CC BY
63
YflK 502.521:504.5:622.32
H.-H. Liste
RHIZOSPHERE BACTERIA COMMUNITY AND PETROL HYDROCARBON (PHC) BIODEGRADATION IN SOIL PLANTED TO FIELD CROPS
Julius Kuhn-Institute (JKI), Federal Research Centre for Cultivated Plants, D-14195 Berlin, Germany, Konigin-Luise-Strasse 19; [email protected]
A greenhouse pot experiment was conducted for investigating the capability of a grass (annual ryegrass), a legume (summer vetch), and a crucifer (white mustard) to grow in a soil from a former coal gasification site and promote the biodegradation of petrol hydrocarbons (PHCs). Soil concentrations of 1517 mg kg-1 of total petrol hydrocarbons (TPHs), including 71.4 mg kg-1 of total US EPA priority polycyclic aromatic hydrocarbons (TPAHs) have caused a significant (P<0.05) reduction in shoot and root yields by more than 50%. Abundance and bacterial community composition in soil and rhizosphere were distinctly altered by the PHC contaminants and also depended on crop species and age. After 95 days,
68.7% of initial TPH amounts and 59% of the TPAHs had disappeared from unplanted soil.
The removal of PHCs was fostered in soil planted to mustard and vetch reaching final TPH concentrations that were 15.6% and 12%, respectively, lower than in unplanted soil. Mustard and vetch elicited the greatest degradative root activities and sustained particularly great populations of rhizosphere bacteria that are known hydrocarbon degraders. None of the crops aided the biodegradation of TPAHs in soil.
Keywords: bacteria; bioavailability; biodegradation; crops; PAHs; petrol hydrocarbons; phytoremediation; rhizosphere.
Introduction
Bioremediation often is used to clean up hazardous waste sites that are contaminated with petrol hydrocarbons, but current in situ techniques are often ineffective for the removal of the more recalcitrant, toxic, mutagenic, and carcinogenic PHC constituents such as PAHs [57; 5]. Accelerating the biodegradation of petrol hydrocarbons in general and of PAHs in particular is thus a major challenge to improving the performance and acceptance of cost-saving soil bioremediation techniques.
Plants are suggested to enhance the degradation of organopollutants because of greater numbers and activities of microbes in the rhizosphere [10; 42]. A plant-aided biodegradation was evident for pesticides [57], petrol hydrocarbons [56], PAHs [4; 29] and many other organic compounds [32]. Yet, there is also a number of research studies demonstrating neutral [14] or inhibitory [51] plant effects with many data indicating that plant species effect the fate of particular chemicals differently [7]. A more systematic approach to selecting plant species for phytoremediation purposes is thus essential but has rarely been attempted [11; 56]. Especially differences in microbial community features and contaminant degradation between rhizosphere soils of various plant species or types are not well documented in the literature [7].
The objective of this study was to investigate three field crops from different botanical groups for their rhizosphere bacterial communities and capabilities to grow in a contaminated soil from a former coal gasification site and foster the biodegradation of petrol hydrocarbons, including PAHs. Field crops are of particular interest for phytoremediation purposes because they are bred for fast growth and high yields.
Materials and methods
Soils and treatments
Loamy sand (2.1% organic carbon, pH 7.6) from Berlin and soil contaminated with petrol hydrocarbons (46.3 g TPHs kg-1 dry soil), including PAHs (2129.8 mg TPAHs kg-1 dry soil) from a former coal gasification plant site in Groditz, Saxony, were air dried and passed through a 6-mm sieve to remove large rocks and debris. The loamy sand (100 kg) and fertilizer (0.42 kg) (6% NO3-N, 7% NH4-N, 13% P2O5, 21% K2O) were placed in a cement mixer and blended for 10 min. The contaminated soil (3350 g) was added to the cement mixer and diluted thoroughly with the fertilized loamy sand for 20 min. The procedure was aimed to have contaminated and control soil of similar physical properties and to decrease the toxicity of polluted soil. The contaminated loamy sand mixture (2.4% organic carbon, pH 7.7) was stored for equilibration (aging) for 30 days in the dark at 15°C and final concentrations were reached of 1517 mg TPHs kg-1 dry soil, including 71.4 mg TPAHs kg-1 dry soil.
