HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS		QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

This Article Abstract Figures Only Full Text (PDF) Free Alert me when this article is cited Alert me if a correction is posted Services Related articles in JEQ Similar articles in this journal Similar articles in ISI Web of Science Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via ISI Web of Science (7) Citing Articles via Google Scholar Google Scholar Articles by Chaignon, V. Articles by Hinsinger, P. Search for Related Content PubMed PubMed Citation Articles by Chaignon, V. Articles by Hinsinger, P. GeoRef GeoRef Citation Agricola Articles by Chaignon, V. Articles by Hinsinger, P. Related Collections Lysimeter/Rhizosphere Studies Ecological Risk Assessment Heavy Metals Plant and Soil Interactions Soil Pollution

TECHNICAL REPORTS
Heavy Metals in the Environment

A Biotest for Evaluating Copper Bioavailability to Plants in a Contaminated Soil

ENSA.M-INRA, UMR Rhizosphère & Symbiose, Place Viala, F-34060 Montpellier Cedex 1, France

^* Corresponding author (philippe.hinsinger{at}ensam.inra.fr)

Received for publication April 12, 2001.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS AND DISCUSSION CONCLUSIONS REFERENCES

 
This work aimed at defining the optimal conditions for a novel ecotoxicological test designed for evaluating the bioavailability and phytotoxicity of metals to plants. This biotest, which provided easy access to roots, shoots, and rhizosphere soil, was applied to a vineyard calcareous soil that had been contaminated by the application of Cu fungicides. A preliminary hydroponic experiment comparing various levels of solution Cu concentration enabled us to determine the no observable adverse effects concentration (NOAEC), which was in the range 5 to 20 µM total Cu (0.01–0.06 µM free Cu ion) for rape (Brassica napus L. cv. Goeland). For the biotest, rape was grown in hydroponic conditions for 21 d in pots designed so that plants developed a planar mat of roots at the surface of a polyamide mesh. By then, the plants were transferred for 4 or 8 d onto a 1- or 3-mm-thick soil layer that was separated from the root mat by the mesh and connected to a reservoir of nutrient solution or deionized water via a filter paper wick. An 8-d period was the best option as it enabled plant growth to be significant. The use of 1-mm soil thickness was recommended if the biotest aimed at investigating root-induced changes in the rhizosphere. Although it may cause some artifacts, compared with deionized water, nutrient solution provided better standardized conditions for comparing widely differing soil samples. The studied soil did not induce any Cu phytotoxicity in spite of its fairly large total Cu content.

Abbreviations: DM, dry matter • DTPA, diethylene triaminepentaacetic acid • EDTA, ethylenediaminetetraacetic acid • NOAEC, no observable adverse effects concentration

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS AND DISCUSSION CONCLUSIONS REFERENCES

 
THE CONTAMINATION of agricultural land can arise from various origins such as the use of fertilizers and pesticides, the application of sewage sludge, the disposal of industrial wastes or the deposition of atmospheric contaminants (Alloway, 1995). In all cases, the potential risks for terrestrial and aquatic ecosystems need to be estimated. Although this problem is of increasing concern, it is surprising that, compared with aquatic ecosystems, risk evaluation of terrestrial ecosystems still relies on only a few standardized ecotoxicological tests. Short-term plant toxicity tests have originally been developed from simple measurements used in plant physiology and weed science (Cushman and Meyer, 1990). The two short-term phytotoxicity tests that involve terrestrial plants and are currently used are (i) the seed germination test and (ii) the root elongation test (Kapustka, 1997; International Organization for Standardization, 1999). The seed germination test is relatively insensitive to many toxic substances: many of these are not readily taken up into the plants because at this stage, the seedlings derive their nutrients mostly from seed reserves rather than from the soil. This test thus provides a poor indication of soil contamination, while the root elongation assay provides a more sensitive measure of phytotoxic effects (Kapustka, 1997). However, the root elongation test is conducted in quartz sand with soil eluates (water extracts) and is therefore quite far from realistic exposure conditions. In comparison with these two standardized tests, the early seedling growth test (American Society for Testing and Materials, 1994) and related growth tests based on pot experiments (International Organization for Standardization, 1999) are closer to "real-world" conditions of exposure (Kapustka, 1997). In addition to their duration and cost, the major drawbacks of these tests are that the exposure conditions, relevant endpoints, interspecies extrapolation, and lab-to-field extrapolation are not clearly defined (American Society for Testing and Materials, 1994; Kapustka, 1997; International Organization for Standardization, 1999). Standardized plant tests (as early seedling growth tests) that are published in the literature dealing with phytotoxicity assessment authorize modifying the test procedure to make the test more relevant (American Society for Testing and Materials, 1994). However, in all cases, only aerial parts of plants are considered while root contamination and bioaccumulation or bioavailability and speciation of the metal in the soil are disregarded.

