Plant Soil (2008) 302:233–247 DOI 10.1007/s11104-007-9475-2
REGULAR ARTICLE
Rhizosphere microbial community and hexachlorocyclohexane degradative potential in contrasting plant species P. S. Kidd & A. Prieto-Fernández & C. Monterroso & M. J. Acea
Received: 31 May 2007 / Accepted: 2 November 2007 / Published online: 12 December 2007 # Springer Science + Business Media B.V. 2007
Abstract The organochlorine 1,2,3,4,5,6 hexachlorocyclohexane (HCH) is a broad-spectrum insecticide that was used on a large-scale worldwide. The soil– plant–microbe system and its influence on HCH biodegradation are evaluated. A greenhouse experiment was designed to evaluate HCH dissipation and several microbial parameters among rhizosphere and bulk soil of two contrasting plants, Cytisus striatus (Hill) Rothm and Holcus lanatus L. Plants were grown for 180 days in three treatments: uncontaminated soil (control), uncontaminated soil inoculated with soil (3% w/w) from a HCH-contaminated site (INOC), and uncontaminated soil inoculated with soil (3% w/w) from the HCH-contaminated site and artificially contaminated to obtain 100 mg HCH kg−1 dry soil (100HCH-INOC). At harvest, plant biomass, soil water-extractable organic C, pH and Cl concen-
Responsible Editor: Fanjie J. Zhao. P. S. Kidd (*) : A. Prieto-Fernández : M. J. Acea Instituto de Investigaciones Agrobiológicas de Galicia, CSIC, Apdo. 122, Santiago de Compostela 15780, Spain e-mail:
[email protected] C. Monterroso Departamento de Edafología y Química Agrícola, Facultad de Biología, Universidad de Santiago de Compostela, Santiago de Compostela 15782, Spain
tration, rhizosphere microbial densities (total heterotrophs, ammonifiers, amylolytics) and C substrate utilization patterns, and degradation of α-, β-, δ- and γ-HCH isomers were determined in bulk and rhizosphere soils. Soil solution Cl concentration was determined every 30 days throughout the entire growth period. Results demonstrate that both Cytisus striatus and Holcus lanatus can grow in soils with up to 100 mg HCH kg−1. An enhanced degradation of α-HCH, but not β- or δ-HCH, was observed in the rhizosphere. Significant changes in the microbial densities were observed between bulk and rhizosphere soils of Cytisus, and an increase in C source utilization indicated changes in community level physiological profiles (CLPP) in the rhizosphere of this species when grown in contaminated soils. HCH dissipation was also greater in soils planted with this species. In accordance, increases in soil extractable C, Cl concentration and acidity were greater at the rhizosphere of Cytisus. Concentration of Cl in soil solutions also indicates greater HCH dechlorination in soils planted with Cytisus than Holcus. Results suggest that phytostimulation of bacteria present or added to soil is a promising approach to cleaning HCHcontaminated sites, and especially for biodegradation of α-HCH. Keywords Ammonifiers . Amylolytics . Hexachlorocyclohexane isomers . Heterotrophs . Lindane . Phytoremediation . Rhizosphere
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Introduction Persistent organic pollutants (POPs) (such as chlordane, dioxins, 1,1,1-trichloro-2,2-bis(4-chlorophenyl) ethane (DDT), polychlorinated biphenyls (PCBs) and polycyclic aromatic hydrocarbons (PAHs)) are of great environmental concern due to their recalcitrance, global transport and distribution, and toxicity (Wania and Mackay 1996). Recent studies demonstrate significantly enhanced dissipation and/or mineralisation of POPs at the root–soil interface or rhizosphere (rhizodegradation) (Anderson et al. 1993; Anderson and Coats 1995; Kuiper et al. 2004; Chaudhry et al. 2005; Krutz et al. 2005). This rhizosphere effect is generally attributed to an increase in microbial density, diversity and/or metabolic activity due to the release of plant root exudates, mucigel and root lysates (enzymes, amino acids, carbohydrates, low-molecular-mass carboxylic acids, flavonones and phenolics; Curl and Truelove 1986). Rhizodeposits not only provide a nutrient-rich habitat for microorganisms but can potentially enhance biodegradation in different ways: they may facilitate the co-metabolic transformation of pollutants with similar structures, induce genes encoding enzymes involved in the degradation process, increase contaminant bioavailability (surfactant activity), and/or selectively increase the number and activity of pollutant degraders in the rhizosphere (Anderson and Coats 1995; Schnoor et al. 1995; Nichols et al. 1997; Burken and Schnoor 1998; Miya and Firestone 2001; Shaw and Burns 2003). Plants can also improve the physical and chemical