©ListeH.-H., 2011
Portions of 1000 g of the contaminated or uncontaminated loamy sand were placed in black plastic pots (13 cm diam.), and seeds (Saaten-Union GmbH, Hannover, Germany) of summer vetch (Vicia sativa L.), white mustard (Sinapsis alba L.), and annual ryegrass (Lolium multiflorum Lam.) were sown into the soil (15 seeds per pot). Seedlings were later thinned to give five plants per pot. Tap water was added to the soils to bring the moisture level to approximately 80% of the water-holding capacity. The plants were grown in a greenhouse maintained at 15°C during a 16-h day and at 10°C during an 8-h night. The light intensity was according to sunlight conditions of early summer and additional light was provided as needed by sodium vapor lamps (Osram Violox T 400) at an intensity of 10,000 cdm-2. The soils were watered daily and fertilized every two weeks with 10 ml of a 0.3-% solution of Poly Crescal (AGLUKON GmbH, Dusseldorf, Germany) per 100 g of soil. The 12 replicate pots of each treatment were completely randomized in the greenhouse.
Plant analysis
After 95 days of plant growth, from six pots of each treatment the shoots were excised from the roots. All soil was removed from the pots and air dried before the roots that had completely permeated the entire soil could be separated from the planted soil by passing the soil through a 6-mm sieve and picking the remaining roots by hand. The roots and shoots were washed in tap water, dried for 24 h at 105°C, and weighed.
Soil analysis
For extraction of TPHs, air-dried soil (10 g) from each sampled pot was homogenized with 30 g of anhydrous Na2SO4 and subjected to extraction using 1h sonication with 110 ml of 1,1,2-trichlorotrifluoroethane (IR spectroscopy grade, Merck AG, Darmstadt, Germany). The hydrocarbon extracts were cleaned on neutral aluminum oxide powder (50-200 ^m, pH 7.0-7.8) and analyzed by an infrared analyzer (Horiba OMCA-200) as TPH-IR according to DIN method 38409-18 (DIN, 1981).
For PAH extraction, air-dried soil (10 g) from each sampled pot was mixed with 20 g of anhydrous Na2SO4 and subjected to extraction using 2h sonication with 100 ml of tetrahydrofuran (Merck AG, Darmstadt, Germany). The PAH extracts were analyzed with HPLC using UV detection based on U.S. EPA Method 610 (US EPA, 1998). Extraction and analysis of TPHs and PAHs were conducted by an accredited commercial service laboratory (Chemisches Labor Dr. Betz GmbH, Berlin, Germany).
Bacterial enumeration and identification
After 15 days and when crops began to bloom (mustard, 57 d; vetch, 61 d) or after 60 days (ryegrass and soil), plants were carefully pulled from six pots of each treatment and mildly shaken to remove soil loosely adhering to the roots. The roots with the remaining adherent soil were separated from the shoots and portions of 1 g (15-day samples) or 10 g (wet wt.) were transferred either to 50-ml glass tubes filled with 9 ml or into 250-ml Erlmeyer flasks containing 90 ml of sterile saline (0.9%). Tubes and flasks were closed with an autoclaved cotton plug, and shaken for 20 min at 200 rev min-1 on an orbital shaker (B. Braun Biotech International GmbH, Melsungen, Germany). After allowing the soil slurry to settle for 20 min, 1 ml of the supernatant was serially diluted in 50-ml glass tubes containing 9 ml of sterile saline (0.9%), and aliquots (0.1 ml) of the proper dilution (producing 30 to 300 colonies per plate) were spread on glycerin-peptone agar plates [17] in duplicate. The unplanted soil was processed the same way. The plates were incubated for 5 to 7 days at 25°C prior conducting discriminatory colony counting for determining abundance and composition of the bacterial community. In more detail, all colonies of a plate were grouped and counted according to their morphological appearance [44; 52] and one representative of each colony morphotype (cmt) was isolated and purified. The isolates were taxonomically identified by gas-chromatographic analysis of their whole-cell fatty acid methyl esters (FAME) with the Hewlett-Packard HP5898A Microbial Identification System (MIS®) using versions 3.80 and 4.01 of the Aerobic Library (TSBA 40) (MIDI, Inc., Newark, DE, USA) and according to the procedure as specified by the manufacturer (Hewlett-Packard, Avondale, Pennsylvania, USA). Microbes that vanished during cultivation were summarized as "Deserter" and those that failed FAME identification have been designated by the system as "no match". Organisms not identifiable by FAME analysis were coarsely grouped on the basis of biochemical or colony morphology features. Bacteria were discriminated by Gram reaction using KOH (3%) [45]. Actinomycetes colonies (firm consistency, powdery or chalky appearance) were categorized according to the release of brownish pigments into the agar as either actinomycetes(+) (pigment positive) or actinomycetes1'"'1 (pigment negative). Very small, undifferentiable colonies were assigned to as "Pinpoints" [53].