The fraction of soil metal that can be taken up by plants (i.e., the bioavailable fraction) (Singh and Rakipov, 1988) is most often estimated with chemical extractions that are routinely used in soil testing (Lebourg et al., 1996). Many chemical extractants (CaCl₂ and other dilute salts, DTPA [diethylene triaminepentaacetic acid] and EDTA [ethylenediaminetetraacetic acid]) have been used to identify the portion of metals in the soil that is the most readily available (Beckett, 1989; Lebourg et al., 1996). Such chemical extractions can hardly account for the various, complex processes that are involved in the acquisition of metals by plants (Marschner, 1995; McLaughlin et al., 1998; Hinsinger, 2001), and are thus not satisfactory for estimating soil metal bioavailability to plants. Biotests conducted with plants provide an alternative approach to estimate metal bioavailability in contaminated soils. However, the biotests that have been used by various authors for this purpose are based on pot experiments, for which conditions vary considerably. Being poorly standardized, they cannot easily be compared with one another.

The present work is focused on the case of a vineyard soil that has been contaminated by the long-term field application of copper salts (Bordeaux mixture) as fungicides against powdery mildew. For most plant species, the critical toxicity level of copper in the leaves is 20 to 30 mg kg^-1 dry matter (Reuter and Robinson, 1997). This criterion is, however, not a sufficient indication of the copper toxicity and bioavailability for the plant (Mitchell et al., 1978; Lexmond, 1980; Brun et al., 2001). Indeed, a large copper supply can inhibit root growth before shoot growth (Marschner, 1995) and Cu can accumulate in the roots without any significant increase in the Cu content of the aerial parts (Mitchell et al., 1978; Brun et al., 2001). Therefore, a proper biotest should provide easy access not only to the shoots, but also to the roots to evaluate Cu contamination of plants (Brun et al., 2001). But when plants are grown in soils, especially the plants with fine roots (Brun et al., 2001), root collection is easily biased and tedious. Indeed, at harvest, small amounts of soil can adhere to roots, resulting in overestimates of root Cu content (Brun et al., 2001), and the finest roots are often discarded from the analysis because of their difficult and almost impossible recovery.

The aim of this work was to define the optimal conditions (time duration, amount of soil, and nutrients supplied) for a novel biotest that provides an easy access to roots, shoots, and rhizosphere soil to evaluate the bioavailability and phytotoxicity of Cu to oilseed rape plantlets. A preliminary biotest was performed in nutrient solution over a range of Cu concentrations to evaluate the sensitivity of rape roots and shoots to Cu toxicity.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS AND DISCUSSION CONCLUSIONS REFERENCES

 
General Principles of the Proposed Biotest
The cropping device that was used for the present biotest was adapted from that of Guivarch et al. (1999), which evolved from those designed by Kuchenbuch and Jungk (1982) and Niebes et al. (1993). The major advantage of this technique is that roots are physically separated from the soil, which enables an easy recovery of the shoots, roots, and soil. Another advantage of this device is that the thickness of the soil layer used and, hence, the distance from the roots in the soil compartment can be as little as a few millimeters. It can thereby be considered as rhizosphere soil and provide enough rhizosphere material to evaluate root-induced changes in metal speciation. In that respect, the proposed biotest is just a simplified version of the "root mat" technique that has been extensively used in rhizosphere research over the last decades (Jaillard et al., 2003). Despite the physical separation between roots and soil, intense chemical interactions can occur (e.g., reviews by Hinsinger, 1998; Jaillard et al., 2003) and plants can thereby successfully take up nutrients (e.g., Kuchenbuch and Jungk, 1982; Niebes et al., 1993; Morel and Hinsinger, 1999) and pollutants (Guivarch et al., 1999; Kruyts et al., 2000) from the soil, at fairly large fluxes. This root mat approach has also been successfully used to study the toxicity of Al in the rhizosphere of maize (Zea mays L.) (Calba et al., 1996).

The proposed biotest is a two-step procedure: in the first step, plants are grown for three weeks in hydroponic conditions ("preculture") and in the second step, they are transferred onto the soil for another few days of growth ("soil experiment"). In a preliminary experiment, preculture plants were grown for another two weeks in hydroponic conditions with various levels of Cu ("hydroponic experiment").

Hydroponic Preculture
The plant containers were prepared as follows: a polyamide grid (900-µm pore diameter) was pasted at the bottom side of a PVC cylinder (25-mm i.d.). Another PVC cylinder (32-mm i.d.) was closed by a fine polyamide mesh (30-µm pore diameter, Fyltis/Nytel 03-30/18; Sefar Filtration, Rüschlikon, Switzerland) at the bottom side. The small cylinder was inserted into the large one. A space of 3 mm was left between the grid and the mesh so that roots could develop as a planar mat (roughly 3-mm-thick disk with a 32-mm diameter) in between.