properties of contaminated soil, and increase contact between the root-associated microorganisms and the soil contaminants. Terpenes (such as cymene, α-pyrene and α-terpinene) and phenolics (such as salicylate) have been shown to induce biphenyl dioxygenase in PCB-degrading bacteria (Chen and Aitken 1999). While phenolic compounds such as naringen, coumarin or catechin, released by roots of certain plants have been shown to support the growth of rhizospheric PCB-degrading bacteria (Donnelly et al. 1994; Chaudhry et al. 2005). In contrast, phenanthrene-degrading activity of Pseudomonas putida ATCC 17484 was repressed after incubation with plant root extracts (Rentz et al. 2004). However, these authors suggested that the enhanced microbial growth on rhizodeposits is likely to compensate for this partial repression since a larger microbial population leads to a faster degradation rate.
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Although the mechanisms involved are not always determined, rhizosphere-enhanced biodegradation has been demonstrated for a wide range of organic pollutants. Enhanced mineralisation of 2,4dichlorophenoxyacetic acid (2,4-D) was shown in soil collected from the rhizosphere of Trifolium pratense (Shaw and Burns 2005). Chekol et al. (2002) showed enhanced transformation of the explosive, trinitrotoluene (TNT) by the forage grasses Phalaris arundinacea and Panicum virgatum. Concentrations of the organochlorine p,p′-DDE (2,2-bis(p-chlorophenyl) 1,1-dichloroethylene), a metabolite of DDT, were significantly reduced in the rhizosphere of field-grown zucchini, pumpkin and spinach compared to bulk soil (White 2001). Numerous studies have demonstrated increased degradation of petroleum hydrocarbons (such as phenanthrene, benzo[a]pyrene, benzo[a]anthracene, chrysene, hexadecane, benzene, toluene etc.) as a result of modified microbial activity in the rhizosphere of grasses and legumes (Nichols et al. 1997; Miya and Firestone 2001; Corgié et al. 2003; Muratova et al. 2003; Phillips et al. 2006). The organochlorine 1,2,3,4,5,6 hexachlorocyclohexane (HCH) is a broad-spectrum insecticide that was used on a large-scale worldwide since the 1940s, and is available in two formulations: technical-grade HCH (a mixture of different isomers, mainly α- (60– 70%), β- (5–12%), γ- (10–15%), and δ-HCH (6–10%)) and lindane (almost pure γ-HCH). Lindane, the only isomer with insecticidal properties, is isolated from technical-grade HCH by crystallization (Turnbull 1996). All HCH isomers are acutely toxic to mammals, due to their mutagenic, teratogenic and carcinogenic properties (Willett et al. 1998). Due to its widespread use, and strong persistence, residues of lindane and other HCH isomers are found all over the world in air, water, sediments and soils (Willett et al. 1998; Schwitzguébel et al. 2006). Although nowadays its use is restricted or completely banned in most countries, it continues to pose serious environmental and health concerns, particularly on highly contaminated former production or dumping sites (Lal et al. 2006; Schwitzguébel et al. 2006). Due to its physicochemical characteristics, HCH isomers tend to sorb to organic material in the environment and have a low bioavailability (Rodríguez Garrido 2003). The low water solubility and high hydrophobicity (logKOW 3.7–4.1; Willett et al. 1998) of HCH isomers make their uptake and translocation
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within the plant unlikely. HCHs are more likely to be strongly adsorbed on the root epidermis (Calvelo Pereira et al. 2006; Schwitzguébel et al. 2006). In contrast, several bacterial strains (predominantly Sphingomonas strains) capable of degrading HCH isomers under aerobic conditions have been isolated from contaminated soils (Imai et al. 1989; Böltner et al. 2005; Mohn et al. 2005; Phillips et al. 2005). The phytostimulation of rhizospheric HCH-degrading microorganisms is therefore likely to be a successful strategy for the remediation of HCH-contaminated soils. Further studies evaluating the soil–plant–microbe system and its influence on HCH biodegradation are necessary so as to better explore and exploit an undoubtedly huge potential. The objective of the present research was to study the effect of two plant species (Cytisus striatus and Holcus lanatus) on the rhizosphere microflora and its degradative potential in response to soil contamination by isomers of hexachlorocyclohexane (α-HCH, β-HCH, δ-HCH and γ-HCH). There is a lack of information about the response of different physiological groups of bulk and rhizosphere soil microorganisms to HCH contamination.
