic phase was separated, dried, and concentrated just to dryness [24]. The organic phase was dissolved in 5 mL of hexane then cleaned up via passing through a column prewashed with 50 ml of hexane + acetone (9: 1 v/v). The column was filled with acidic alumina (5 g) + sodium sulphate (2 mg) and was eluate with 100 mL of a mixture of hexane + acetone, 9: 1 v/v [25]. The elute was evaporated to dryness and the residue was dissolved in 1.0 mL methanol and then analyzed by high-performance liquid chromatography (HPLC) with a UV-detector at 236 nm. A C18 column was used, and the mobile phase was a mixture of methanol and water (70:30, v/v). The flow rate was 1.0 mL/min. The retention time of cyanophos was 3.46 min. The metabolite 4-cyanophenol synthesized in our laboratory by hydrolyzing cyanophos with methanolic sodium hydroxide [5] and identified by HPLC with the same condition of cyanophos. Under these conditions, the retention time of 4-cyanophenol was 1.33 min.

Data analysis

The rate of degradation (K) and half-life (t 1/2) was obtained from the following Equation: The rate of degradation (K) = 2.303 × slope. Half-life (t 1/2) = 0.693/K [26].

In this study, all statistical analyses were performed with CoStat 6.311 CoHort Software. Significant differences between controls and contaminated samples were determined by the one-way ANOVA test.

Calibration curve was obtained by plotting peak areas in ‘y’ axis against concentrations of the pesticide in ‘x’ axis within the investigated range (0.18 to 12.5 μg/ml) of concentrations. Each solution was injected in triplicate. The linearity was significant with an excellent correlation coefficient of R2 = 0.994. The Limit of Detection (LOD) and Quantification (LOQ) of cyanophos were evaluated using the following equations: LOD = 3.3S0/b (3) and LOQ= 10S0/b (4) [27]. Where S0 is the standard deviation of the calibration line and b is the slope. The Limit of Detection LOD and Quantification LOQ of cyanophos in this study were found to be 0.34 μg/mL corresponding to 0.08 μg/g and 1.02 μg/mL corresponding to 0.26 μg/g, respectively. The extraction efficiency of the analytical procedure was evaluated via recovery experiments conducted in triplicate using the fortified blank Plantago major L. samples at two different concentrations, 0.2 and 0.5 μg/g. The average percentage recoveries obtained were between 93.1± 5.3%, 90.9± 4.5% and 88.3± 3.6% in water, leaves and roots, respectively.

Results and discussion

Biosorption assays

Data in Table 1 shows the biosorption capacity for cyanophos by the dry roots, fruits (seeds with shells) and leaves of Plantago major L. after four hours of exposure. The Experimental data of biosorption of cyanophos onto Plantago major L. showed that, the correlation coefficients (R2) for the Freundlich isotherm were 0.971, 0.997 and 0.921 for dry roots, leaves and fruit, representing a good fit of the observed data. Dry root biosorption KF was much higher than that of dry fruit and dry leaves at all exposure concentrations. The amount of absorption of cyanophos can be ranked as dry roots (76.91 μg/g), dry fruits (26.18 μg/g) and dry leaves (21.09 μg/g). The equilibrium constant (n-1) that related to the extent or degree of biosorption was 0.896, 0.928, and 1.04 for dry roots, dry fruit, and dry leaves of Plantago major L., respectively. The biosorption (KF) of dry roots was 3.64 and 2.93 folds higher than that of dry leaves and dry fruit, while the KF of dry fruit was 1.24 fold larger than that of dry leaves of Plantago major L. The increase of biosorption KF for dry roots than dry leaves and fruits may be due to the lipid content of plant roots, in which protein, fats, nucleic acids, cellulose tissues, and other components all contain lipophilic components which appear to be more efficiency in pesticides adsorption [28]. Hetero-geneous accumulation of pesticides in different plant parts of same crop species, which could be attributed to their diverse morphological characteristics [29]. Biosorption of similar pesticides by different biomass depended on the number of sites on the biosorbent, the accessibility of the sites, the chemical state of the site (i.e., availability) and affinity between site and xenobiotic (i.e., binding strength) [30]. Also, physico-chemical parameters, such as temperature, pH of the contact solution and conditions of the reaction, e.g. continuous or batch mode, are reported to influence the adsorption results [8]. The high level of KF suggested that the adsorption capacity of dry roots were high [21]. Freundlich model was more appropriate to describe the adsorption characteristics of trichloroethylene (TCE) onto Carbon nanotubes [31]. The larger Freundlich constant KF showed an easy uptake of phenol from aqueous solution [32]. Agricultural fibers were more efficiency in phenol removal [33]. Furthermore, wheat ash were highly effective sorbent for herbicide MCPA [34]. Accumulation of pesticides on agricultural adsorbents is generally achieved through interactions with the hydroxyl and carboxyl groups particularly abundant in polysaccharides (cellulose and hemicelluloses) and lignin, both of which constitute about 90% of dry lignocellulosic materials [35]. The biosorption capacity (KF) of dry roots of Plantago major L. was significantly higher than that of dry leaves and dry fruits at all concentrations of imidacloprid [11]. Several researchers reported on biosorption uptake of pesticides by biosorption [8, 36].
Table 1