Seeds of oilseed rape (Brassica napus L. cv. Goeland) were surface-sterilized with 6% H₂O₂ for 10 min and rinsed with deionized water. Ten seeds were sown in each container onto the surface of the grid. Plants were grown in hydroponic conditions for 21 d to obtain a large planar mat of roots that fully covered the polyamide mesh. Ten containers were placed on top of a 6-L bucket, which contained an aerated nutrient solution (Fig. 1) . A daily supply of nutrient solution was necessary to constantly wet the mesh during the 21-d "preculture" period. During the first week, plants were supplied with 600 µM CaCl₂ and 2 µM H₃BO₃. The buckets were then filled for two weeks with a complete nutrient solution of following composition: 2 mM KNO₃, 2 mM Ca(NO₃)₂, 1 mM MgSO₄, 0.5 mM KH₂PO₄, 0.1 mM FeNaEDTA, 10 µM H₃BO₃, 2 µM MnCl₂, 1 µM ZnSO₄, 0.05 µM NaMoO₄, and 0.2 µM CuCl₂. The nutrient solution initially was pH 5.5 and was renewed weekly to avoid large pH changes and nutrient depletion. The experiment was conducted in a growth chamber with the following day–night conditions: 16 h, 25°C, relative humidity 75%, and a light intensity of about 500 µmol photons m^-2 s^-1 (in the range 400–700 nm) during the day, and 8 h, 20°C, and relative humidity of 100% at night. After 21 d, the containers (taken from all the buckets) were ranked according to the size of the plants and those containing the smallest and tallest plants were discarded. Groups of five replicates were then selected randomly among the leftover containers that contained plants with homogeneous sizes. At this stage (i.e., at 0 d), a group of five replicates was harvested to determine the biomass and Cu concentration of shoots and roots. The analyses of these containers provided references (starting point or "control") for either the hydroponic or soil experiment. The other containers with 3-wk-old rape plants were then used for either the hydroponic or soil experiment.

View larger version (46K): [in this window] [in a new window]   Fig. 1. Cropping device used (i) during the first step of the biotest (hydroponic preculture period) to obtain a dense mat of roots and (ii) during the hydroponic experiment.

Soil Experiment—Biotest for Assessing Soil Copper Bioavailability
This experiment corresponds to the second step of the biotest during which 21-d-old plants ("preculture") were transferred onto a contaminated soil for estimating the bioavailability and phytotoxicity of soil Cu. One day before being placed in contact with the plants for either 4 or 8 d, soil containers were prepared as follows: the lower part consisted of a 3-mm-thick PVC plate (7 by 5 cm) while the upper part was either a 1.1- or 3-mm-thick PVC plate that had the same dimensions and a central hole with a 32-mm i.d. An ashless filter paper (#40; Whatman, Maidstone, UK) was inserted between the two plates and connected to a solution reservoir to function as a wick (Fig. 2) . Before being poured in the soil containers, the soil had been incubated for 6 d in the growth chamber by adding 44 mL of solution to 200 g air-dried soil in a polyethylene bag. Two different bags were prepared: one with deionized water, the other one with a nutrient solution that had the same composition as the above-mentioned complete solution minus CuCl₂ and FeNaEDTA. The FeNaEDTA was omitted, as EDTA might have affected the speciation of Cu in the soil. Copper was not supplied so that soil Cu was the only source of Cu for the plants at this stage of the biotest. The assembled soil container was then filled with either 2.4 or 4.4 g of incubated, wet soil (treatments called 1 or 3 mm, respectively) and left for incubating for another day in the growth chamber. They were covered with an aluminium foil to prevent their dehydration and the development of algae. When filling the soil containers, some extra soil was added so that the soil layer was actually thicker than the hollow PVC plate, to ensure a good contact with the plant container at the last stage of the biotest. This means that the actual thickness of the soil layers was close to 1.5 to 2 and 3.5 to 4 mm. Nevertheless, these two treatments will thereafter be called 1 mm and 3 mm, respectively, which are the dimensions of the corresponding soil containers.

View larger version (52K): [in this window] [in a new window]   Fig. 2. Cropping device used during the second step of the biotest, that is, soil experiment with plants being grown on top of a thin layer of soil.

The design of this experiment consisted of two time durations (4 and 8 d) x two soil thicknesses (1 and 3 mm) x two nutrient treatments (nutrient solution and deionized water). For each of these eight treatments, five replicates were prepared by placing five containers with preculture plants onto five soil containers. Five additional soil containers were prepared for each treatment, covered with aluminium foil, and incubated in the same conditions as for the soil experiment (same procedure as above but without plants) to serve as "control" soils, to evaluate the root-induced changes occurring in the rhizosphere soil. In this experiment, as in the hydroponic experiment, the five replicates were not fully independent, as they were sharing the same reservoir of nutrient solution. As this was large compared with their requirements, negligible interactions between the replicates were expected and the replicates were assumed to be independent when conducting the statistical tests.

Plant and Soil Analyses
The roots and shoots were harvested at the end of the hydroponic preculture period (0 d), and after 14 d of hydroponic culture (in the "hydroponic experiment") or after 4 and 8 d of soil–plant contact (in the "soil experiment"). For each treatment, the five replicates were collected and analyzed separately. They were weighed fresh; shoots were oven-dried at 105°C and roots were frozen at -20°C. Copper bound to the outer root cell walls (apoplastic Cu) was determined as follows: a subsample of 0.8 g of fresh roots (after thawing) was shaken end over end with 40 mL of 0.001 M HCl during 3 min; then 360 µL of 1 M HCl were added to yield a final concentration of 0.01 M HCl. After shaking for another 5 min, the suspensions were filtered with an ashless filter paper (Whatman 40) and Cu was assayed by flame atomic absorption spectrometry (FAAS) (SpectrAA-600; Varian, Palo Alto, CA). Iwasaki et al. (1990) and preliminary experiments designed to measure K efflux from roots with a microelectrode showed that such experimental conditions (acid concentrations and duration of extraction) did not damage the plasma membrane of either fresh or frozen or thawn roots (which would have resulted in overestimating apoplastic Cu). Similar results were obtained for both fresh and frozen–thawn roots, which indicated that roots could be frozen before apoplastic Cu determination. The remaining portion of roots was oven-dried at 105°C. The plant roots and shoots were digested separately in a 1:1 mixture of hot, concentrated HNO₃ and HClO₄ (Association of Official Analytical Chemists, 1975). Copper in the digest was assayed by FAAS.

