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Cadmium uptake kinetics in parts of the seagrass Cymodocea nodosa at high exposure concentrations

  • 1Email author,
  • 2,
  • 1 and
  • 2
Journal of Biological Research-Thessaloniki201825:5

https://doi.org/10.1186/s40709-018-0076-4

  • Received: 22 November 2017
  • Accepted: 26 February 2018
  • Published:

Abstract

Background

Seagrass species have been recommended as biomonitors of environmental condition and as tools for phytoremediation, due to their ability to concentrate anthropogenic chemicals. This study aims to provide novel information on metal accumulation in seagrasses under laboratory conditions to support their use as a tool in the evaluation and abatement of contamination in the field. We investigated the kinetics of cadmium uptake into adult leaf blades, leaf sheaths, rhizomes and roots of Cymodocea nodosa in exposure concentrations within the range of cadmium levels in industrial wastewater (0.5–40 mg L−1).

Results

A Michaelis–Menten-type equation satisfactorily described cadmium accumulation kinetics in seagrass parts, particularly at 0.5–5 or 10 mg L−1. However, an S equation best described the uptake kinetics in rhizomes at 5 mg L−1 and roots at 10 and 20 mg L−1. Equilibrium concentration and uptake rate tended to increase with the exposure concentration, indicating that seagrass displays a remarkable accumulation capacity of cadmium and reflect high cadmium levels in the surrounding medium. Concerning leaf blades and rhizomes, the bioconcentration factor at equilibrium (range 73.3–404.3 and 14.3–86.3, respectively) was generally lower at higher exposure concentrations, indicating a gradual reduction of available binding sites. Leaf blades and roots accumulated more cadmium with higher rate than sheaths and rhizomes. Uptake kinetics in leaf blades displayed a better fit to the Michaelis–Menten-type equation than those in the remaining plant parts, particularly at 0.5–10 mg L−1. A marked variation in tissue concentrations mainly after the steady state was observed at 20 and 40 mg L−1, indicative of the stress induced on seagrass cells. The maximum concentrations observed in seagrass parts at 5 and 10 mg L−1 were comparatively higher than those previously reported for other seagrasses incubated to similar exposure concentrations.

Conclusions

Cymodocea nodosa displays a remarkable cadmium accumulation capacity and reflects high cadmium levels in the surrounding medium. Kinetic models satisfactorily describe cadmium uptake in seagrass parts, primarily in adult leaf blades, at high exposure concentrations, permitting to predict cadmium accumulation in field situations. Cymodocea nodosa appeared to be a valuable tool in the evaluation and abatement of cadmium contamination in coastal areas.

Keywords

  • Metal
  • Accumulation kinetics
  • Seagrass parts
  • Cymodocea nodosa
  • Biomonitor
  • Phytoremediation

Background

Seagrasses are present in most shallow coastal waters throughout the world, providing a multitude of ecological services and functions. Seagrass beds are highly productive ecosystems, provide habitat and nursery areas for a variety of invertebrates, fish and mammals, and enhance water quality by stabilizing sediments and removing nutrients [1]. Due to their ability to concentrate and retain non-nutrient anthropogenic chemicals in their tissues, seagrass species have been also recommended as biomonitors of environmental condition and as potent tools for phytoremediation [24]. The usefulness of seagrasses in the abatement of contamination in coastal areas is corroborated by available data suggesting that these plants appear to be relatively tolerant to many anthropogenic chemicals, particularly more tolerant than other marine flora (macroalgae, kelp, saltmarsh plants, mangroves) and macroinvertebrates [3].

With regard to trace metals, which are of greatest environmental concern [5], several studies, mainly in situ, have examined the accumulation of these contaminants in seagrasses [3, 6]. Laboratory-derived results for metal uptake kinetics are available for about ten seagrass species, for example Heterozostera tasmanica [7], Posidonia oceanica [810] and, most commonly, Zostera species [1113] and Halophila stipulacea [1417]. Metal uptake kinetics only in one or two plant parts, namely leaves or leaves and roots-rhizomes, were investigated in most of these studies [7, 1217]. Of the trace metals, cadmium, lead and zinc were more commonly used in uptake experiments [7, 9, 1114, 16]. Recently, the linkage between metal uptake into intermediate-juvenile leaf blades and toxic effects was also examined in a seagrass species (Cymodocea nodosa) [18, 19]. However, only in a few of these studies, an attempt was made to describe the uptake patterns by kinetic models [18, 19]. The fit of kinetics data to appropriate models would permit the calculation of uptake kinetic parameters and the prediction of metal accumulation in field situations. Such information could support the use of seagrasses as a tool in the evaluation of contamination in coastal areas or the clean-up of contaminated sites.

The main goal of the present study is to contribute to a better understanding of metal accumulation in seagrasses and to increase the utility of these marine plants as a potent tool in the evaluation and abatement of metal contamination in coastal areas. We assessed the capacity of various parts of Cymodocea nodosa to accumulate cadmium from the surrounding medium, as well as the speed of this process. Cadmium was chosen as a contaminant because it is considered highly toxic; it may enter the aquatic environment from various anthropogenic sources, such as zinc, copper and lead mining, various industries, nickel–cadmium batteries and phosphate fertilizers [20]. Cadmium concentrations in the range of 0.1–100 mg L−1 are typical in wastewater from several industries (chemical and metal product facilities, leather and tanning processes, electricity and gas production and sanitary industries) [21]. Cymodocea nodosa (Ucria) Ascherson along with Posidonia oceanica (L.) Delile are the most important and widespread seagrass species in the Mediterranean Sea. Cymodocea nodosa is distributed from the intertidal zone to depths of 33–35 m, and can colonize different environmental types, such as open coastal waters, estuaries and coastal lagoons; it is considered as a species with great phenotypic plasticity and a high capacity to adapt to environmental variability and thereby to colonize new substrates [2224]. Cymodocea nodosa is also considered as a good biomonitor for trace elements, with leaves being the best part for the determination of element loads [25]. In addition, microtubule integrity in leaf cells of C. nodosa is regarded as an early marker of metal-induced stress [18, 19, 26]. Under laboratory conditions, we investigated the kinetics of cadmium uptake into adult leaf blades, leaf sheaths, rhizomes and roots of Cymodocea nodosa exposed to concentrations of cadmium ranging from 0.5 to 40 mg L−1; the uptake kinetics data were fitted to different models.