Materials and methods Plant species, preparation of soils and greenhouse pot experiments Common velvet grass (Holcus lanatus L. (Gramineae)) and Portuguese broom (Cytisus striatus (Hill) Rothm. (Leguminosae)) were chosen for this study because they are found growing in a HCH-contaminated area in Porriño (Pontevedra, NW Spain) where residues from lindane fabrication were disposed of during the 1950s and 1960s. Soil concentrations of ∑-HCH (calculated as the sum of α-HCH, β-HCH, δ-HCH and γ-HCH) at this site typically range between 2 and 100 mg kg−1, although at some local points concentrations of up to 20,000 mg kg−1 are found (Concha-Graña et al. 2006). Cytisus striatus is a well-known leguminous plant which ensures nitrogen-supply to the soil microflora. Grass species and leguminous plants have been shown to be suitable for rhizoremediation (Kuiper et al. 2004). The seeds of both species were collected from the contaminated (Porriño) site in July 2005.
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For the pot experiment, a granite derived soil, with no prior exposure to HCHs, was collected under shrub vegetation (predominantly Cytisus and Ulex sp.) close to the sea from the Pontevedra region (NW Spain). The soil has a pH of 5.5, 2.4% organic matter content, cation exchange capacity (CEC) of 8.7 cmolc kg−1 and 80.2% sand, 0.9% silt and 18.9% clay content. The soil was air-dried, ground and passed through a 5-mm sieve. The soil was limed with agricultural limestone (applied at rates equivalent to 2,500 kg CaCO3 ha−1), and fertilised at rates equivalent to 150 kg P ha−1, 200 kg K ha−1 with KH2PO4 and 100 kg N ha−1 with NH4NO3 before the experiment. Fertilisers were thoroughly mixed with the soil, and soils were adjusted to 80% of their water holding capacity and left for 4 weeks to attain equilibrium. For HCH application, residues produced during the fabrication of γ-HCH were obtained from a former factory in Porriño and ground in a mortar to pass a 1-mm sieve. The mean composition of the residue was: 77% α-HCH, 16% β-HCH, 5% δ-HCH, and 2% γ-HCH (Calvelo Pereira et al. 2006). Appropriate amounts of lindane residues were dissolved in 50-ml acetone and thoroughly mixed with sand. After mixing, the solvent was allowed to evaporate for 24 h and the sand was subsequently mixed with the treatment soils (at field capacity). Non-contaminated soils received an equal volume of sand and acetone (evaporated prior to mixing with soils). For inoculation, top soil was collected from the contaminated (Porriño) site and thoroughly mixed with the treatment soils at 3% w/w (at field capacity). Previous studies showed that degradation of HCH isomers was not detectable in the absence of microorganisms from the contaminated site (unpublished data). Hexachlorocyclohexanedegrading populations have been shown to be clearly associated with HCH-contaminated soils (Mohn et al. 2005). Plants were grown in ten replicated 1,000 g pots and three different treatments were prepared: (a) uncontaminated soil (control), (b) uncontaminated soil inoculated with soil from the contaminated site (INOC), or (c) uncontaminated soil inoculated with soil from the contaminated site and artificially contaminated to obtain a total of 100 mg HCH kg−1 dry soil (100 HCH-INOC). The final level of ∑-HCH in the INOC soil (due to the presence of HCH in the soil used for inoculation purposes) was 39 mg/kg (49% α-HCH, 44% β-HCH, 3% δ-HCH and 4% γHCH), and in the 100 HCH-INOC soil was 110 mg/kg