Biosorption of cyanophos by plantago major L. on a dry weight basis after 4 hours exposure

Concentrations in water (μg/mL) ±*S.D.

Concentrations in water (μg/mL) and adsorption on a dry weight basis

Significantly

 

Roots

Leaves

Fruits (seeds with shells)

 
 

(μg/mL) ±*S.D.

(μg/g)

(μg/mL) ±*S.D.

(μg/g)

(μg/mL) ±*S.D.

(μg/g)

 

34.07 ± 1.0(a)

15.17 ± 0.67(b)

2890

11.43 ±0.5(c)

652.8

12.12 ± 0.39(c)

798.75

***

17.89 ±0.34

8.43 ± o.45(b)

946

2.23 ±0.13(c)

313.2

2.50 ±0.23(c)

384.75

***

9.10 ±0.1

4.15 ±0.13(b)

495

0.91 ±0.08(c)

182.8

0.92 ±0.03(c)

204.5

***

4.47 ± 0 .13

2.23 ± .05(b)

224

0.092 ± 0.01(c)

87.56

0.074 ± 0.002(c)

109.9

***

0.91 ±0.01

Undetected

90.0

Undetected

18.2

Undetected

22.8

***

Freundlich parameters

1Kf

76.91

21.09

26.18

1an-1

0.896

0.928

1.047

1bR2

0.971

0.997

0.921

*SD, standard deviation; 1Kf, adsorption capacity (ug/g); 1an-1, intensity of adsorption; 1bR2, the correlation coefficient.

*P< 0.01, ***P< 0.001.

Uptake and distribution

Table 2 show the phytoremediation potential of Plantago major L. to remove cyanophos insecticide from contaminated water. Viable whole plant of Plantago major L. in water solution significantly reduced cyanophos residues by 11.0 & 94.7% during 2 & 216 hours of exposure periods as compared with 0.8 & 36.9% in water solution without the plantain (Table 2). The half-life value (t1/2) of cyanophos, calculated by first-order reaction, for water planted with Plantago major L. and in unplanted water was found to be 1.73 and 13.63 days, respectively (Table 2). These data demonstrated that, most of the cyanophos disappearance by a Plantago major L. may be attributed to the uptake potential and transformation or degradation by the enzyme induction capability of the plant or by microorganisms in the plant root zone. Only one of them contributes to the reduction of a contaminant or connected them [37]. The growing cells of short-rod gram-negative bacteria that isolated from the water solution containing Plantago major L. was able to induce 93.34% loss of imidacloprid as a source of both carbon and nitrogen within a short period (48 hrs) compared with 31.90% in un inoculated medium [11]. The t1/2 value of cyanophos for Nile water was found to be 7 days [38]. Under illumination, the half-life was estimated to be 2 and 4 days on soil and silica, respectively, while it was evaluated to be 120 min in acetone solution [39]. Viable whole Plantago major L. in water solution reduced imidacloprid residues by 55.81–95.17%, during 1–10 days of exposure periods compared with 13.71–61.95% in water solution without Plantago major L. [11]. The disappearance rate constant (kr) is 8.0 times greater in water solution with plantain than in the cyanophos control. The areas under the curve (AUC) represent compound concentration during the period of study. AUC in the water solution with plantain was decreased than cyanophos control (Table 2).
Table 2

Concentrations of cyanophs uptake during plantago major L

Determinations

2 hours

Days after application

  