Methods

Plant collection

Cymodocea nodosa was collected from the eastern coast of the Gulf of Thessaloniki, Northern Aegean Sea at the Viamyl site (site V, 40°33′N, 22°58′E). At this site, C. nodosa grows from 0.4 m to around 2 m depth, forming a continuous monospecific meadow. Leaf, rhizome and root biomass displays an annual mean value of approx. 60, 122 and 65 g dry wt m−2, respectively [27]. Leaf biomass and leaf blade length display an almost unimodal annual pattern; both markedly increase from March to July–August attaining a maximum value of approx. 150 g dry wt m−2 and 542.2 mm, respectively, while rhizome biomass and root biomass peak in mid or late summer and in mid autumn to early winter attaining a maximum value of approx. 225 and 125 g dry wt m−2, respectively ([27], unpublished data).

Plants were collected at the site V at 0.7–1.0 m depth in July 2011 with a 20 cm diameter acrylic corer, which penetrated to a depth of 30 cm; all the above- and below-substrate plant material rooted within this area was collected. All plants were rinsed in seawater at the collection site and transported to the laboratory in plastic containers containing seawater.

Treatments

Fresh green plants without epiphytes were kept for 24 h in seawater under laboratory conditions in order to equilibrate. Plants were incubated in plastic aquaria containing 10 L of cadmium sulphate hydrate (3CdSO4·8H2O 98.7%, insoluble matter ≤ 0.005%, chloride ≤ 0.001%, total nitrogen ≤ 0.0005%, Ca ≤ 0.005%, Cu ≤ 0.0005%, Fe ≤ 0.0005%, K ≤ 0.01%, Na ≤ 0.005%, Pb ≤ 0.002%, Zn ≤ 0.002%; lot number: 1.02027.0100; Merck, Darmstadt, Germany) dissolved in filtered (Whatman GF/C) seawater at one of the following cadmium concentrations: 0.5, 5, 10, 20 and 40 mg L−1, corresponding to 4.44, 44.48, 88.95, 177.94 and 355.88 μΜ, respectively. Plant material of about 12.5 g dry wt, including about 70 leaf shoots and the corresponding rhizomes and roots, was incubated in each aquarium; no sediment and no complexing agents were added. The seawater used for the experiments was also collected from the sampling site. The seawater used in the experiments had salinity 36.7 psu, pH 7.9, dissolved O2 5.88 mg L−1, N-NO2 0.02 μΜ, N-NO3 0.37 μΜ, Ν-NH4+ 0.43 μΜ and cadmium 0.267 μg L−1. The media in the aquaria were changed every 2 days in order to maintain the original levels. The aquaria were aerated constantly using aquarium pumps and covered with plastic film (Sanitas, Sarantis S.A., Athens, Greece) in order to prevent evaporation. The experiments were conducted under a constant 16:8 h day:night regime at ambient temperature of 21 ± 1 °C and irradiance of 120 μmοl m−2 s−1. After 0, 3, 5, 7 and 9 days, at least three samples, each one including 3 or 4 leaf shoots (about 10 leaves, approx. 270 mg dry wt) and the corresponding rhizomes (approx. 210 mg dry wt) and roots (approx. 140 mg dry wt), were removed at random from each aquarium. Plant material was separated into leaves, rhizomes and roots. Seagrass leaves were characterized as adult or intermediate-juvenile; adult leaves, which accounted for about 85% of the total leaf material, were used for cadmium analysis. Adult leaf material was further partitioned into leaf blades and leaf sheaths. Leaf age estimation was based on the morphological features of the sheath; leaves with a well-developed sheath that encompassed other leaves in their interior were classed as adults [28]. Similar procedures have been used in previous studies [1416].

Cadmium determination

The material from each plant compartment (adult leaf blades, leaf sheaths, rhizomes and roots) of the samples from a single aquarium collected on the same day was pooled. The pooled samples were washed in double-distilled water, dried to a constant weight (60 °C) and ground in an agate mill. Three subsamples of each powdered sample were wet digested in HNO3/HClO4 (4:1) at 50 °C for 1 h and then at 130 °C for 3 h. Similar methods have been frequently used in previous studies [1416, 18]. Cadmium concentrations were measured using graphite furnace atomic absorption spectrophotometry (AAS, AANALYST 700 Perkin-Elmer). Pro-analysis grade reagents were used and reagent blanks were run concurrently. Standards were prepared by serial dilution of stock solutions. The accuracy of the technique was checked by analysis of standard sea lettuce reference material (Ulva lactuca no 279, Community Bureau of Reference BCR, Brussels, Belgium); one sample of the standard reference material was included in each analytical batch. Results were in agreement with certified values (certified value, mean ± SD: 0.274 ± 0.032 μg g−1 dry wt; measured value, mean ± SD: 0.280 ± 0.020 μg g−1 dry wt; recovery: 102%).

Data analysis

Cadmium accumulation kinetics were fitted to a version of the Michaelis–Menten equation; this Michaelis–Menten-type equation (Eq. 1) is frequently used in metal kinetic studies, where C represents the tissue metal concentration reached in time t, Cmax the maximum or saturation concentration, and K m the time taken to reach half of the value of Cmax [2931].
$${\text{C}} = {{\left( {{\text{C}}_{\text{max} } \times t} \right)} \mathord{\left/ {\vphantom {{\left( {{\text{C}}_{\hbox{max} } \times t} \right)} {\left( {K_{m} + t} \right)}}} \right. \kern-0pt} {\left( {K_{m} + t} \right)}}$$
(1)

The data were analyzed using IBM Statistics SPSS® 19 (New York, NY, USA), by means of nonlinear regression, and with sequential quadratic programming as the estimation method [31]. The rate of the initial uptake (found by dividing half of the value of Cmax by K m ) was also estimated.