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(57% α-HCH, 38% β-HCH, 3% δ-HCH and 2% γ-HCH). Each pot was planted with either 30-day-old seedlings of Cytisus or 7-day-old seedlings of Holcus, which were previously grown on a potting compost/ sand mixture and thoroughly rinsed in deionised water before transplanting. Plants were grown for 180 days in a greenhouse, and watered as required. An additional four replicate pots per treatment were left unplanted. At the end of the experiment, plants were harvested, rhizosphere soil was separated, and shoot and root dry weights were determined after air drying. The whole soil and root system was gently crushed and loosely held soil was separated by shaking. This is referred to as the bulk soil. The remaining tightly held soil was considered rhizosphere soil, and was removed by shaking and/or gently crushing in a plastic bag after a brief period of airdrying. To the extent possible, root debris included in the collected rhizosphere soil were removed using tweezers or by sieving. Fresh soil samples were used for microbial analyses and dry soil samples for physico-chemical analyses and HCH determination. As a control, uncontaminated soil was placed in glass conical flasks with screw caps and sterilised by autoclaving three times (40 min, 121°C) with 24 h intervals. Half of the flasks were contaminated to obtain a total of 100 mg HCH kg −1 dry soil (following the same methods as above but lindane residues dissolved in acetone were filter-sterilised before mixing with sand (0.22 μm Millex® LG sterile filters)). Soils were wet with sterilised deionised water at 80% field capacity and flasks were incubated at 25°C for 49 days. Microbial analyses of bulk and rhizosphere soil samples Culturable heterotrophic bacteria, and the ammonifying and amylolytic populations were determined in bulk and rhizosphere soils by the most probable number (MPN) technique, as follows. Five grams of rhizosphere/bulk soil were suspended in 45 ml sodium hexametaphosphate solution (1%) and shaken for 30 min in an end-over-end shaker. These soil suspensions were diluted in 10-fold series and 50 μl aliquots were used to inoculate microtiter plates containing selective liquid media (150 μl/well). Five wells were
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inoculated per suspension-dilution. Heterotrophic population was estimated in yeast-extract medium (1.0 g yeast extract, 1.0 g glucose, 0.5 g KNO3, 0.2 g MgSO4.7H2O, 0.5 g K2HPO4, 0.1 g CaCl2, 0.1 g NaCl, 0.01 g FeCl3, in 1 l deionised water) plus oligoelements (1.5 mg FeSO4·7H2O, 0.3 mg H3BO3, 0.19 mg CoCl2, 0.1 mg MnCl2·4H2O, 0.08 mg ZnSO4.7H2O, 0.02 mg CuSO4·5H2O, 0.036 mg NaMoO4, 0.024 mg NiCl2·6H2O). Amylase-producers were cultured in Winogradsky’s saline medium (containing 0.25 g K2HPO4, 0.1 g MgSO4·7H2O, 1.0 g NH4NO3, 0.1 g NaCl, 2 mg Fe2(SO4)3·7H2O, 2 mg MnSO4 and 1.5 g soluble starch in 1 l deionised water) plus oligoelements (as above). Ammonifiers