0.17

1

3

6

9

1t1/2(Days-1)

2Kr(Day-1)

3AUCmg l1(Day-1)

In water solution

μg/mL ± *S.D

9.92 ±0.14a

9.85 ±0.1a

9.20 ±0.21a

8.59 ±0.12a

7.19 ±0.17a

6.31 ±0.1a

13.63

0.05

71.37

% loss

0.8

1.5

8.0

14.1

28.1

36.9

 

In water solution with plantain

μg/mL ± *S.D

8.55 ±0.11c

7.70 ±0.10c

4.53 ±0.1c

2.89 ±0.03c

1.13 ±0.03

0.53 ±0.02c

1.73

0.40

20.13

% loss

11.0

23.0

54.7

71.1

88.7

94.7

 

Significantly

***

***

***

***

***

***

In plantain roots

μg/g ± S.D

243.5 ±1.32a

113.49 ±1.56a

56.25 ±1.13a

49.21 ±1.24a

45.26 ±1.25a

42.11 ±1.35a

 

In plantain leaves

μg/g ± *S.D

53.63 ±0.87b

73.0 ±1.04b

40.29 ±0.67b

36.4 ±0.86b

31.62 ±0.71b

20.00 ±1.0b

 

Total uptake

297.13

186.49

96.54

85.64

76.88

30.0

Significantly

***

***

***

***

***

***

1T1/2, half-life; 2kr, disappearance rate constant; 3AUCs, areas under the curve represent compound concentration during the period of study; *SD, standard deviation

*P± 0.01, ***P± 0.001.