When the fit to Eq. 1 was not significant at the 0.01 level, the data were fitted to different regression models (linear, Eq. 2; logarithmic, Eq. 3; inverse, Eq. 4; exponential, Eq. 5; power, Eq. 6; S, Eq. 7), also using IBM Statistics SPSS® 19; in each case, the model that provided the best fit (i.e., at the highest significance level) was chosen [31].
$${\text{C}} = \alpha + b \times t$$
(2)
$${\text{C}} = \alpha + b \times \ln \left( {t + 1} \right)$$
(3)
$${\text{C}} = \alpha + \, \left[ {b{{} \mathord{\left/ {\vphantom {{} {\left( {t + 1} \right)}}} \right. \kern-0pt} {\left( {t + 1} \right)}}} \right]$$
(4)
$${\text{C}} = \alpha \times e^{b\; \times \;t}$$
(5)
$$\text{lnC} = \ln \alpha + b \times \ln \left( {t + 1} \right)$$
(6)
$$\text{lnC} = \alpha + \left[ {{b \mathord{\left/ {\vphantom {b {\left( {t + 1} \right)}}} \right. \kern-0pt} {\left( {t + 1} \right)}}} \right]$$
(7)
In order for the information on accumulation kinetics to be completed, the time required to reach equilibrium (Teq), the equilibrium concentration (Ceq) and the mean rate of uptake (Vc) were calculated. Ceq was estimated from the equation used in each case as the tissue concentration at which the daily increase in concentration was less than 1% of that of the previous day, Teq was estimated as the time required to reach Ceq, and Vc by dividing Ceq by Teq [29, 31]. Bioconcentration factor (BCF) at equilibrium was also calculated (Eq. 8), where Ceq is the equilibrium concentration, Ci the initial tissue metal concentration (at day 0) and Cw the metal concentration in the water [31, 32].
$${\text{BCF}}\, = {{\left( {{\text{C}}_{\text{eq}} \;{-}\;{\text{C}}_{\text{i}} } \right)} \mathord{\left/ {\vphantom {{\left( {{\text{C}}_{\text{eq}} \;{-}\;{\text{C}}_{\text{i}} } \right)} {{\text{C}}_{\text{w}} }}} \right. \kern-0pt} {{\text{C}}_{\text{w}} }}\,$$
(8)

The Wilcoxon matched-pairs signed-rank test was used to compare experimental metal concentrations and uptake parameters in different plant parts. Spearman’s rank correlation coefficient (ρ) was calculated to identify correlations.

Results

The initial concentrations of cadmium (0 day, mean ± standard error) in adult leaf blades, leaf sheaths, rhizomes and roots of C. nodosa were 1.773 ± 0.188, 1.066 ± 0.258, 0.655 ± 0.036 and 0.258 ± 0.008 μg g−1 dry wt, respectively.

The accumulation kinetics of cadmium in C. nodosa parts were rapid in the first 3–5 days of exposure; this initial rapid accumulation was generally followed by a slower accumulation phase and/or a steady state (Figs. 1 and 2). A variation on the last incubation day was observed at the higher exposure concentrations, particularly a marked increase in leaf sheaths at 20 mg L−1 and rhizomes at 20 and 40 mg L−1, and, on the contrary, a marked decrease in adult leaf blades at 20 and 40 mg L−1 and roots at 10 and 20 mg L−1. In addition, the initial rapid accumulation in roots at 40 mg L−1 was followed by a marked decrease after the third day of exposure.
Fig. 1
Fig. 1

Kinetics of cadmium accumulation in adult leaf blades and leaf sheaths of Cymodocea nodosa at different concentrations of cadmium in water. Values plotted are mean tissue concentration ± standard error (n = 3); bold lines are the accumulation kinetics calculated using a Michaelis–Menten-type equation

Fig. 2
Fig. 2

Kinetics of cadmium accumulation in rhizomes and roots of Cymodocea nodosa at different concentrations of cadmium in water. Values plotted are mean tissue concentration ± standard error (n = 3); bold lines are the accumulation kinetics calculated using a Michaelis–Menten-type equation (rhizomes at 0.5, 10, 20 and 40 mg L−1 and roots at 0.5 and 5 mg L−1) or an S equation (rhizomes at 5 mg L−1 and roots at 10 and 20 mg L−1)

For each incubation day, tissue cadmium concentrations generally tended to increase with the exposure concentration (Cw; Figs. 1 and 2); in particular, a significant and positive correlation was found between tissue metal content and Cw for all plant parts and all incubation times (ρ = 0.9, p < 0.05 or ρ = 1.0, p < 0.001; Table 1). Cadmium concentrations reached during the incubation period generally differed among plant parts (Figs. 1 and 2); in particular, adult leaf blades and roots displayed significantly higher cadmium concentrations than either leaf sheaths or rhizomes, when both the lower exposure concentrations (0.5–10 mg L−1) and the full set of exposure concentrations (0.5–40 mg L−1) were examined (Wilcoxon test, p < 0.01 and 0.001, respectively).
Table 1

Spearman’s rank correlation coefficient values between exposure concentration and experimental tissue cadmium concentration (C) at each incubation day

Variables

Blades

Sheaths

Rhizomes

Roots

C at day 3

1.0***

1.0***

0.9*

0.9*

C at day 5

1.0***

1.0***

1.0***

1.0***

C at day 7

1.0***

1.0***

1.0***

0.9*

C at day 9

0.9*

1.0***

1.0***

0.9*

p < 0.05; *** p < 0.001

The fit of the kinetics of cadmium accumulation in adult leaf blades, leaf sheaths and rhizomes at the 0.5–10 mg L−1 treatments and in roots of C. nodosa at the 0.5 and 5 mg L−1 treatments to the Michaelis–Menten-type equation was generally significant (r2: 0.812–0.997, p < 0.01 or 0.001; Figs. 1 and 2, Table 2). Metal uptake kinetics in adult leaf blades generally displayed a better fit (i.e., at a higher significance level) to the Michaelis–Menten-type equation, compared to those in leaf sheaths and roots. Concerning rhizomes at 5 mg L−1, the fit to the Michaelis–Menten-type equation was not significant at the 0.01 level; in this case, an S equation was found to best describe the uptake kinetics (r2: 0.993, p < 0.01; Fig. 2, Tables 3 and 4).
Table 2

Kinetics of cadmium accumulation in parts of Cymodocea nodosa exposed to different concentrations of cadmium in water

Exposure concentration (mg L−1)

 

0.5

5

10

20

40

Blades

 Cmax

306.5 (± 194.8)

611.2 (± 39.0)

3289.5 (± 1438.7)

1748.5 (± 559.9)a

7386.1 (± 6749.5)a

 K m

17.1 (± 15.3)

3.8 (± 0.6)

17.9 (± 10.9)

3.3 (± 2.6)

16.7 (± 20.4)

 Cmax/(2 × K m )

9.0

80.4

91.9

264.9

221.1

 r2

0.968***

0.997***

0.986***

0.971**

0.975**

Sheaths

 Cmax

13.5 (± 3.0)