The soils were collected at harvest and oven-dried at 105°C. Two independent chemical extractions were performed with two subsamples of soils collected from either cropped soil containers or uncropped, control soil containers (without plants). Copper was extracted with 0.05 M CaCl₂ to estimate the exchangeable fraction of soil Cu (McLaren and Crawford, 1973). The amount of soil used in the present work was modified, but the soil to solution ratio was the same: 1 g of dry soil was shaken for 2 h with 5 mL of 0.05 M CaCl₂. The suspensions were then centrifuged at 15 000 x g for 3 min. The other extraction was done with DTPA. This method had been initially designed by Lindsay and Norvell (1978) for assessing the plant-available fraction of soil Cu in calcareous soils that were low or deficient in Cu. However, it has since been used also for Cu-contaminated soils (e.g., Beckett, 1989; Brun et al., 2001). Briefly, the extractant solution was composed of 0.005 M DTPA, 0.01 M CaCl₂, and 0.1 M triethanolamine (TEA) at pH 7.3. The CaCl₂ and TEA were added in the extractant solution to prevent CaCO₃ excessive dissolution and metal release (O'Connor, 1988). In comparison with the initial procedure, the amount of soil used here was much reduced to adapt to the small amount of soil collected from each soil container, but the soil to solution ratio was the same: 500 mg of dry soil was shaken for 2 h with 1 mL of extractant solution. The suspensions were then centrifuged at 15 000 x g for 3 min. Copper in all soil extracts was assayed by FAAS.

	RESULTS AND DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS AND DISCUSSION CONCLUSIONS REFERENCES

 
Hydroponic Experiment—Biotest for Assessing Copper Phytotoxicity
Plant Growth Response
The biomass of both shoots and roots showed a maximum at 5 µM and then decreased when Cu concentration further increased (Fig. 3a,b) . This decrease was significant, however, only for concentrations above 20 µM. For roots and shoots, no significant biomass difference was observed between 0.2, 5, and 20 µM. Similarly, no significant biomass differences were observed for concentrations between 80 and 200 µM (Fig. 3a,b). For shoots, at 200 µM the biomass had decreased by a factor of three relative to that at 0.2 µM and was equal to that of preculture plants, indicating that no growth had occurred. When considering biomass as a simple endpoint, the no observable adverse effect concentration (NOAEC), was thus lying somewhere between 20 and 80 µM. In comparison, for maize the NOAEC was lying between 0.16 and 1.6 µM Cu concentration as deduced from shoot and root growth measurements by Lexmond and van der Vorm (1981). This was found in their experiment that most closely compared with the present experiment in terms of solution composition (only NO₃–N supplied and pH at 5.6–6.1). However, a major difference was that they supplied Fe as FeSO₄ instead of FeNaEDTA and hence most of solution Cu was occurring as free Cu ion (90–95%). The NOAEC found for rape in the present work was ranging between 0.06 and 1.1 µM free Cu ion concentration, which thus compares fairly well with that deduced from Lexmond and van der Vorm (1981) for maize. Figures 3a and 3b also show that the EC₅₀, that is, the exposure concentration at which 50% of the maximum biomass is achieved, was about 80 µM for shoots and 50 µM for roots. This further shows a larger sensitivity of roots relative to shoots. An even more sensitive indicator than root growth might have been used, in light of the approach that has been developed for assessing Al toxicity, such as the relative elongation of selected roots (e.g., Kinraide and Parker, 1987; Parker, 1995; Calba et al., 1996). This indicator is more sensitive because decreased root elongation, the primary effect of Al rhizotoxicity, is often accompanied by some root thickening that somewhat compensates the ultimate effect on root growth as assessed via the whole biomass. This indicator recently proved applicable to evaluate the response of plant roots to Cu toxicity (Parker et al., 1998) and an EC₅₀ of about 0.3 µM of free Cu ion was found for wheat (Triticum aestivum L.), to compare with an EC₅₀ of about 0.25 µM for root biomass in the present work (equivalent to 50 µM total Cu). Comparison of the two approaches for an identical plant species grown in identical hydroponic conditions would be needed to evaluate which is the best suited (most sensitive) method for measuring Cu rhizotoxicity.

View larger version (28K): [in this window] [in a new window]   Fig. 3. Dry biomass of shoots (a) and roots (b) expressed as g per container (per 10 plants), and Cu concentrations (mg kg-1 dry matter) of shoots (c) and roots (d) as a function of Cu concentration in the nutrient solution (µM) in the hydroponic experiment. Open symbols stand for values of preculture plants. Corresponding free Cu ion concentrations (µM) as calculated from the SOILCHEM computer code (Sposito and Coves, 1988) are also indicated. Mean values with different letters are significantly different (p < 0.05) as measured by the Ryan–Einot–Gabriel–Welsch–Range test.