Data analysis
Means based on six, eight, or twelve replicates for each treatment were compared by analysis of variance (ANOVA) followed by a Student-t test (P<0.05) or a Student-Newman-Keuls Test (P<0.05 or 0.1) using COSTAT 5.0 (CoHort Software, Minneapolis, MN, USA).
Results
Plant yields
Petrol hydrocarbons in soil (1517 mg TPHs kg-1) have affected plant dry matter yields during 95 days (Fig. 1). Shoot yields had decreased significantly (P<0.05) for ryegrass by 38.9% and for vetch by 49.8% and root yields were reduced (P<0.05) for ryegrass by 52.6%. Still, ryegrass had the most massive root system of all three crops in contaminated soil.
Fig. 1. Shoot and root dry matter yields in petrol hydrocarbon-contaminated soil (1517 mg TPHs kg-1) after 95 days of crop growth in the greenhouse.
The error bars indicate the least significant difference (LSD) (P<0,05; n=6)
Fig. 2. Relative frequency distribution of MIS® similarity indices for all FAME-identified soil and rhizosphere bacteria (920 isolates)
Soil and rhizosphere bacterial community
Abundance and composition of the culturable soil and rhizosphere bacteria community were examined after two and eight weeks into the experiment. At 15 days, contaminated soil contained as much as 18-fold more bacteria than unpolluted soil (Table 1). Juvenile mustard and vetch raised the number of total bacteria in the root zone in unpolluted soil (R/S ratios of 1.8 and 8.2) whereas in contaminated soil, bacteria in the rhizospheres of all three crops were less abundant as compared to unplanted soil (R/S ratios between 0.3 and 0.5) (Table 1). After eight weeks, total bacteria counts in contaminated and pristine soil were alike and most
numerous in the rhizospheres (Table 1). Bacteria were particularly abundant in the root zones of flowering mustard and vetch reaching 11 to 30-fold the numbers of unplanted soil.
Table 1
Number of culturable bacteria in pristine and petrolhydrocarbon (PHC)-contaminated fallow
and rhizosphere soil
Contaminants Plant Total counts (106 cfu g-1) R/S ratio3 Colony types (cmtb plate)
2 weeksc 8 weeksd 2 weeks 8 weeks 2 weeks 8 weeks
None None 32 Bae 18 Ba NA1 NA 9 Aa 7 Bb
Mustard 58 Bb 545 Aa 1,8 Bb 30,3 Aa 9 Aa 11 Aa
Vetch 261 Aa 529 Aa 8,2 Ab 29,4 Aa 7 Aa 9 Ba
Ryegrass 12 Bb 214 Ba 0,4 Bb 11,9 Ba 6 Ab 8 Ba
PHCs2 None 591 Aa 45 Cb NA NA 9 ABa 10 Ba
Mustard 275 Bb 488 Aa 0,5 Ab 10,8 Aa 11 Aa 13 Aa
Vetch 211 Cb 436 Aa 0,4 Bb 9,7 Aa 7 BCb 10 Ba
Ryegrass 156 Cb 249 Ba 0,3 Bb 5,5 Ba 7 Cb 10 Ba
a R/S ratio - rhizosphere to soil ratio of total bacteria counts. b cmt - colony morphotypes as obtained from original soil dilution agar plates. c 2 weeks - sampling after 15 days of plant growth.
d 8 weeks - sampling after 57 days (mustard), 61 days (vetch), or 60 days (ryegrass, unplanted soil) of plant growth. e Values in a column for either contamination level that are followed by different capital letters and values in a row for either parameter that are followed by different lowercase letters are statistically different (P<0,05, n=8).
1 NA - not applicable.
2 PHCs - petrolhydrocarbons.
A total of 1413 representative colony morphotypes were isolated from the original soil dilution plates and 65.2% of the isolates could be taxonomically classified by FAME analysis. Of all FAME-identified isolates, 66.3% matched a known organism in the MIS® database at similarity indices > 0.5 and 82.6% had matches with a similarity index > 0.3 (Fig. 2).
The bacterial community diversity was similar in contaminated vs. pristine soil but the species composition has differed (Fig. 3). At fifteen days, a total of 66 classified bacteria (including actinomycetes(+)/(-)) were isolated from polluted compared to 69 from pristine soil. Most notably, Bacillus spp. were much less common in PHC-polluted soil (4 species) than in uncontaminated soil (16 species) (Fig.