134.8 (± 22.8)

272.8 (± 132.6)

568.4 (± 73.3)a

942.6 (± 180.6)

 K m

0.1 (± 1.1)

4.7 (± 1.8)

3.1 (± 4.3)

4.6 (± 1.3)

1.4 (± 1.3)

 Cmax/(2 × K m )

67.5

14.3

44.0

61.8

336.6

 r2

0.881**

0.985***

0.837**

0.997***

0.950**

Rhizomes

 Cmax

63.7 (± 33.9)

 

405.2 (± 204.8)

514.2 (± 158.2)a

852.0 (± 82.0)a

 K m

14.1 (± 11.2)

 

12.7 (± 9.8)

2.4 (± 2.2)

0.1 (± 0.4)

 Cmax/(2 × K m )

2.2

 

16.0

107.1

4260.0

 r2

0.968***

 

0.968***

0.965**

0.992**

Roots

 Cmax

138.8 (± 27.8)

333.3 (± 106.0)

   

 K m

0.9 (± 1.2)

0.1 (± 1.5)

   

 Cmax/(2 × K m )

77.1

1666.5

   

 r2

0.937**

0.812**

   

The exposure concentrations of 0.5, 5, 10, 20 and 40 mg L−1 correspond to 4.44, 44.48, 88.95, 177.94 and 355.88 μM, respectively

The fits correspond to a Michaelis–Menten-type equation: C = (Cmax × t)/(Km+ t)

C, tissue concentration (μg g−1 dry wt) reached in a given time; Cmax, maximum tissue concentration; Km, time (in days) to reach half of the value of Cmax; t, time (in days); standard errors are given in parentheses

** p < 0.01; *** p < 0.001

aOnly the initial four points were fitted

Table 3

Regression models tested in those cases in which the Michaelis–Menten-type model did not provide a satisfactory fit to the uptake kinetics data

 

Exposure concentration (mg L−1)

Model

r2

F

df1

df2

Significance

Rhizomes

5

Linear

0.778

7.003

1

2

0.118

Logarithmic

0.868

13.130

1

2

0.068

Inverse

0.815

8.811

1

2

0.097

Exponential

0.702

4.717

1

2

0.162

Power

0.917

22.175

1

2

0.042

S

0.993

292.077

1

2

0.003

Roots

10

Linear

0.945

34.576

1

2

0.028

Logarithmic

0.802

8.106

1

2

0.104

Inverse

0.639

3.533

1

2

0.201

Exponential

0.825

9.429

1

2

0.092

Power

0.973

73.230

1

2

0.013

S

0.998

1200.35

1

2

0.001

20

Linear

0.903

18.599

1

2

0.050

Logarithmic

0.749

5.972

1

2

0.134

Inverse

0.602

3.021

1

2

0.224

Exponential

0.792

7.606

1

2

0.110

Power

0.954

41.675

1

2

0.023

S

0.997

581.401

1

2

0.002

The exposure concentrations of 5, 10 and 20 mg L−1 correspond to 44.48, 88.95 and 177.94 μM, respectively

Table 4

Kinetics of cadmium accumulation in parts of Cymodocea nodosa exposed to different concentrations of cadmium in water

 

Exposure concentration (mg L−1)

5

10

20

Rhizomes

 α

5.512 (± 0.180)

  

 b

− 5.909 (± 0.346)

  

 r2

0.993**

  

Roots

 α

 

8.366 (± 0.148)a

8.980 (± 0.225)a

 b

 

− 9.745 (± 0.281)

− 10.313 (± 0.428)

 r2

 

0.998**

0.997**

The exposure concentrations of 5, 10 and 20 mg L−1 correspond to 44.48, 88.95 and 177.94 μM, respectively

The fits correspond to an S equation: lnC = α + [b/(t + 1)]

C, tissue concentration (μg g−1 dry wt) reached in a given time; α and b constants; t, time (in days); standard errors are given in parentheses

** p < 0.01

aOnly the initial four points were fitted

The fit of cadmium uptake kinetics in adult leaf blades, leaf sheaths and rhizomes at the higher exposure concentrations (20 and 40 mg L−1) to the Michaelis–Menten-type equation was also significant (r2: 0.950–0.997, p < 0.01 or 0.001; Figs. 1 and 2, Table 2); as for roots at the 10 and 20 mg L−1 treatments, an S equation best described the uptake kinetics (r2: 0.997–0.998, p < 0.01; Fig. 2, Tables 3 and 4). However, in most of these cases, the initial four points only were used because of the marked variation in tissue metal content on the last day of incubation. The kinetics of cadmium uptake in roots at 40 mg L−1 were not fitted to any model because of the marked decrease in tissue metal content after the third day of incubation (Fig. 2).

The values of the uptake parameters obtained for C. nodosa parts are shown in Tables 2 and 5. The values of the maximum concentration (Cmax) and the equilibrium concentration (Ceq) generally tended to increase with Cw; in particular, a significant and positive correlation was found between Cmax and Cw for adult leaf blades, leaf sheaths and rhizomes, and between Ceq and Cw for adult leaf blades, leaf sheaths and roots (ρ = 0.9, p < 0.05 or ρ = 1.0, p < 0.001; Table 5). The values of the rate of initial uptake (Cmax/(2 × K m )) and the mean rate of the uptake (Vc) also generally tended to increase with Cw (Tables 2 and 5); a significant and positive correlation was particularly found between Cmax/(2 × K m ) and Cw for adult leaf blades and rhizomes (ρ = 0.9, p < 0.05 and ρ = 1.0, p < 0.001, respectively), and between Vc and Cw for all plant parts (ρ = 1.0, p < 0.001; Table 6). Thereby, higher Cmax and Ceq values were reached with higher uptake rates.
Table 5

Equilibrium concentration, (Ceq in μg g−1 dry wt), time to reach equilibrium (Teq, in days), mean rate of uptake (Vc, in concentration/days) and bioconcentration factor at equilibrium (BCF)

Exposure concentration (mg L−1)

 