The concentration of Cu in the roots might be considered an even more sensitive endpoint, as several authors have pointed out that Cu can accumulate in the roots to a large extent while Cu concentration in plant shoots remains unaltered (Mitchell et al., 1978; Lexmond and van der Vorm, 1981; Marschner, 1995; Brun et al., 2001). Indeed, much larger Cu concentrations have been found in the roots, ranging between 24 and 24 700 mg kg^-1 DM, than in the shoots (Fig. 3c,d). Although constantly increasing with increasing Cu concentration in the nutrient solution, they were not significantly different for concentrations ranging from 0.2 to 80 µM. Root Cu concentration peaked at 100 µM, reaching 24 700 mg kg^-1 DM (Fig. 3d). Because of much larger variabilities in the measured values for roots, a significant increase in root Cu concentration was found only at a solution Cu concentration of 100 µM. It thus appears for this plant species that this indicator is not best suited for an early detection of Cu phytotoxicity. It is nonetheless interesting to point out that when Cu concentration in the shoots increased from 6 mg kg^-1 DM (marginal–adequate range) at 0.2 µM, to 16 mg kg^-1 DM (critical range) at 5 µM, and to 49 mg kg^-1 DM (phytotoxic range) at 20 µM, Cu concentrations in the roots increased in the meantime from 24 to 65 and 533 mg kg^-1 DM, respectively. When plant growth started to be significantly altered (at 80 µM), Cu concentration had reached about 400 mg kg^-1 DM in the shoots and more than 5000 mg kg^-1 DM in the roots, which is many-fold higher than phytotoxic levels (Fig. 3c). The present experiment thus clearly confirms that Cu preferentially accumulates in the roots and more so when solution Cu concentration increases, in agreement with former results obtained for maize (Mocquot et al., 1996). Shoot to root Cu concentration ratios decreased from 0.25 at 0.2 µM to 0.07 at 80 µM, which compares fairly well with the observations for maize by Lexmond and van der Vorm (1981). However, a striking difference between the two species was the much larger increase in Cu concentration in the shoots of rape exposed to slightly phytotoxic concentrations, relative to maize. In addition, the ratio of shoot to root bioaccumulated Cu in rape (3.4, 2.5, 1.3, and 1.1 for 0.2, 5, 20, and 80 µM Cu concentrations, respectively) was about 5- to 10-fold larger than that deduced from the results obtained for maize by Lexmond and van der Vorm (1981). Rape was thus more prone than maize to translocate a substantial proportion of Cu into its shoots. This propensity to translocate metals to shoots has been observed in many species of the Brassica genus and offers some prospects for using these species for phytoextraction purposes as shown, for instance, by Ebbs et al. (1997).

Soil Experiment—Biotest for Assessing Copper Bioavailability
Plant Growth and Copper Concentration in Plant Parts
The biomass of both shoots and roots was greater than that of rape harvested at the end of the preculture (Table 1). This increase was, however, significant only for plants supplied with nutrient solution for 8 d. Plant growth did not seem to be affected by soil layer thickness. The poor growth of plants relative to preculture plants (Table 1) might be interpreted at first sight as a response to Cu phytotoxicity. Plant growth was notably reduced when deionized water was supplied (Table 1). However, it is likely that the growth was restricted as a consequence of a shortage of nutrients supplied by the small amount of soil when deionized water was used. Indeed, supplying nutrient solution alleviated those nutrient deficiencies and significantly improved plant growth (after 8 d at least, see Table 1).

View larger version (35K): [in this window] [in a new window]   Fig. 4. Concentrations (a) and amounts (b) of Cu in plant shoots and roots expressed as mg kg-1 dry matter and µg per container (per 10 plants), respectively. NS and H2O stand for nutrient solution and deionized water treatments, respectively. Percentages of total root Cu bound to root cell walls are indicated in (a). Mean values with different letters are significantly different (p < 0.05) as measured by the Ryan–Einot–Gabriel–Welsch–Range test.

Root Cu concentrations did not exceed 127 mg kg^-1 DM when deionized water was supplied and half of this (about 60 mg kg^-1 DM) when nutrient solution was supplied (Fig. 4a). These values remain in the order of Cu concentrations observed in the roots of hydroponically grown plants supplied with nonphytotoxic Cu concentrations (24 mg kg^-1 DM at 0.2 µM Cu, 65 mg kg^-1 DM at 5 µM Cu). This is an additional indication that this soil did not induce any Cu toxicity in rape plants. These results confirm that when adequate shoot Cu concentrations (unchanged relative to uncontaminated soils) are found, fairly large Cu concentrations can occur in roots of plants grown in Cu-contaminated calcareous soils, although much larger concentrations have been found in roots in earlier reports (Mitchell et al., 1978; Brun et al., 2001). There is thus some prospect for using the concentration of Cu in roots as an early indicator of plant contamination for soil-grown plants, as suggested by Mitchell et al. (1978). However, for this purpose, references on the critical levels of root Cu that are indicative of a toxicity need to be established. In addition, the present experiment showed that a substantial proportion (26–78%) of total root copper was bound to the cell wall fraction (see apoplastic Cu in Fig. 4a), which is in line with previous results (Iwasaki et al., 1990; Parker et al., 1998). Lower proportions of apoplastic Cu were found in plants supplied with nutrient solution, possibly because of increased competition with cations (such as Ca or Mg) supplied by the solution, as previously shown for wheat by Parker et al. (1998). As stressed by these authors, there is a need to better understand the processes involved in the rhizotoxicity of Cu and its relationship with the partitioning of root Cu between the apoplasm and symplasm compartments, similar to research completed for Al rhizotoxicity. This is a prerequisite for selecting which of total, symplasmic, or apoplasmic Cu in roots is the best indicator of the Cu toxicity in soil-grown plants.