In water, cyanophos uptake and distribution by both roots and leaves of Plantago major L. are shown in the Table 2. Cyanophos was taken up by roots much faster than shoots. In water, Cyanophos significantly accumulated in the roots of Plantago major L. to reach the maximum levels after 2 hrs (243.5±1.32 μg/g). Afterwards concentration decrease gradually throughout the test. Cyanophos translocated into the leaves (53.63±0.87 μg/g) within 2 hrs and reached the maximum after 0.17 day of exposure (73.0 ±1.04 μg/g), then decreased until the end of testing (Table 2). It has been observed that roots were important in accumulating compounds due to their direct exposure of toxic chemicals with underground parts, and transporting the compounds to above-ground organs (shoots) [40]. The uptake and translocation of organic compounds are dependent on hydrophobicity (lipophilicity), solubility, polarity, molecular weight, plant species and environmental factors [41]. Lipophilicity is the most important property of a chemical in determining its movement into and within a plant and is related to the n-octanol/water partition coefficient (Kow) value. For uptake, log Kow must be typically between 0.5 and 3.0. Compounds with larger log Kow are hydrophobic and may adsorb strongly onto roots. With smaller log Kow are too hydrophilic to pass through cell membrane [42]. Cyanophos is a moderately hydrophobic compound (log Kow 2.65) and is likely to partially adsorb onto roots or be taken up by roots and move across cell membranes to reach the aboveground portion of plants. Foliar uptake directly into the aboveground portion of plants is an important route compared with root uptake, especially for volatile and semi-volatile compounds [43]. However, the volatility of cyanophos was found to be substantially high, V.p. 105 at 20°C [2] therefore, cyanophos residue accumulated in the roots and moved to the aerial part of the plant material. Also, potential uptakes of organic contamination are influenced by evapotranspiration [44]. Pesticide remediation is enhanced in emergent and floating vegetation by high transpiration rates and lipids associated with plant cuticles [44], therefore higher transpiration rates of emergent vegetation could be expected to increase volatilization of pesticide metabolites resulting in greater remediation capabilities. Most of the transport of pesticides occurs in aquatic plants via rhizomes. Once moved inside the plants, pesticides are acropetally distributed from the roots primarily into the leaves and lost via diffusion if volatile [45]. The rooted portion of Acorus. gramineus plays an import role in the total pesticide-uptake due to the relatively high weight-based accumulation of pesticides and its contribution to organ biomass proportion [46]. Among the organophosphorus pesticide, malathion was an efficient accumulator (81%) in the roots, followed by fenitrothion (76%), diazinon (65%), and parathion (64%), respectively. In water solution with Plantago major L., imidacloprid significantly accumulated in plantain roots, leaves and fruits to reach the maximum levels after 6, 1 and 3 days of treatment, respectively. The maximum levels were 15.74, 37.21, and 5.74 μg/g, respectively. These values were decreased to 6.95, 1.46, and 0.12 μg/g after 10 days of treatment [11]. Plants can accumulate or metabolize a variety of organic compounds, including, imidacloprid [13], triazophos [14], chlorpyrifos [12, 15], methyl parathion [16] and atrazine [17, 47]. Several grass species were utilized to create grass waterways and buffer strips to contain polluted surface waters [4850]. The disappearance of cyanophos was coupled with the appearance of the metabolites in the roots and the leaves of Plantago major L. Three major degradation products were detected at roots and leaf samples (Figures 1, 2 and 3). Two major metabolites 4-cyanophenol and product with retention time (R.T.) of 1.43 minutes were detected in roots and leaves, respectively. One other metabolite was detected in both roots and leaves at a R.T. of 3.05 minutes (Figures 1 and 2). In the water solution, the degradation products 4-cyanophenol and the product at a R.T. of 1.43 minutes were detected in the roots and the leaves after two hours, respectively. After1 day, the product at a R.T. of 1.43 minutes was decreased gradually in leaves until the end of sampling while the 4-cyanophenol was no detectable in roots at 6 days (Figures 1 and 2). The metabolite at a R.T. of 3.05 minutes was detected in both roots and leaves after 1 day and 2 hours, respectively. The metabolite at a R.T. of 3.05 minutes increased gradually to 6 days then decreased throughout the 9 days of exposure (Figure 1). These data indicated that Plantago major L., degrade cyanophos enzymatically in water. Cyanophos was taken up rapidly and metabolized by the root of bean in hydroponic solution [5]. After 9 days, 24% remained as unchanged and 24% as desmethyl-cyanophos via demethylation. The important metabolite 4-cyanophenol a probably occurs principally by the hydrolysis of Oxon in plants. The same author found that when cyanophos was applid to bean leaves cyanophos oxon and desmethyl-cyanophos oxon were also detected. The aerobic metabolism of cyanophos in water-sediment systems undergoes the cleavage of the P–O-methyl and P–O-aryl bonds together with the oxidation of the P=S to the oxon group and the hydrolysis of the cyanophos moiety [51]. Photo-oxidation reaction leading to the formation of the oxon derivative and the scission of the P–O bond generating 4-cyanophenol, as a major product are reported to be the main observed processes [39]. Wheat plants enhanced uptake/degradation of methyl parathion, p-nitrophenol and hydroquinone in unsterilized soil by 64.85%, 94.7% and 55.8% respectively. Methyl parathion hydrolyzes to p-nitrophenol, which is further metabolized to hydroquinone with nitrite release The enzyme p-nitrophenol 4-hydroxylase is active as evidenced by release of nitrite by leaf and root extracts and also by the appearance of hydroquinone in the reaction mixture [16]. Data also in Figures 2 and 3 clear role Plantago major L. in phytoremediation of cyanophos and degradation products in water.
Figure 1

Uptake of cyanophos degradation products in water by Plantago major L. roots and leaves.

Figure 2

HPLC Chromatogram of cyanophos and its degradation products in water by Plantago major L. roots. Note, Retention time (R.T) of cyanophos and its degradation product 4-cyano phenol were 3.38 and 1.33 min, respectively.

Figure 3

HPLC Chromatogram of cyanophos and its products in water by Plantago major L. leaves.

Conclusions

The use of plants to detoxify contaminated water is a potentially cost-effective alternative to traditional remediation technologies. From the results of this study, it can be ended that the existence of plants increased the removal t1/2 of cyanophos in water system. Plantago major L. is able to take up cyanophos from water by roots as well as by leaves, so Plantago major L. may be used for phytoremediation of water contaminated with cyanophos insecticide.

Declarations

Acknowledgements

The author is most grateful to the laboratory staff of pesticides analysis and environmental pollution Laboratory, Plant Production Department, Faculty of Technology and Development, Zagazig University, Zagazig, Egypt for their collaboration in this research.