0.5

5

10

20

40

Blades

 Ceq

203.9

504.7

2155.0

1467.3a

4908.6a

 Teq

34

18

34

17

33

 Vc

6.0

28.0

63.4

86.3

148.7

 BCF

404.3

100.6

215.3

73.3

122.7

Sheaths

 Ceq

11.7

109.1

228.5

457.6a

836.2

 Teq

14

20

16

19

11

 Vc

0.8

5.5

14.3

24.1

76.0

 BCF

21.3

21.6

22.7

22.8

20.9

Rhizomes

 Ceq

43.8

193.6

144.0

439.0a

824.5a

 Teq

31

22

7

14

3

 Vc

1.4

8.8

20.6

31.4

274.8

 BCF

86.3

38.6

14.3

21.9

20.6

Roots

 Ceq

126.2

322.5

3138.9a

5694.9a

 

 Teq

9

3

29

29

 

 Vc

14.0

107.5

108.2

196.4

 

 BCF

251.9

64.4

313.9

284.7

 

The exposure concentrations of 0.5, 5, 10, 20 and 40 mg L−1 correspond to 4.44, 44.48, 88.95, 177.94 and 355.88 μM, respectively

aOnly the initial four points were fitted

Table 6

Spearman’s rank correlation coefficient values between exposure concentration uptake parameters; n = 5

Variables

Blades

Sheaths

Rhizomes

Roots

Cmax

0.9*

1.0***

1.0***

 

Km

− 0.3ns

0.1ns

− 1.0***

 

Cmax/(2 × Km)

0.9*

0.4ns

1.0***

 

Ceq

0.9*

1.0***

0.8ns

1.0***

Teq

− 0.41ns

− 0.3ns

− 0.9*

0.737ns

Vc

1.0***

1.0***

1.0***

1.0***

BCF

− 0.5ns

0.0ns

− 0.7ns

0.6ns

For abbreviations, see Tables 2 and 5

nsNon significant; * p < 0.05; *** p < 0.001

The values of the time taken to reach half of the Cmax (K m ) and the time taken to reach equilibrium (Teq) obtained for adult leaf blades displayed no clear pattern with increasing Cw; in leaf sheaths, these parameters displayed their lowest values at the lowest and the highest exposure concentration respectively, while in roots, Teq displayed its highest value at the higher exposure concentrations. As for rhizomes, K m and Teq generally tended to decrease with increasing Cw (ρ = − 1.0, p < 0.001 and ρ = − 0.9, p < 0.05, respectively; Tables 2, 5 and 6).

Comparable BCF values were obtained for leaf sheaths at all of the treatments (range 20.9–22.8); as for roots, BCF values (range 64.4–313.9) showed no clear trend with increasing Cw, while as for adult leaf blades and rhizomes, BCF values (range 73.3–404.3 and 14.3–86.3, respectively) were generally lower at higher exposure concentrations (Tables 5 and 6).

Ceq and BCF values obtained for adult blades were significantly higher than those obtained for leaf sheaths and rhizomes; Vc values estimated for adult leaf blades and rhizomes were also significantly higher than those estimated for leaf sheaths (Wilcoxon test, p < 0.05). In addition, at each exposure concentration, Ceq, Vc and BCF generally had the highest values in adult leaf blades and roots, and the lowest values in leaf sheaths and rhizomes (Table 5).

Discussion

The initial concentrations of cadmium in adult leaf blades, leaf sheaths, rhizomes and roots of Cymodocea nodosa collected for the experiments were within the range of cadmium concentrations earlier measured in C. nodosa parts from various localities along Mediterranean coasts (see review in [28]). The maximum experimental concentration observed in adult leaf blades of C. nodosa (101.3 μg g−1 dry wt) at the lowest exposure concentration studied (0.5 ml L−1) was within the wide range of reported cadmium concentrations in leaves of seagrass species (0.1–266 μg g−1 dry wt) from various geographical locations (see review in [3]).

Leaf blades and roots are the main sites of ionic uptake in seagrasses [33, 34] and metal uptake is considered to follow two pathways-from surrounding water to leaves and then to rhizomes, or from interstitial water into roots to rhizomes and leaves [35]. Translocation of cadmium within a seagrass species (Zostera marina) has been previously demonstrated in laboratory experiments; in particular, both a bidirectional translocation and a translocation from leaves to root-rhizomes have been reported [36, 37].

The uptake kinetics of cadmium into parts of C. nodosa at the lower exposure concentrations examined (0.5–5 or 10 mg L−1), namely at concentrations within the range of cadmium concentrations in industrial wastewater [21], generally displayed a similar pattern: an initial rapid accumulation was followed by a slower accumulation phase and an equilibrium state. This pattern is consistent with previous observations concerning the accumulation of cadmium into parts of marine angiosperms [7, 11, 14, 18, 36]. A Michaelis–Menten-type equation satisfactorily described cadmium accumulation kinetics in most cases, while an S equation best described the uptake kinetics in rhizomes at 5 mg L−1 and roots at 10 mg L−1, both permitting to calculate uptake parameters.

The observed accumulation kinetics was most probably the net result of several different processes. A potential release by the seagrass into the medium of organic ligands, capable to complex dissolved metals, may have played a role in controlling the cadmium uptake [3840]. Cadmium accumulation may involve a combination of adsorption onto the outer cell wall and uptake into the cells; the steady state may correspond to an equilibrium attained between the metal in the medium and the metal bound on the cell surface, while intracellular uptake may include both a diffusion across the plasma membrane into the protoplasm and an active accumulation of metal within the plant cells [29, 32, 41, 42]. Hence, cadmium is expected to be accumulated in different cellular compartments. In addition, cadmium accumulation, particularly in leaf sheaths and rhizomes, may have resulted from both a direct uptake from the surrounding medium and some internal transport.

Tissue cadmium concentrations reached at each incubation time at the lower exposure concentrations (0.5–5 or 10 mg L−1) and, thereby, uptake parameters too, appeared to be a function of cadmium concentration in water. The maximum concentration and the equilibrium concentration, as well as the rate of initial uptake and the mean rate of uptake respectively, generally tended to increase with the exposure concentration, indicating that C. nodosa parts display a remarkable absorption capacity of cadmium and an abundance of cell wall or intracellular binding sites [43]. This finding also indicates that cadmium in C. nodosa is correlated with that in the surrounding medium over a wide range of exposure concentrations [30]. However, as for adult leaf blades and rhizomes, the bioconcentration factor at equilibrium generally displayed lower values at higher exposure concentrations, suggesting a reduction of available binding sites [7] and, thereby, a lower sensitivity in detecting high levels of cadmium contamination [31].