The absence of any Cu phytotoxicity in the present soil, in spite of its large total Cu content, might partly be due to its calcareous nature, as Cu is known to be more bioavailable in acidic than neutral or calcareous soils (McLaren and Crawford, 1973; Mitchell et al., 1978; Alloway, 1995). Alternatively, it could be due to the procedure used in the present biotest, which is based on a rather short duration of exposure to the contaminated soil (4–8 d). This might have resulted in a dilution effect in the 3-wk-old plants used in this biotest. No periods of contact longer than 4 to 8 d were considered here, because a major advantage of the present biotest relative to conventional pot experiments was to provide a short-term, rapid, and thus reasonably inexpensive test. There is thus a need to investigate other soils with different properties, including noncalcareous soils and soils that are known to be responsible for phytotoxicity.

Uptake and Bioavailability of Copper
Little or no significant differences in amounts of bioaccumulated Cu were found between the various treatments (Fig. 4b). The uptake of Cu, however, increased when nutrient solution was supplied compared with deionized water. As Cu concentration tended to decrease in the meantime (Fig. 4a), the enhanced uptake of Cu was due to the increased growth of plants supplied with nutrient solution relative to those supplied with deionized water. For similar reasons, Cu uptake also increased with increasing soil–plant contact period. Although twice as much soil (and hence Cu) was supplied in the 3-mm treatment relative to the 1-mm treatment, no significant increase in Cu uptake was observed. Copper uptake by rape per unit of soil mass supplied, which is an estimate of soil Cu bioavailability (Table 2), was deduced from the previous figures (amounts of bioaccumulated Cu, Fig. 4b) by subtracting the initial amount of Cu in the preculture plants and expressing the final result relative to the mass of dry soil supplied (which differed between the two soil thickness treatments). Bioavailable Cu amounted to 0.5 to 2% of total soil Cu. The effect of time duration was clearly visible (Table 2). Again, soil thickness (i.e., the amount of soil supplied) had little significant effect on Cu bioavailability (Table 2). This indicates that a major proportion of the amount of Cu taken up by rape plants was derived from the first millimeter of the rhizosphere, which is not surprising owing to the poor mobility of Cu in soils. In that respect, working with 1 mm would be justified, especially if the biotest also aims at investigating the root-induced changes that occur in the rhizosphere, as was the case in the present work. Rhizosphere effects typically occur as gradients as a function of the distance from the root surface (Hinsinger, 1998). They might therefore be increasingly diluted and eventually become hardly detectable with increasing distance from the roots (i.e., investigated soil thickness).

	CONCLUSIONS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS AND DISCUSSION CONCLUSIONS REFERENCES

 
The present work aimed at (i) defining the responses of oilseed rape to Cu phytotoxicity as assessed in hydroponic conditions and (ii) optimizing the operational conditions of a novel ecotoxicological test designed for evaluating soil bioavailability and phytotoxicity of metals such as Cu. The NOAEC ranged between 5 and 20 µM total Cu (0.01 and 0.06 µM free Cu ion). The soil used for investigating the operational conditions of the soil-based biotest did not induce any significant phytotoxicity, in spite of its large total Cu content. It can nonetheless be concluded that the best conditions for conducting this biotest are the following: (i) the duration should be 8 rather than 4 d to be able to use plant growth as an indicator of phytotoxicity, (ii) the thickness of the soil layer (container) should be 1 mm (roughly 2 g dry soil per container) rather than 3 mm, as most of the bioavailable Cu is derived from this portion of the rhizosphere and (iii) nutrient solution should be supplied instead of deionized water, for a better comparison of soils with different nutrient status.