Authors’ Affiliations

(1)
Plant Production Department, Faculty of Technology and Development, Zagazig University

References

  1. Mullie W, Diallo A, Gadji B, Ndiaye M: Environmental hazards of mobile ground spraying with cyanophos and fenthion for Quelea control in Senegal. Ecotoxicol Environ Saf 1999, 43: 1–10. 10.1006/eesa.1998.1744View ArticleGoogle Scholar
  2. Tomlin C: The e- pesticide Manual. A world Compendium. 13 2004 edition. United kingdom: British Crop Protection Council: Farniham, surey; 2004:184.Google Scholar
  3. Bruggers RL, Elliott CCH: quelea, Africa1s Bird Pest. Oxford: Oxford University Press; 1989.Google Scholar
  4. Floesser-Mueller H, Swack W: Photochemistry of organophosphorus insecticides. Rev Environ Contam Toxicol 2001, 172: 129–228.Google Scholar
  5. Chiba M, Shlgeru K, Izuru Y: Metabolism of cyanox and surecide in bean plants and degradation in soil. J Pesticide Sci 1976, 1: 179–191. 10.1584/jpestics.1.179View ArticleGoogle Scholar
  6. Kiso Y, Sugiura Y, Kitao T, Nishimura K: Effects of hydrophobicity and molecular size onrejection of aromatic pesticides with nanofiltration membranes. J Mem Sci 2001, 92: 1–10.View ArticleGoogle Scholar
  7. Rahman MA, Hasegawa H: Aquatic arsenic: Phytoremediation using floating macrophytes. Chemosphere 2011, 83: 633–646. 10.1016/j.chemosphere.2011.02.045View ArticleGoogle Scholar
  8. Yadamari T, Kalyan Y, Gangadhar B, Ramakrishna N: Biosorption of malathion from aqueous solutions using herbal leaves powder. AJAC 2011, 2: 37–45. 10.4236/ajac.2011.228122View ArticleGoogle Scholar
  9. Susarla S, Medina VF, McCutcheon SC: Phytoremediation: an ecological solution to organic chemical contamination. Ecol Eng 2002, 18: 647–658. 10.1016/S0925-8574(02)00026-5View ArticleGoogle Scholar
  10. McCutcheon SC, Schnoor JL: Phytoremediation, Transformation and Control of Contaminants. Hoboken, New Jersey: John Wiley and Sons; 2003:1–58.Google Scholar
  11. Romeh A: Phytoremediation of water and soil contaminated with imidacloprid pesticide by plantago major , L. Int J Phytoremediation 2010, 12: 188–199.View ArticleGoogle Scholar
  12. Prasertsup P, Naiyanan A: Removal of chlorpyrifos by water lettuce ( Pistia stratiotes l.) and duckweed ( Lemna minor l.). Int J Phytoremediation 2011, 13: 383–395. 10.1080/15226514.2010.495145View ArticleGoogle Scholar
  13. Byrne FJ, Toscano NC: Uptake and persistence of imidacloprid in grapevines treated by chemigation. Crop Prot 2006, 25: 831–834. 10.1016/j.cropro.2005.11.004View ArticleGoogle Scholar
  14. Cheng S, Jin X, Huiping X, Liping Z, Zhenbin W: Phytoremediation of triazophos by canna indicalinn. in a hydroponic system. Int J Phytoremediation 2007, 9: 453–463. 10.1080/15226510701709531View ArticleGoogle Scholar
  15. Romeh AA, Hendawi MY: Chlorpyrifos insecticide uptake by plantain from polluted water and soil. Environ Chem Lett 2013, 11: 163–170. 10.1007/s10311-012-0392-0View ArticleGoogle Scholar
  16. Khan NU, Bhavya V, Nazeeb I, Paddu KS: Phytoremediation using an indigenous crop plant (wheat): the uptake of methyl parathion and metabolism of p-nitrophenol. Indian J Sci Technol 2011, 4: 1661–1667.Google Scholar
  17. Wang Q, Wei Z, Cui. Bo X: Phytoremediation of atrazine by three emergent hydrophytes in a hydroponic system. Water Sci Technol 2012, 66: 282–1288.Google Scholar
  18. Sharifa AA, Neoh YL, Iswadi MI, Khairul O, Abdul Halim MM, Jamaludin Mohamed A, Hing HL: Effects of methanol, ethanol and aqueous extract of plantago major on gram positive bacteria, gram negative bacteria and yeast. Ann Microsc 2008, 8: 42–44.Google Scholar