3). Actinomycetes(+)/(-) colonies were not isolated at all from polluted soil, which, however, harbored various plant pathogens (Acidovorax avenae, Clavibacter michiganense, Ps. marginalis, Ps. syringae, Ps. viridiflava, Rathayibacter tritici) and Risk Group 2 human pathogens (BGCh, 2002) (Acinetobacter junii, Actinobacillus lignieresii, Alcaligenes xylosoxydans, Bordetella bronchiseptica, Bordetella parapertussis, Enterococcus faecium, Kluyvera ascorbata, Moraxella catarrhalis, Ochrobactrum anthropi, Yersinia pseudotuberculosis) not found in pristine soil. Particularly abundant in PHC soil were known petrol hydrocarbon degraders such as Alcaligenes piechaudii, Ps. putida, and Stenotrophomonas maltophilia (Table 2). After eight weeks, 75 bacteria species were identified in polluted compared to 80 in pristine soil (Fig. 3). Interestingly, the bacilli diversity has become similar in polluted and pristine soil and actinomycetes(+)/(-) have also appeared in PHC soil. Potential petrol hydrocarbon degraders such as Flavobacterium johnsoniae, Ps. fluorescens, Ps. putida, Ps. stutzeri, Ps. syringae, and Vibrio furnissii have still occurred only or in particular great numbers in contaminated soil (Fig. 3, Table 2).
Table 2
Abundance (106 cfu g-1) of common bacteria populations in petrolhydrocarbon (PHC)-contaminated
fallow and rhizosphere soil
Plant
Bacteria 2 weeka 8 weekb
None Mustard Vetch Ryegrass None Mustard Vetch Ryegrass
Acidovorax avenae 1ac 0a 0a 0a 0a 6a 11a 0a
Alcaligenes piechaudii 0a 6a 26a 2a 0a 0a 5a 0a
Bordetella parapertussis 18a 0a 0a 0a 0 0 0 0
Burkholderia cepacia 0 0 0 0 0a 5a 0a 0a
Enterobacter intermedius 0 0 0 0 0a 0a 13a 0a
Flavobacterium j ohnsoniae 1b 7ab 13a 0b 0b 8b 34a 5b
IVlltl U Ucltiei 1U1U esteraromaticum 0a 13a 0a 0a 1a 4a 0a 4a
Nesterenkonia halobia 0a 0a 0a 1a 0a 1a 6a 3a
Pseudoalteromonas haloplanktis 0 0 0 0 0a 3a 1a 0a
Pseudomonas balearica 0 0 0 0 0a 3a 1a 0a
Pseudomonas chlororaphis 2b 12b 0b 1b 0 0 00
Pseudomonas fluorescens 3ab 8a 4ab 0b 2a 11a 9a 0a
Pseudomonas putida 13b 46a 30ab 36ab 0b 31a 31a 1b
Pseudomonas stutzeri 0 0 0 0 0b 0b 11a 0b
Pseudomonas syringae 0a 2a 7a 0a 0a 4a 6a 0a
Rahnella aquatilis 0b 4b 13a 0b 0 0 00
Serratia liquefaciens 13a 0a 0a 0a 0 0 00
Serratia plymuthica 0b 1b 19a 0b 0 0 00
Sphingobacterium spiritivorum 9a 2a 1a 0a 0a 3a 13a 3a
Stenotrophomonas maltophilia 28ab 44a 18ab 3b 0a 3a 0a 0a
Vibrio fur nissii 0 0 0 0 0a 4a 0a 0a
Yersinia pseudotuberculosis 6a 2a 2a 0a 0b 4b 30a 1b
Actinomycetes(+) 0 0 0 0 0b 3a 1b 0b
Actinomycetes(-) 0 0 0 0 0b 3a 0b 0b
a 2 weeks - sampling after 15 days of plant growth.
b 8 weeks - sampling after 57 days (mustard), 61 days (vetch), or 60 days (ryegrass, unplanted soil) of plant growth. c Means in rows for either sampling date that are followed by different lowercase letters are statistically different (P<0,10). Means were obtained from 12 replicates (2 weeks) and from 8 replicates (8 weeks). Means in rows for either sampling date that are not followed by letters were not comparable by ANOVA (no replicate values >0).