Compared with glasshouse pot experiments that are conventionally used in terrestrial ecotoxicology, this biotest has several definitive advantages: (i) easy recovery of all the roots, (ii) cost effectiveness, as it lasts for only 8 d and does not require manual watering, (iii) small size that is adapted for conducting a large number of treatments and replicates over a small surface area, that is, in growth chambers that can provide standardized and reproducible climatic conditions. The proposed biotest, however, suffers of several disadvantages, especially related to the particular arrangement of soil and roots: (i) the roots are not growing within the soil and may therefore be less exposed to toxicities, (ii) some plant species are poorly adapted as they hardly form a dense, planar mat of roots (e.g., species with a strong, thick tap root), (iii) the small ratio of soil to plant requires the addition of a nutrient solution with associated, potential artifacts; therefore, the soil is almost water-saturated, being connected to a large reservoir of nutrient solution that may affect the cation exchange, complexation, and redox equilibria. This biotest would now need to be further evaluated for a range of soils varying in properties and levels of metal contamination. It should also be applied to conventional pot experiments that are currently being used in terrestrial ecotoxicology and, ultimately, to in situ measurements. In that respect, it is interesting to point out that a slightly different biotest derived from the same original method of Niebes et al. (1993) has been recently validated against a conventional pot experiment for evaluating the bioavailability of ¹³⁷Cs in a range of soils (Kruyts et al., 2000; Thiry et al., 2000). This biotest has also been used for studying chemical changes occurring in the rhizosphere that ultimately affect the bioavailability of trace elements to plants (Guivarch et al., 1999; Hinsinger, 2001). It is a unique tool for assessing responses of roots to metal toxicities in soil-grown plants, as it provides an easy access to the roots and hence to sensitive endpoints (e.g., root elongation, as shown for Al by Calba et al., 1996) or biomarkers (biochemical responses) of metal toxicities that would express in the roots. The proposed biotest thus can be seen as a research tool that also accommodates the requirements of standardized ecotoxicological testing procedures.

	ACKNOWLEDGMENTS

 
The authors thank Dr. P. Andrieux for providing access to the experimental vineyard watershed, G. Souche and M. Clairotte for their help in sampling the soil on this site, and Dr. C. Plassard for designing and conducting the K efflux measurements. The help provided by Professor P. Schwab and anonymous referees for improving the manuscript is gratefully acknowledged. This work was made possible by the financial support of the French Ministry of Environment, within the framework of its PNETOX program.

	REFERENCES

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS AND DISCUSSION CONCLUSIONS REFERENCES

 