Cadmium concentrations reached during the incubation period at the lower exposure concentrations (0.5–5 or 10 mg L−1) and, thereby, uptake kinetic parameters too, appeared to differ to some extent among C. nodosa parts, indicating within-plant differences in cadmium accumulation capacity. Adult leaf blades and roots, which present the highest surface/volume ratio and are the main sites of ionic uptake [3335, 43], generally accumulated more cadmium with higher uptake rates than leaf sheaths and rhizomes, in which, as noted earlier, metal accumulation may have resulted from both a direct uptake from the surrounding medium and some internal transport. In addition, cadmium uptake kinetics in adult leaf blades generally displayed a better fit to the Michaelis–Menten-type equation, compared to those in the remaining plant parts. Variability in the distribution pattern of cadmium among C. nodosa parts might be expected in field situations due to differences in the bioavailability of this element in the water column and sediment. Nevertheless, our findings are consistent with most of the previous field observations, whereby C. nodosa leaves and roots accumulated the highest cadmium loads and rhizomes the lowest ones (see review in [28]); in particular, at the same sampling area (Viamyl site), C. nodosa displayed over the year significantly lower concentrations of cadmium in rhizome parts than in blades, sheaths and roots [28]. A comparison of the experimental cadmium concentrations reached in adult leaf blades of C. nodosa during the incubation period with the corresponding concentrations in intermediate-juvenile leaf blades of this seagrass species incubated to the same exposure concentrations [18] reveals that adult leaf blades accumulate more cadmium than intermediate and juvenile ones; this finding could be explained by a synthesis of more binding sites over time [44, 45].

The above findings suggest that C. nodosa could be considered as a potent tool for the assessment of cadmium levels of 0.5–5 or 10 mg L−1 in the surrounding medium in coastal areas after an accidental discharge of untreated metal-bearing industrial effluents or in inshore wastewater treatment areas. Considering both our data and operational criteria (e.g. ease of sampling), adult leaf blades appear to be the most suited plant part for the determination of cadmium levels in the surrounding environment. The above findings also suggest that C. nodosa could be considered as a valuable tool for the treatment of cadmium from industrial effluents or in the abatement of cadmium contamination in coastal environments receiving untreated industrial effluents, when exposure concentrations range from 0.5 to 5 or 10 mg L−1. For instance, contaminant amounts could be removed by harvesting mainly seagrass adult leaves when the equilibrium state in metal accumulation is achieved [46].

A comparison of the maximum experimental cadmium concentrations observed in parts of C. nodosa with those previously reported for the respective parts of other seagrass species incubated to similar concentrations of cadmium in seawater corroborates the finding that C. nodosa displays a remarkable accumulation capacity of cadmium. For instance, the maximum experimental cadmium concentrations in rhizomes and roots of C. nodosa (142.1 and 405.6 μg g−1 dry wt, respectively) at the 5 mg L−1 treatment were higher than those in rhizomes and roots of Zostera marina (approx. 56.0 and 320.5 μg g−1 dry wt, respectively) exposed for 19 days to seawater containing 5.62 mg L−1 cadmium [11]; the maximum experimental cadmium concentration in adult leaf blades of C. nodosa (1063.6 μg g−1 dry wt) at the 10 mg L−1 treatment was also higher than that in leaves of Halophila stipulacea (721 μg g−1 dry wt) incubated for 16 days in seawater containing cadmium in concentration of 11.24 mg L−1 [14]. In addition, the maximum experimental cadmium concentration in adult leaf blades of C. nodosa (1063.6 μg g−1 dry wt) at the 10 mg L−1 treatment was higher compared to the respective one in leaves of the submerged brackish-water angiosperm Ruppia maritima (772.0 μg g−1 dry wt) exposed to lagoon water (8.6 psu salinity) also containing cadmium in concentration of 10 mg L−1 [47], despite a decreased cadmium accumulation with increasing salinity is expected due to the decreased availability of cadmium in the medium because of the complexes formed between chloride and metal [48].

The marked variation in tissue cadmium concentrations mainly observed at the last day of incubation at the higher exposure concentrations studied (20–40 mg L−1) could be related to the stress induced on seagrass cells. A potential explanation for the marked decrease in adult leaf blade and root cadmium concentrations is a release of accumulated metal quantities to the exterior [29]. Seagrass cells may possess active efflux systems for non-essential elements, as observed for cadmium in marine diatoms [42], to minimize the toxic impact of metal exposure [49]. A potential release (leakage) of cadmium by tissues of a seagrass species (Zostera marina) has been previously reported by Brinkhuis et al. [36]. This marked decrease could be also ascribed to an extended internal transport of accumulated metal quantities from roots and leaf blades towards to rhizomes and leaf sheaths; this interpretation is corroborated by the observation that leaf sheath and rhizome cadmium concentrations markedly increased at the last day of exposure at the 20 mg L−1 treatment (as for sheaths) or at both the 20 and 40 mg L−1 treatments (as for rhizomes). This marked increase in leaf sheath and rhizome cadmium concentrations could also be ascribed to an increase in available binding sites due to potential cell deterioration [11, 12, 14].

Conclusions

Cymodocea nodosa displays a remarkable accumulation capacity of cadmium, varying among plant parts and reflects high cadmium levels in the surrounding medium. Kinetic models, usually a Michaelis–Menten-type equation, satisfactorily describe cadmium uptake patterns in plant parts, primarily in adult leaf blades, at exposure concentrations within the range of cadmium levels in industrial wastewater, permitting to calculate uptake parameters and to predict cadmium accumulation in field situations. The data presented contribute to a better understanding of metal accumulation in seagrasses and highlight the usefulness of C. nodosa as a biomonitor of cadmium contamination and as a valuable tool for phytoremediation, when exposure concentrations range from 0.5 to 5 or 10 mg L−1.

Declarations

Authors’ contributions

PM contributed extensively to all aspects of the work presented in this manuscript. TK participated in the conception and design of the project, and in the sample collection, data analysis and writing the manuscript. KRC was involved in sample pretreatment and AM in cadmium determination procedures. All authors read and approved the final manuscript.

Acknowledgements

Thanks are due to the anonymous referees for thoughtful critiques and comments.

Competing interests

The authors declare that they have no competing interests.

Availability of data and materials

The authors declare that data analyzed in the current study are available within the article.

Consent for publication

Not applicable.

Ethics approval and consent to participate

Not applicable.

Funding

Not applicable.