Occurrence and frequency of bacteria species in soil also depended on the crops. Most colony morphotypes were detected on agar plates from mustard rhizosphere samples (9 to 13 cmt plate-1) and mainly from polluted soil after eight weeks (Table 1). Important hydrocarbon-degraders, including Alcaligenes piechaudii, Flavobacterium johnsoniae, Pseudomonas spp., and Vibrio furnissii were particularly promoted in the contaminated root zones of mustard and vetch (Table 2). Interestingly, the appearance of actinomycetes(+)/(-) in PHC soil after eight weeks was confined to the rhizospheres of mustard and vetch. Petrol hydrocarbon degradation
Initial concentrations of TPHs (1517 mg kg-1) and TPAHs (71.4 mg kg-1) have declined significantly (P<0.05) in all treatments during 95 days (Table 3). In unplanted soil, 68.7% of TPHs and 59% of the TPAHs had disappeared. Mustard and vetch plants have further fostered the biodegradation of petrol hydrocarbons reaching final TPH concentrations that were 15.6% and 12%, respectively, lower than in unplanted soil. Ryegrass had no beneficial effect on the removal of TPHs and none of the crops had aided the biodegradation of TPAHs in soil.
When the extents of additional biodegradation in planted soil relative to root mass are compared, it is evident that the greatest degradative activities were associated with the small root systems of vetch and mustard (Table 3). Ryegrass did not show any degradative activity for TPHs. None of the crops exerted a root activity that aided extra biodegradation of TPAHs.
Fig. 3. Bacterial community composition in pristine (P) and petrol hydrocarbon-contaminated (C) soil (unplanted and rhizosphere) after 2 weeks (-A-) and 8 weeks (-B-) of crop growth.
Grey fields indicate the occurrence of the particular microorganisms
Bacterias: (1) Acidovorax avenae, (2) Acidovorax delafieldii, (3) Acinetobacter baumannii, (4) Acinetobacter haemolyticus, (5) Acinetobacter junii, (6) Acinetobacter lwoffii, (7) Actinobacillus lignieresii, (8) Aeromonas sobria, (9) Agrobacterium radiobacter, (10) Agrobacterium rubi, (11) Agrobacterium tumefaciens, (12) Alcaligenes faecalis, (13) Alcaligenes piechaudii, (14) Alcaligenes xylosoxydans, (15) Amycolatopsis orientalis, (16) Arthrobacter agilis, (17) Arthrobacter atrocyaneus, (18) Arthrobacter aurescens, (19) Arthrobacter citreus, (20) Arthrobacter crystallopoietes, (21) Arthrobacter globiformis, (22) Arthrobacter ilicis, (23) Arthrobacter mysorens, (24) Arthrobacter nicotianae, (25) Arthrobacter oxydans, (26) Arthrobacter pascens, (27) Arthrobacter protophormiae/ramosus, (28) Azospirillum brasilense, (29) Bacillus cereus, (30) Bacillus circulans, (31) Bacillus coagulans, (32) Bacillus dipsosauri, (33) Bacillus flexus, (34) Bacillus globisporus, (35) Bacillus laterosporus, (36) Bacillus lentimorbus, (37) Bacillus licheniformis, (38) Bacillus marinus, (39) Bacillus megaterium, (40) Bacillus mycoides, (41) Bacillus psychrophilus, (42) Bacilluspumilus, (43) Bacillus simplex, (44) Bacillus sphaericus, (45) Bacillus subtilis, (46) Bacillus thuringiensis canadensis sv., (47) Bacillus thuringiensis kurstakii, (48) Bordetella avium, (49) Bordetella bronchiseptica, (50) Bordetella parapertussis, (51) Brevibacillus agri, (52) Brevibacillus centrosporus, (53) Brevibacterium helvolum, (54) Brevibacterium mcbrellneri, (55) Brevundimonas diminuta, (56) Brevundimonas vesicularis, (57) Burkholderia cepacia, (58) Cellulomonas biazotea, (59) Cellulomonas fimi, (60) Cellulomonas turbata, (61) Chromobacterium violaceum, (62) Chryseobacterium meningosepticum, (63) Clavibacter michiganense, (64) Comamonas acidovorans, (65) Corynebacterium ammoniagenes, (66) Corynebacterium aquaticum, (67) Curtobacterium citreum, (68) Curtobacterium flaccumfaciens, (69) Dactylosporangium fulvum, (70) Deinococcus erythromyxa, (71) Deserter, (72) Enterobacter intermedius, (73) Enterococcus faecalis, (74) Enterococcus faecium, (75) Flavobacterium aquatile, (76) Flavobacterium johnsonae, (77) Flavobacterium resinovorum, (78) Gluconobacter asaii, (79) Gordona bronchialis, (80) Hafnia alvei, (81) Hydrogenophaga pseudoflava, (82) Kluyvera ascorbata, (83) Kocuria