• Alloway, B.J. 1995. Heavy metal in soils. 2nd ed. Blackie Academic and Professional, London.
• American Society for Testing and Materials. 1994. Standard practise for conducting early seedling growth tests. E 1598-94. ASTM, Philadelphia, PA.
• Association Française de Normalisation. 1994. Qualité des Sols—Recueil de normes Françaises. AFNOR, Paris.
• Association of Official Analytical Chemists. 1975. Official methods of analysis. 12th ed. AOAC, Washington, DC.
• Beckett, P.H.T. 1989. The use of extractants in studies on trace metals in soils, sewage sludges and sludge-treated soils. Adv. Soil Sci. 9:143–179.
• Brun, L.A., J. Maillet, P. Hinsinger, and M. Pépin. 2001. Evaluation of copper availability to plants in copper-contaminated vineyard soils. Environ. Pollut. 111:293–302.[Medline]
• Calba, H., B. Jaillard, P. Fallavier, and J.C. Arvieu. 1996. Agarose as a suitable substrate for use in the study of Al dynamics in the rhizosphere. Plant Soil 178:67–74.
• Cushman, R., and R.D. Meyer. 1990. Improving the utilization of non-traditional agricultural products through coordination during the registration process. Commun. Soil Sci. Plant Anal. 21:1531–1540.
• Ebbs, S.D., M.M. Lasat, D.J. Brady, J. Cornish, R. Gordon, and L.V. Kochian. 1997. Phytoextraction of cadmium and zinc from a contaminated soil. J. Environ. Qual. 26:1424–1430.[Abstract/Free Full Text]
• Guivarch, A., P. Hinsinger, and S. Staunton. 1999. Root uptake and distribution of radiocesium from contaminated soil and the enhancement of Cs adsorption in the rhizosphere. Plant Soil 211:131–138.
• Hinsinger, P. 1998. How do plant roots acquire mineral nutrients? Chemical processes involved in the rhizosphere. Adv. Agron. 24:225–265.
• Hinsinger, P. 2001. Bioavailability of trace elements as related to root-induced chemical changes in the rhizosphere. p. 25–41. In G.R. Gobran, W.W. Wenzel, and E. Lombi (ed.) Trace elements in the rhizosphere. CRC Press, Boca Raton, FL.
• International Organization for Standardization. 1999. Soil quality—Guidance on the ecotoxicological characterisation of soils and soil materials. Guidelines no. ISO TC 190/SC 7 and ISO/DIS 15799. ISO, Geneva, Switzerland.
• Iwasaki, K., K. Sakurai, and E. Takahashi. 1990. Copper binding by the root cell walls of Italian ryegrass and red clover. Soil Sci. Plant Nutr. (Tokyo) 36:431–439.
• Jaillard, B., C. Plassard, and P. Hinsinger. 2003. Measurement of H⁺ fluxes and concentrations in the rhizosphere. p. 231–266. In Z. Rengel (ed.) Handbook of soil acidity. Marcel Dekker, New York.
• Kapustka, L.A. 1997. Selection of phytotoxicity tests for use in ecological risk assessments. p. 515–548. In W. Wang, J.W. Gorsuch, and J.S. Hughes (ed.) Plants for environmental studies. CRC Press, Boca Raton, FL.
• Kinraide, T.B., and D.R. Parker. 1987. Cation amelioration of aluminum toxicity in wheat. Plant Physiol. 84:546–551.
• Kruyts, N., Y. Thiry, and B. Delvaux. 2000. Respective horizon contributions to cesium-137 soil to plant transfer: A rhizospheric experimental approach. J. Environ. Qual. 29:1180–1185.[Abstract/Free Full Text]
• Kuchenbuch, R., and A. Jungk. 1982. A method for determining concentration profiles at the soil–root interface by thin slicing rhizospheric soil. Plant Soil 68:391–394.
• Lebourg, A., T. Sterckeman, H. Ciesielski, N. Proix, and A. Gomez. 1996. Estimation of soil trace metal bioavailability using unbuffered salt solutions: Degree of saturation of polluted soil extracts. Environ. Technol. 19:243–252.
• Lexmond, T.M. 1980. The effect of soil pH on copper toxicity to forage maize grown under field conditions. Neth. J. Agric. Sci. 28:164–184.
• Lexmond, T.M., and P.D.J. van der Vorm. 1981. The effect of pH on copper toxicity to hydroponically grown maize. Neth. J. Agric. Sci. 29:217–238.
• Lindsay, W.L., and W.A. Norvell. 1978. Development of DTPA soil test for zinc, iron, manganese, and copper. Soil Sci. Soc. Am. J. 42:421–428.
• Lorenz, S.E., R.E. Hamon, and S.P. McGrath. 1994. Differences between soil solutions obtained from rhizosphere and non-rhizosphere soils by water displacement and soil centrifugation. Eur. J. Soil Sci. 45:431–438.
• Marschner, H. 1995. Mineral nutrition of higher plants. 2nd ed. Academic Press, San Diego, CA.
• McLaren, R.G., and D.V. Crawford. 1973. Studies on soil copper. II. The specific adsorption of copper by soils. J. Soil Sci. 24:443–452.
• McLaughlin, M.J., E. Smolders, and R. Merckx. 1998. Soil–root interface: Physicochemical processes. p. 233–277. In Soil chemistry and ecosystem health. SSSA Spec. Publ. 52. SSSA, Madison, WI.
• Mitchell, G.A., F.T. Bingham, and A.L. Page. 1978. Yield and metal composition of lettuce and wheat grown on soils amended with sewage sludge enriched with cadmium, copper, nickel and zinc. J. Environ. Qual. 7:165–171.[Abstract/Free Full Text]
• Mocquot, B., J. Vangronsveld, H. Clijsters, and M. Mench. 1996. Copper toxicity in young maize (Zea mays L.) plants: Effect on growth, mineral and chlorophyll contents, and enzyme activities. Plant Soil 182:287–300.
• Morel, C., and P. Hinsinger. 1999. Root-induced modifications of the exchange of phosphate ion between soil solution and soil solid phase. Plant Soil 211:103–110.
• Niebes, J.F., P. Hinsinger, B. Jaillard, and J.E. Dufey. 1993. Release of non exchangeable potassium from different size fractions of two highly K-fertilised soils in the rhizosphere of rape (Brassica napus cv Drakkar). Plant Soil 155/156:403–406.
• O'Connor, G.A. 1988. Use and misuse of the DTPA soil test. J. Environ. Qual. 17:715–718.[Abstract/Free Full Text]
• Parker, D.R. 1995. Root growth analysis: An underutilised approach to understanding aluminium rhizotoxicity. Plant Soil 171:151–157.
• Parker, D.R., J.F. Pedler, D.N. Thomason, and H. Li. 1998. Alleviation of copper rhizotoxicity by calcium and magnesium at defined free metal-ion. Soil Sci. Soc. Am. J. 62:965–972.[Abstract/Free Full Text]
• Reuter, D.J., and J.B. Robinson. 1997. Plant analysis: An interpretation manual. CSIRO Publishing, Collingwood, VIC, Australia.
• Singh, S.P., and N.G. Rakipov. 1988. Effects of Cd, Cu and Ni on barley and their removal from the soil. Int. J. Environ. Stud. 31:291–295.
• Sposito, G., and J. Coves. 1988. SOILCHEM: A computer program for the calculation of chemical speciation in soils. The Univ. of California, Berkeley.
• Thiry, Y., N. Kruyts, and B. Delvaux. 2000. Respective horizon contributions to cesium-137 soil to plant transfer: A pot experiment approach. J. Environ. Qual. 29:1194–1199.[Abstract/Free Full Text]

Related articles in JEQ:

This Issue in Journal of Environmental Quality

JEQ 2003 32: 745-750. [Full Text]  

This Article Abstract Figures Only Full Text (PDF) Free Alert me when this article is cited Alert me if a correction is posted Services Related articles in JEQ Similar articles in this journal Similar articles in ISI Web of Science Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via ISI Web of Science (7) Citing Articles via Google Scholar Google Scholar Articles by Chaignon, V. Articles by Hinsinger, P. Search for Related Content PubMed PubMed Citation Articles by Chaignon, V. Articles by Hinsinger, P. GeoRef GeoRef Citation Agricola Articles by Chaignon, V. Articles by Hinsinger, P. Related Collections Lysimeter/Rhizosphere Studies Ecological Risk Assessment Heavy Metals Plant and Soil Interactions Soil Pollution