  19. Ahmaruzzaman MD: Adsorption of phenolic compounds on low-cost adsorbents: a review. Adv Colloid Interface Sci 2008, 143: 48–67. 10.1016/j.cis.2008.07.002View ArticleGoogle Scholar
  20. Felsot A, Dahm A: Sorption of organophosphorus and carbamate insecticides by soil. J Agric Foods Chem 1979, 27: 557–563. 10.1021/jf60223a013View ArticleGoogle Scholar
  21. Tebbutt THY: Principles of Water Quality Control. 3rd edition. Oxford, England: Pergamon Press; 1991:219–220.Google Scholar
  22. Wang W: Toxicity tests of aquatic pollutants by using common duckweed. Environ Poll 1986, 11: 1–14.View ArticleGoogle Scholar
  23. EL-Sheamy MK, Hussein MZ, El-Sheak AA, Khater AA: Residue behavior of certain organophosphorus and Carbamate insecticides in water and fish. Egypt J Appl Sci 1991, 6: 94–102.Google Scholar
  24. Luke MA, Froberg JE, Doose GM, Masumato HT: Improved multiresidue gas chromatographic determination of orgonophosphorus, orgononitrogen and orgonohalogene pesticides in procedure, using flame photometric and electrolytic conductivity detectors. Journal of AOAC 1981, 64: 1187–1195.Google Scholar
  25. Zaalok A, Sherif A: Combined effect of applied equipment and formulation of pesticide on spray and dust drift in relation to harmful effects for some non-target organisms. Agric Biol J N Am 2011, 2: 1059–1065.View ArticleGoogle Scholar
  26. Gomaa EA, Belal MH: Determination of dimethoate residues in some vegetables and cotton plant. Zagazig J Agric Res 1975, 2: 215–221.Google Scholar
  27. Thomsen V, Schatzlein D, Mercuro D: Limits of detection in spectroscopy. Spectroscopy 2003, 18: 112–114.Google Scholar
  28. Kang F, Dongsheng C, Yanzheng G, Yi Z: Distribution of polycyclic aromatic hydrocarbons in subcellular root tissues of ryegrass ( Lolium multiflorum Lam.). BMC Plant Biol 2010, 10: 210. 10.1186/1471-2229-10-210View ArticleGoogle Scholar
  29. Mohamed AE, Rashed MN: Assessment of essential and toxic elements in some kinds of vegetables. Ecotoxicol Environ Saf, Environ Res 2003, 55: 251–260. 10.1016/S0147-6513(03)00026-5View ArticleGoogle Scholar
  30. Ghaly AE, Snow Kamal AM: Kinetics of manganese uptake by wetland plants. American Appl Sci 2008, 5: 1415–1423. 10.3844/ajassp.2008.1415.1423View ArticleGoogle Scholar
  31. Naghizadeh A, Nasseri S, Nazmara S: Removal of trichloroethylene from water by adsorption on to multiwall carbon nanotubes. Iran J Environ Health Sci Eng 2011, 8: 317–324.Google Scholar
  32. Rengaraj S, Seuny H, Sivabalan MR: Agricultural solid waste for the removal of organics: adsorption of phenol from water and wastewater by Palm seed coat activated carbon. Waste Manag 2002, 22: 543–548. 10.1016/S0956-053X(01)00016-2View ArticleGoogle Scholar
  33. Mahvi AH: Application of agricultural fibers in pollution removal from aqueous solution. Int J Environ Sci, Tech 2008, 5: 275–285. 10.1007/BF03326022View ArticleGoogle Scholar
  34. Hiller E, Fargas A, Zemanova L, Barta M: Influence of wheat Ash on the MCPA immobilization in agricultural soils. Bull Environ Contam Toxicol 2008, 81: 285–288. 10.1007/s00128-008-9400-2View ArticleGoogle Scholar
  35. Ofomaja AE: Kinetic study and sorption mechanism of methylene blue and methyl violet onto mansonia wood sawdust. Chem Eng J 2008, 143: 85–95. 10.1016/j.cej.2007.12.019View ArticleGoogle Scholar
  36. Senthilkumaar S, Krishna SK, Kalaamani P, Subburamaan CV, Ganapathi N: Adsorption of organophosphorous pesticide from aqueous solution using “waste” jute fiber carbon. Mod Appl Sci 2010, 4: 67–83.View ArticleGoogle Scholar