kristinae, (84) Kocuria rosea, (85) Kocuria varians, (86) Kytococcus sedentarius, (87) Leuconostoc mesenteroides dextranicum, (88) Methylobacterium extorquens, (89) Methylobacterium mesophilicum, (90) Methylobacterium organophilum, (91) Microbacterium barkeri, (92) Microbacterium esteraromaticum, (93) Microbacterium lacticum, (94) Microbacterium liquefaciens, (95) Microbacterium saperdae, (96) Micrococcus luteus, (97) Micrococcus lylae, (98) Micromonospora carbonacea, (99) Moraxella catarrhalis, (100)Myroides odoratus, (101) Nesterenkonia halobia, (102) No match, (103) Nocardia brasiliensis , (104) Nocardia carnea, (105) Nocardia globerula, (106) Nocardioides albus, (107) Ochrobactrum anthropi, (108) Paenibacillus apiarius, (109) Paenibacillus azotofixans, (110) Paenibacillus gordonae, (111) Paenibacillus macerans, (112) Paenibacilluspabuli, (113) Paenibacilluspolymyxa, (114) Pantoea agglomerans, (115) Paracoccus denitrificans, (116) Pedobacter heparinus, (117) Phyllobacterium rubiacearum, (118) Pseudoalteromonas haloplanktis, (119) Pseudomonas balearica, (120) Pseudomonas chlororaphis, (121) Pseudomonas corrugata, (122) Pseudomonas flectens, (123) Pseudomonas fluorescens, (124) Pseudomonas marginalis, (125) Pseudomonas mendocina, (126) Pseudomonas putida, (127) Pseudomonas saccharophila, (128) Pseudomonas savastanoi pv. fraxinus, (129) Pseudomonas stutzeri, (130) Pseudomonas syringae, (131) Pseudomonas viridiflava, (132) Rahnella aquatilis, (133) Rathayibacter tritici, (134) Rhodobacter capsulatus, (135) Rhodococcus erythropolis, (136) Rhodococcus fascians, (137) Rhodococcus globerulus, (138) Rhodococcus rhodochrous, (139) Roseomonas group** Ochrobactrum, (140) Serratia liquefaciens, (141) Serratia plymuthica, (142) Sphingobacterium multivorum, (143) Sphingobacterium spiritivorum, (144) Sphingomonas macrogoltabidus, (145) Staphylococcus hominis, (146)
Staphylococcus lugdunensis, (147) Stenotrophomonas maltophilia, (148) Streptococcus sanguis, (149)
Streptoverticillium reticulum, (150) Tsukamurella paurometabolum, (151) Tsukamurella wratislaviensis, (152) Variovorax paradoxus, (153) Vibrio furnissii, (154) Xanthobacter agilis, (155) Xanthomonas axonopodis, (156) Xenorhabdus nematophilus, (157) Yersinia enterocolitica, (158) Yersinia pseudotuberculosis, (159).
Petrol hydrocarbon degradation
Initial concentrations of TPHs (1517 mg kg-1) and TPAHs (71.4 mg kg-1) have declined significantly (P<0.05) in all treatments during 95 days (Table 3). In unplanted soil, 68.7% of TPHs and 59% of the TPAHs had disappeared. Mustard and vetch plants have further fostered the biodegradation of petrol hydrocarbons reaching final TPH concentrations that were 15.6% and 12%, respectively, lower than in unplanted soil. Ryegrass had no beneficial effect on the removal of TPHs and none of the crops had aided the biodegradation of TPAHs in soil.
When the extents of additional biodegradation in planted soil relative to root mass are compared, it is evident that the greatest degradative activities were associated with the small root systems of vetch and mustard (Table 3). Ryegrass did not show any degradative activity for TPHs. None of the crops exerted a root activity that aided extra biodegradation of TPAHs.
Table 3
Concentration of TPHs and TPAHs in unplanted petrolhydrocarbon (PHC)-contaminated soil
and PHC soil planted to field crops
Plant Concentration in soil (mg kg-1) Root wt. (g pot-1) Degradative activity (mg g-1 root)
TPHsa TPAHsb TPHs TPAHs
0 days 95 days 0 days 95 days 95 days 95 days 95 days
None 1517a 475ABbc 71,4a 29,3Ab NAd NA NA
Mustard 1517a 401Bb 71,4a 29,4Ab 0,85 B +100 Be -0,1 Af
Vetch 1517a 418Bb 71,4a 30,5Ab 0,26 B +259 A -5,9 A
Ryegrass 1517a 541Ab 71,4a 30,7Ab 3,67 A -19 C -0,4 A
a TPHs - total petrolhydrocarbons.
b TPAHs - total of 16 US EPA priority polycyclic aromatic hydrocarbons.
c Values in a column that are followed by different capital letters and values in a row for either contaminant fraction that are followed by different lowercase letters are statistically different (P<0,05, n=6). d NA - not applicable. e (+) - plant-promoted degradation. f (-) - plant-inhibited degradation.