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Authors’ Affiliations

(1)
Department of Botany, School of Biology, Aristotle University of Thessaloniki, 54124 Thessaloniki, Greece
(2)
Laboratory of Environmental Research and Education, Democritus University of Thrace, Nea Hili, 68100 Alexandroupolis, Greece

References

  1. Virnstein RW, Morris LJ. Setting seagrass targets for the Indian River Lagoon, Florida. In: Bortone SA, editor. Seagrasses: monitoring, ecology, physiology and management. Boca Raton: CRC Press; 2000. p. 211–29.Google Scholar
  2. Huesemann MH, Hausmann TS, Fortman TJ, Thom RM, Cullinan V. In situ phytoremediation of PAH- and PCB-contaminated marine sediments with eelgrass (Zostera marina). Ecol Eng. 2009;35:1395–404.View ArticleGoogle Scholar
  3. Lewis MA, Devereux R. Nonnutrient anthropogenic chemicals in seagrass ecosystems: fate and effects. Environ Toxicol Chem. 2009;28:644–61.View ArticlePubMedGoogle Scholar
  4. Pergent G, Labbe C, Lafabrie C, Kantin R, Pergent-Martini C. Organic and inorganic human-induced contamination of Posidonia oceanica meadows. Ecol Eng. 2011;37:999–1002.View ArticleGoogle Scholar
  5. Kennish MJ. Estuary restoration and maintenance. Boca Raton: CRC Press; 2000.Google Scholar
  6. Ferrat L, Pergent-Martini C, Roméo M. Assessment of the use of biomarkers in aquatic plants for the evaluation quality: application to seagrasses. Aquat Toxicol. 2003;65:187–204.View ArticlePubMedGoogle Scholar
  7. Fabris GJ, Harris JE, Smith JD. Uptake of cadmium by the seagrass Heterozostera tasmanica from Corio Bay and Western Port, Victoria. Aust J Mar Freshw Res. 1982;33:829–36.View ArticleGoogle Scholar
  8. Cristiani G, Gassend R, Augier H. Etude de la contamination experimentale de la phanerogame marine Posidonia oceanica (L.) Delile par les composes mercuriques: Partie 1—Modalites de la contamination par le chlorure mercurique. Environ Pollut (Series A). 1980;23:153–62.View ArticleGoogle Scholar
  9. Warnau M, Fowler SW, Teyssié JL. Biokinetics of selected heavy metals and radionuclides in two marine macrophytes: the seagrass Posidonia oceanica and the alga Caulerpa taxifolia. Mar Environ Res. 1996;41:343–62.View ArticleGoogle Scholar
  10. Ferrat L, Bingert A, Roméo M, Gnassia-Barelli M, Pergent-Martini C. Mercury uptake and enzymatic response of Posidonia oceanica after an experimental exposure to organic and inorganic forms. Environ Toxicol Chem. 2002;21:2365–71.View ArticlePubMedGoogle Scholar
  11. Lyngby JE, Brix H. The uptake of heavy metals in eelgrass Zostera marina and their effect on growth. Ecol Bull. 1984;36:81–9.Google Scholar
  12. Bond AM, Bradbury JR, Hudson HA, Garnham JS, Hanna PJ, Strother S. Kinetic studies of lead (II) uptake by the seagrass Zostera muelleri in water by radiotracing, atomic absorption spectrometry and electrochemical techniques. Mar Chem. 1985;16:1–9.View ArticleGoogle Scholar
  13. Bond AM, Reust V, Hudson HA, Arnup KR, Hanna PJ, Strother S. The effects of temperature, salinity and seagrass species on the uptake of lead (II) from seawater by excised leaves. Mar Chem. 1988;24:253–60.View ArticleGoogle Scholar
  14. Malea P. Uptake of cadmium and the effect on viability of leaf cells in the seagrass Halophila stipulacea (Försk.) Aschers. Bot Mar. 1994;37:67–73.Google Scholar
  15. Malea P, Haritonidis S, Kevrekidis T. The short-term uptake of copper by the two parts of the seagrass Halophila stipulacea (Försk.) Aschers and leaf-cells viability. Fresen Environ Bull. 1995;4:117–22.Google Scholar
  16. Malea P, Kevrekidis T, Haritonidis S. The short-term uptake of zinc and cell mortality of the seagrass Halophila stipulacea (Försk.) Aschers. Isr J Plant Sci. 1995;43:21–30.View ArticleGoogle Scholar
  17. Malea P, Haritonidis S. Toxicity and uptake of aluminium by the seagrass Halophila stipulacea (Försk.) Aschers., in response to aluminium exposure. Fresen Environ Bull. 1996;5:345–50.Google Scholar
  18. Malea P, Adamakis ID, Kevrekidis T. Kinetics of cadmium accumulation and its effects on microtubule integrity and cell viability in the seagrass Cymodocea nodosa. Aquat Toxicol. 2013;144–5:257–64.View ArticleGoogle Scholar
  19. Malea P, Adamakis ID, Kevrekidis T. Effects of lead uptake on microtubule cytoskeleton organization and cell viability in the seagrass Cymodocea nodosa. Ecotoxicol Environ Saf. 2014;104:175–81.View ArticlePubMedGoogle Scholar
  20. Nosier SA. Removal of cadmium ions from industrial wastewater by cementation. Chem Biochem Eng Q. 2003;17:219–24.Google Scholar
  21. Vullo DL, Ceretti HM, Daniel MA, Ramírez SA, Zalts A. Cadmium, zinc and copper biosorption mediated by Pseudomonas veronii 2E. Bioresour Technol. 2008;99:5574–81.View ArticlePubMedGoogle Scholar
  22. Terrados J, Ros JD. Growth and primary production of Cymodocea nodosa (Ucria) Ascherson in a Mediterranean coastal lagoon: the Mar Menor (SE Spain). Aquat Bot. 1992;43:63–74.View ArticleGoogle Scholar
  23. Reyes J, Sansón M, Afonso-Carrillo J. Distribution and reproductive phenology of the seagrass Cymodocea nodosa (Ucria) Ascherson in the Canary Islands. Aquat Bot. 1995;50:171–80.View ArticleGoogle Scholar
  24. Cancemi G, Buia MC, Mazzella L. Structure and growth dynamics of Cymodocea nodosa meadows. Sci Mar. 2002;66:365–73.View ArticleGoogle Scholar