were evaluated in Winogradsky`s saline solution and Lasparagine (0.2 g l−1) as the only N and C source. The redox dye, tetrazolium violet (2,5-Diphenyl-3-(αnaphthyl)tetrazolium chloride (TV), 15 mM) was used to indicate growth, and was added to media using the ratio 1:100 (v/v, TV:media). Tetrazolium violet serves as an indicator of dehydrogenase activity, forming a deep purple precipitate upon reduction by electrons flowing through the electron transport system and by superoxide radicals (Kennedy 1994). All microbial groups were cultured at pH 6.8 and counts were performed after 1 and 3 weeks of incubation at 25°C. Each MPN was determined from the appropriate table, corrected for the initial dilution and inoculant volume, and expressed as log10MPN g−1 dry soil. The rhizosphere effect (R/B) was calculated for each physiological group as the ratio of the number (MPN g−1 soil) of microorganisms in the rhizosphere over the number (MPN g−1 soil) of microorganisms in bulk soil. The ability of microbial communities to utilise different C substrates was determined (community level physiological profiles (CLPP) analysis). Carbon substrate microtiter (MT) plates were made up by selecting 28 carbon sources (Table 1) from those included in EcoplateTM from Biolog Inc. or recommended by Kennedy (1994). Individual carbon source stock solutions were prepared at a concentration of 10%, passed through a sterile filter and added to sterile saline medium (1.75 g K2H2PO4, 0.5 g KHPO4, 0,582 g NH4Cl and 0.25 g MgSO4·7H2O in 1 l deionised water, adjusted to pH 7.0) using the ratio 1:100 (v/v, C source:saline medium). The indicator TV was added as above (1:100 TV:media (v/v)). Each well of microtiter plates was filled by dispensing
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Table 1 List of carbon sources used in metabolic profiling of microbial communities Chemical group
Carbon substrate
Carbohydrates
D-(+)
Carboxylic acids
Amino acids
Amines Polymers
Miscellaneous
cellobiose α-lactose β-methyl D-glucoside D-(+) xylose i-erythritol Maltose N-acetyl-D-glucosamine Glyceraldehyde D-galactonic acid δ-lactone Galacturonic acid o-hydroxybenzoic acid p-hydroxybenzoic acid Malonic acid α-keto butyric acid Malic acid L-arginine L-asparagine L-phenylalanine L-serine L-threonine L-glutamic acid β-phenyl ethylamine Putrescine Tween 40 Tween 60 α-cyclodextrin Glycogen α- D-glucose-1-phosphate
100 μl of C substrate:saline medium-TV, and each set of carbon sources was replicated three times in a single 96 well MT plate. Wells containing saline medium-TV without C substrates were also prepared as controls. A tenfold dilution of the first soil suspension (10−2) was used to inoculate the MT plates (bulk and rhizosphere soils). The soil suspension was allowed to sediment for 2 h and 100 μl of the supernatant were inoculated into each well of the MT plate. Substrate utilisation was indicated by colour development of the tetrazolium violet redox dye, and recorded after 3 and 7 days incubation at 25°C. The total number of C sources utilised, and the number of C sources utilised within each group of substrates (amines, amino acids, carbohydrates, carboxylic acids, glucose 1-phosphate, polymers), was determined.