Discussion
Ryegrass, vetch, and mustard were compared in the greenhouse for their capabilities to enhance the biodegradation of PHCs in soil from a former gas plant site. All three field crops have grown in the soil that contained 1517 mg TPHs, including 71.4 mg TPAHs per kg of dry soil. Still, plant vigor (photosynthetic activity) was depressed by PHCs (H.-H. Liste, 2003, Habilitation thesis) and shoot and root dry matter yields were reduced by more than 50% for ryegrass and vetch.
Toxicity of light petrol constituents (e.g. octane, BTEX) to plants and also unfavorably changed chemical, physical, and biological soil conditions can affect plant growth [36]. Poor plant growth in hydrocarbon-contaminated soils is often the result from severe water repellency that affects the water regime in soil [27; 37]. Phytosanitary aspects also may have contributed to the reduced crop growth in PHC-contaminated soil. Several Bacillus spp. and actinomycetes had vanished from PHC soil and these microbes were linked to plant disease suppression [49]. Correspondingly, plant pathogens, e.g. Clavibacter michiganense and Pseudomonas syringae, were more abundant in the polluted than in the pristine rhizosphere of young plants.
Despite plant growth depression, PHCs were biodegraded more rapidly in planted soil. Within 95 days, 73.6% or 72.4% of the initial 1517 mg TPH kg-1 have disappeared from soil planted to mustard or vetch as compared with 68.7% from unplanted soil. This is consistent with results from numerous laboratory, greenhouse, and field trials, which provide persuasive evidence that the roots of plants can aid soil bioremediation [14; 29; 30; 40].
The ability to accelerate petrol hydrocarbon degradation in soil has differed among the crops. The PHC removal during 95 day was greatest for mustard while vetch had the highest degradative activity. No remedial effect was seen for ryegrass despite its large root mass. It has repeatedly been shown that the potential to ameliorate contaminated soils differs among species and even varieties and no correlation exists between root mass and a plants capacity to foster pollutant degradation [28; 34; 56].
Plant physiological traits, namely rhizodeposition seem very important for pollutant degradation. Rhizodeposits can supply nitrogen or oxygen to microorganisms in contaminated soils that are often deficient of these elements [42], stimulate phosphate-solubilizing microbes and hence phosphorous availability [12], may enhance contaminant bioavailability [54], induce pollutant degradation pathways [24], or fortuitously select microbes that metabolize or cometabolize particular organic compounds [15]. Interestingly, mustard and vetch in particular have promoted the proliferation of bacteria, including Alcaligenes piechaudii, Flavobacterium johnsoniae, Ps. fluorescens, Ps. putida, Ps. stutzeri, Ps. syringae, Stenotrophomonas maltophilia, and Vibrio furnissii that are common and efficient degraders of aliphatic and aromatic hydrocarbons in soil [8; 46; 50; 55]. Besides, vetch may have boosted the degradation of PHCs by symbiotic fixation of atmospheric nitrogen even though the number of root nodules was lower in contaminated soil. Legumes can further enhance the clean-up of soils defiled with petrol hydrocarbons because these soils are often nitrogen deficient [23; 42-43].
After all, the promotion by plants of PHC biodegradation in soil was only moderate. Interestingly, in this regard, the number of bacteria in the rhizosphere of juvenile crops was not enhanced, which is important to note, because greater numbers and activities of microbes in the rhizosphere are considered fundamental to a plant-fostered biodegradation of organic contaminants in soil [3; 40]. This absence of an early rhizosphere effect in PHC soil was presumably the result of surplus carbon from PHCs that markedly promoted bacteria proliferation in polluted vs. pristine soil and left root-exuded carbon without the typical stimulation effect on microbial counts. Petrol hydrocarbons contain a variety of easy degradable and available compounds, including BTEX and alkanes that have boosted microbial growth and metabolism while being degraded rapidly in soil even without plants [18; 31]. Weeks later, bacteria in PHC soil were much more abundant in the rhizosphere. Easy available PHC fractions had likely ceased in polluted soil and, therefore, microorganisms became carbon limited as common in soils and only root deposits remained a major source of surplus carbon for increased microbial growth in the root zone. This delayed rhizosphere effect has eventually accelerated the degradation of residual PHCs, but 95 days were obviously too short for more decisive plant effects.