  25. Malea P, Kevrekidis T. Trace element (Al, As, B, Ba, Cr, Mo, Ni, Se, Sr, Tl, U and V) distribution and seasonality in compartments of the seagrass Cymodocea nodosa. Sci Total Environ. 2013;463–4:611–23.View ArticleGoogle Scholar
  26. Malea P, Adamakis ID, Kevrekidis T. Microtubule integrity and cell viability under metal (Cu, Ni and Cr) stress in the seagrass Cymodocea nodosa. Chemosphere. 2013;93:1035–42.View ArticlePubMedGoogle Scholar
  27. Malea P, Zikidou C. Temporal variation in biomass partitioning of the seagrass Cymodocea nodosa at the Gulf of Thessaloniki, Greece. J Biol Res Thessalon. 2011;15:75–90.Google Scholar
  28. Malea P, Kevrekidis T, Potouroglou M. Seasonal variation of trace metal (Mn, Zn, Cu, Pb Co, Cd) concentrations in compartments of the seagrass Cymodocea nodosa. Bot Mar. 2013;56:169–84.View ArticleGoogle Scholar
  29. Fernández JA, Vázquez MD, López J, Carballeira A. Modelling the extra and intracellular uptake and discharge of heavy metals in Fontinalis antipyretica transplanted along a heavy metal and pH contamination gradient. Environ Pollut. 2006;139:21–31.View ArticlePubMedGoogle Scholar
  30. Costa S, Crespo D, Henriques BMG, Pereira E, Duarte AC, Pardal MA. Kinetics of mercury accumulation and its effects on Ulva lactuca growth rate at two salinities and exposure conditions. Water Air Soil Pollut. 2011;217:689–99.View ArticleGoogle Scholar
  31. Díaz S, Villares R, Carballeira A. Uptake kinetics of As, Hg, Sb, and Se in the aquatic moss Fontinalis antipyretica Hedw. Water Air Soil Pollut. 2012;223:3409–23.View ArticleGoogle Scholar
  32. Martins RJE, Boaventura RAR. Uptake and release of zinc by aquatic bryophytes (Fontinalis antipyretica L. ex. Hedw.). Water Res. 2002;36:5005–12.View ArticlePubMedGoogle Scholar
  33. Kuo J, den Hartog C. Seagrass morphology, anatomy, and ultrastructure. In: Larkum AWD, Orth RJ, Duarte CM, editors. Seagrasses: biology, ecology and conservation. Springer: Dordrecht; 2007. p. 51–87.Google Scholar
  34. Romero J, Lee K-S, Pérez M, Mateo MA, Alcoverro T. Nutrients dynamics in seagrass ecosystems. In: Larkum AWD, Orth RJ, Duarte CM, editors. Seagrasses: biology, ecology, and conservation. Springer: Dordrecht; 2007. p. 227–54.Google Scholar
  35. Ralph PJ, Tomasko D, Moore K, Seddon S, Macinnis-Ng CMO. Human impacts on seagrasses: eutrophication, sedimentation and contamination. In: Larkum AWD, Orth RJ, Duarte CM, editors. Seagrasses: biology, ecology and conservation. Springer: Dordrecht; 2007. p. 567–93.Google Scholar
  36. Brinkhuis BH, Penello WF, Churchill AC. Cadmium and manganese flux in eelgrass Zostera marina. II. Metal uptake by leaf and root-rhizome tissues. Mar Biol. 1980;58:187–96.View ArticleGoogle Scholar
  37. Faraday WE, Churchill AC. Uptake of cadmium by the eelgrass Zostera marina. Mar Biol. 1979;53:293–8.View ArticleGoogle Scholar
  38. Naidu R, Harter RD. Effect of different organic ligands on cadmium sorption by extractability from soil. Soil Sci Soc Am J. 1998;62:644–50.View ArticleGoogle Scholar
  39. Vasconcelos MTSD, Leal MFC. Seasonal variability in the kinetics of Cu, Pb, Cd and Hg accumulation by macroalgae. Mar Chem. 2001;74:65–85.View ArticleGoogle Scholar
  40. Karavoltsos S, Sakellari A, Strmečki S, Plavšić M, Ioannou E, Roussis V, et al. Copper complexing properties of exudates and metabolites of macroalgae from the Aegean Sea. Chemosphere. 2013;91:1590–5.View ArticlePubMedGoogle Scholar
  41. Pickering DC, Puia IL. Mechanism for the uptake of zinc by Fontinalis antipyretica. Physiol Plant. 1969;22:653–61.View ArticleGoogle Scholar
  42. Sunda WG, Huntstman SA. Processes regulating cellular metal accumulation and physiological effects: phytoplankton as model system. Sci Total Environ. 1998;219:165–81.View ArticleGoogle Scholar
  43. Sanchiz C, Garcías-Carrascosa AM, Pastor A. Bioaccumulation of Hg, Cd, Pb and Zn in four marine phanerogams and the alga Caulerpa prolifera (Försskal) Lamouroux from the east coast of Spain. Bot Mar. 1999;42:157–64.View ArticleGoogle Scholar
  44. Bryan GW, Hummerstone LG. Brown seaweed as an indicator of heavy metals in estuaries in South-West England. J Mar Biol Assoc UK. 1973;53:705–20.View ArticleGoogle Scholar
  45. Brix H, Lyngby JE. The distribution of some metallic elements in eelgrass (Zostera marina L.) and sediment in the Limfjord. Denmark. Estuar Coast Shelf Sci. 1983;16:455–67.View ArticleGoogle Scholar
  46. Wani RA, Ganai BA, Shah MA, Uqab B. Heavy metal uptake potential of aquatic plants through phytoremediation technique—a review. J Bioremediat Biodegrad. 2017;8:4. https://doi.org/10.4172/2155-6199.1000404.View ArticleGoogle Scholar
  47. Malea P, Kevrekidis T, Mogias A, Adamakis IDS. Kinetics of cadmium accumulation and occurrence of dead cells in leaves of the submerged angiosperm Ruppia maritima. Bot Mar. 2014;57:111–22.View ArticleGoogle Scholar
  48. Fritioff Å, Kautsky L, Greger M. Influence of temperature and salinity on heavy metal uptake by submersed plants. Environ Pollut. 2005;133:265–74.View ArticlePubMedGoogle Scholar
  49. Ralph PJ, Burchett MD. Photosynthetic response of Halophila ovalis to heavy metal stress. Environ Pollut. 1998;103:91–101.View ArticleGoogle Scholar

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