Physico-chemical analyses of bulk and rhizosphere soil samples Soil pH was measured in H2O using a 1:2.5 soil solution ratio. Organic C and chlorine concentration were measured in the 1:2.5 soil/H2O extracts (after 1 h shaking and filtration through 0.2 μm Chromafil® Pet polyester filters). Organic C in water extracts was measured by digestion with 0.4 N K2Cr2O7; the unused dichromate was titrated against Fe(NH4)2 (SO4)2 using a Metrohm 682 Dosimat (Metohm, Switzerland). Chlorine concentration was determined by spectrophotometry using a modified mercury (II) thiocyanate method (Frankenberger et al. 1996). Briefly, sample and standard solutions are mixed with 0.05 M Fe(NO3)3·9H2O and 0.075% Hg(SCN)2 solution. Standard solutions were prepared using NaCl. Absorbance was determined at 460 nm using a 96 well microplate reader (iEMS Reader MF, Labsystems). The concentration of HCH isomers (α-HCH, βHCH, δ-HCH and γ-HCH) soluble in hexane:acetone was measured in 6–8 samples of bulk and rhizosphere soils of Cytisus and Holcus, unplanted soils and sterilised soils. Extraction was carried out in glass test tubes by adding 15 ml of 1:1 (v/v) hexane:acetone to 0.2 g of ground soil sample. The suspension was sonicated for 30 min, filtered through 0.2 μm Chromafil® Pet polyester filters and made up to 25 ml with extractant. This method has previously been shown to efficiently extract HCH isomers (Concha-Graña et al. 2006). The samples were stored at −20°C for posterior chromatographic analysis. Aliquots were diluted appropriately in hexane for HCH determination. Identification and quantification of the different HCH isomers was carried out by GC/ ECD, with a gas chromatographer (Mod. GC 8532 Mega 2 Series, Fisons Instruments, Milan, Italy) equipped with an electron capture detector (Mod. ECD 850, Thermo Quest, Milan, Italy). A split/splitless automatic injector (Mod. AS 800, Fisons Instruments, Milan, Italy) was used in splitless mode, applying an injection volume of 1 μl. The isomers were separated using an Rtx®-ClPesticides capillary column (Restek Corporation, USA, Bellefonte, PA) of 30 m length× 0.25 mm inner diameter with a packed stationary phase. The injection temperature was 270°C and the detector temperature 300°C. Helium (He) was used
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as the carrier gas, at a pressure of 115 kPa, and nitrogen (N2) as make up gas (105 kPa). Oven temperature program applied was: 60°C (3 min), an increase of 30°C min−1 to 180°C (0 min), then an increase of 6°C min−1 to 230°C (0 min) and finally, an increase of 30°C min−1 to 270°C (4 min). The calibration curve data was obtained by injection of standard solutions of the mixture of the four isomers in hexane. The limit of detection for all four isomers was 0.5 μg l−1. Total HCH concentration was calculated as the sum of the four isomers (α-HCH, β-HCH, δ-HCH and γ-HCH).
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Results Plant growth and biomass production Figure 1 shows the mean biomass of Cytisus and Holcus in each treatment (control, INOC and 100HCHINOC). HCH contamination did not significantly
Soil solution extraction and analysis Soil pore water was obtained using in situ Rhizon Flex soil moisture samplers (Rhizosphere Research Products, Wageningen, Netherlands) inserted horizontally in the pots when filling. The samplers consisted of 100-mm porous polymer tubes (2.5 mm o.d., 1.5 mm i.d., pore diameter ∼0.1 μm) connected to a 100-mm PVC tube and a Luer–Lock male connector. Every month, the soils were watered to field capacity, left for 18 h and solution (6–10 ml) was extracted through the samplers by creating a vacuum produced by attaching a 10-ml syringe to the Luer–Lock connection of the sampler. Solutions were analysed for Cl (as described above) and for HCH isomers. HCH present in the soil solutions was extracted with hexane (1:2 (v/v) soil solution: hexane) and quantified by GC/ECD as described above. Statistical analyses The data were processed by a standard analysis of variance. The effects of the factors plant species (Cytisus or Holcus), soil type (rhizosphere or bulk) and treatment (0HCH, INOC, 100HCH-INOC) were studied by 3-way analysis of variance (ANOVA). Tukey’s honestly significant difference (HSD) procedure was applied to separate means. A Student’s t test was used to detect significant differences between rhizosphere and bulk soils. The Kruskall Wallis non parametric ranking test was used to test for differences in C substrate utilisation between species and soil type (bulk or rhizosphere).
Fig. 1 Biomass production (±SE) of Cytisus striatus and Holcus lanatus obtained after 180 days growth in control, INOC and 100HCH-INOC treatments. For each plant tissue different letters indicate significant differences at P