  37. Derbalaha AS, Belalb EB: Biodegradation kinetics of cymoxanil in aquatic system. Chem Ecol 2008, 24: 169–180. 10.1080/02757540802032173View ArticleGoogle Scholar
  38. Al-Makkawy HK, Madbouly MD: Persistence and accumulation of some organic insecticides in Nile water and fish. Resour Conserv Recycl 1999, 27: 105–115. 10.1016/S0921-3449(98)00090-1View ArticleGoogle Scholar
  39. Mikami N, Ohkawa H, Miyamoto J: Photodecomposition of surecide (O-ethyl O-4-cyanophenyl phenylphophonothioate) and Cyanox (O, O-dimethyl O-4-cyanophenyl phosphorothioate). J Pestic Sci 1976, 1: 273–281. 10.1584/jpestics.1.273View ArticleGoogle Scholar
  40. Azmat R, Haider S, Riaz M: An inverse relation between Pb 2+ and Ca 2+ ions accumulation in Phaseolus mungo and Lens culinaris under Pb stress. Pak J Bot 2009, 41: 2289–2295.Google Scholar
  41. Turgut C: Uptake and modeling of pesticides by roots and shoots of parrot feather ( myriophyllum aquaticum ). Environ Sci Pollut Res 2005, 12: 342–346. 10.1065/espr2005.05.256View ArticleGoogle Scholar
  42. Bouldin JL, Farris JL, Moore MT, Smith SJ, Cooper M: Hydroponic uptake of atrazine and lambda-cyhalothrin in Juncus effusus and Ludwigia peploides . Chemosphere 2006, 65: 1049–1057. 10.1016/j.chemosphere.2006.03.031View ArticleGoogle Scholar
  43. Wang MJ, Jones KC: Behaviour and fate of chlorobenzenes (CBs) introduced into soil–plant systems by sewage sludge application: a review. Chemosphere 1994, 28: 1325–1360. 10.1016/0045-6535(94)90077-9View ArticleGoogle Scholar
  44. Chefetz B: Sorption of phenathrene and atrazine by plant cuticular fractions. Environ Toxicol Chem 2003, 22(10):2492–2498. 10.1897/02-461View ArticleGoogle Scholar
  45. Karthikeyan R, Lawrence D, Larry E, Kassim A, Peter A, Philip L, Stacy L, Asil A: Potential for plant-based remediation of pesticide-contaminated soil and water using nontarget plants such as trees, shrubs, and grasses. Crit Rev Plant Sci 2004, 23: 91–101. 10.1080/07352680490273518View ArticleGoogle Scholar
  46. Chuluun B, Janjit I, Jae Seong R: Phytoremediation of organophosphorus and organochlorine pesticides by acorus gramineus . Environ Eng Res 2009, 14: 226–236. 10.4491/eer.2009.14.4.226View ArticleGoogle Scholar
  47. Ibrahim S, Abdel Lateef M, Khalifa H, Abdel Monem A: Phytoremediation of atrazine-contaminated soil using Zea mays (maize). Ann Agric Sci 2013, 58: 69–75. 10.1016/j.aoas.2013.01.010Google Scholar
  48. Rankins A, Shaw R, Boyette M: Perennial grass filter strips for reducing herbicide losses in runoff. Weed Sci 2001, 49: 647–651. 10.1614/0043-1745(2001)049[0647:PGFSFR]2.0.CO;2View ArticleGoogle Scholar
  49. Angier T, McCarty W, Rice P, Bialek K: Influence of a riparian wetland on nitrate and herbicides exported from an agricultural field. J Agric Food Chem 2002, 50: 4424–4429. 10.1021/jf011057nView ArticleGoogle Scholar
  50. Zhao S, Arthur L, Coats R: The use of native prairie grasses to degrade atrazine and metolachlor in soil. In Environmental Fate and Effects of Pesticides. Edited by: Coats JR, Yamamoto H. Washington, DC: ACS Symposium Series 853, ACS; 2003:157–167.View ArticleGoogle Scholar
  51. Kodaka R, Sugano T, Katagi T, Takimoto Y: Comparative metabolism of organophosphorus pesticides in water-sediment systems. J Pestic Sci 2003, 28: 175–182. 10.1584/jpestics.28.175View ArticleGoogle Scholar

Copyright

© Romeh; licensee BioMed Central Ltd. 2014

This article is published under license to BioMed Central Ltd. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Advertisement