Presumably, easy available portions of organic soil contaminants must be consumed first by soil microorganisms before the remedial benefits of the rhizosphere come into full effect. As such, a more rapid depletion of readily available PHC fractions in polluted soil planted to mustard and vetch as indicated by the early appearance of actinomycetes in their root zones after eight weeks may have contributed to the more pronounced beneficial effects of both crop species on overall TPH removal. Actinomycetes only become prominent in soil when easy available organics have diminished [1] and these microorganisms play a key role in the mineralization of the more resistant and less available portions of petrol hydrocarbons such as PAHs [22].
Moreover, the stage of plant development is important for phytoremediation effects. Bacterial growth and metabolism and hence the biodegradation of organics in the rhizosphere increase gradually and often peak after a few months of plant growth [25]. The rhizosphere effect on bacterial numbers was greater for blooming than for juvenile crops in this study. Mature plants enhance the extra break down of organic matter in soil more strongly than younger plants [25], partly because young plants also need time to develop a dense root system that is prerequisite for exposing a maximum of contaminated soil to remedial rhizosphere effects. Plant-aided bioremediation of soils just takes a considerable period of time [10].
No plant-accelerated biodegradation at all was observed for the more persistent petrol hydrocarbon constituents, the PAHs. Fifty nine percent of initial TPAH amounts disappeared from bare soil within 95 days, which suggests good bioavailability. Part of the residual 41% of US EPA-PAHs was likely to be less available to microbial attack since the contaminants were in soil for many years and increasing portions of PAHs are becoming sequestered with time in soil [2; 16]. Because plants are able to enhance mobility and availability of weathered PAHs and other POPs in soil [20; 30; 33; 54] rhizosphere effects on PAH biodegradation may only become evident when bioavailable portions have disappeared and sequestered PAH residues prevail. Also and according to the diauxie phenomenon [39], microbial PAH degradation may not be accelerated, not even by plants, until the more easily degradable PHC constituents are exhausted. Polycyclic aromatic compounds were most resistant to removal from oily soils by biological treatment [19].
Of all crops, only ryegrass has hindered PHC degradation in soil. This compares with a negative rhizosphere priming effect, which is the phenomenon of a depressed decomposition of soil organic matter in the root zone of certain plant species or at certain stages of plant development [25]. This has been attributed to a decreased microbial growth and metabolism in affected rhizospheres, e.g. due to root uptake of large amounts of nitrogen and hence N deficiency in the root zone [38]. Ryegrass takes up from soil much more nitrogen than most other crops [35]. As a possible consequence, available N was very rapidly depleted in the densely rooted grass pots causing a nitrogen limitation to microbial growth. The fewer bacteria in the rhizosphere of ryegrass as compared with mustard and vetch are indicative of this. An adequate fertilization
is thus critical for positive rhizosphere priming effects and an increased break down of hydrocarbons during phytoremediation and not only with respect to nitrogen because all nutrients other than carbon are strongly limited in the root zone [21; 25; 47].
However, negative rhizosphere priming effects can turn into positive ones as plant development advances [9]. For ryegrass, a negative rhizosphere priming effect has slowly become positive after 2.5 months of growth [26]. Consequently, observation periods longer than the 95 days in this study are required to examine for plant effects on the fate of organic pollutants. Yet, the biodegradation of organics in the rhizosphere can perhaps be accelerated by preventing limitations for plant and microbial growth such as by providing sufficient amounts of nitrogen via fertilization or through cropping legumes as pure or mixed stands.
Conclusions
Crops have the potential to foster the biodegradation of petrol hydrocarbons in soil but phytoremediation effects largely correlate with plant species and age and the availability of contaminants and nutrients. As long as petrol hydrocarbons are available in quantities that substantially promote microbial growth in soil, crops are not likely to aid soil bioremediation unless they overcome nitrogen or other nutrient limitations for extra biodegradation. Only as the availability to microbes of PHCs in soil declines with time, beneficial rhizosphere effects on microbial growth will increase and so may the extra break down of PHCs in contaminated soil planted to crops. Further research, particularly on the interrelations between contaminant and nutrient availability and the biodegradation of organic pollutants in plant rhizospheres is crucial for making considerable progress in phytoremediation applications.
Acknowledgements
The work was made possible by funding through the stipend program of the Deutsche Bundesstiftung Umwelt, Osnabruck. I thank Dr. Sigfrid Kohn and Dr. Kristin Peters for helping with the experiments.
References
1) Alexander M. Introduction to Soil Microbiology. 2nd Edition, John Wiley, New York, NY, 1977. 467 pp.
2) Alexander M. How toxic are toxic chemicals in soil? // Environ. Sci. Technol. 1995. №29. pp. 2713-2717.
3) Anderson T.A., Guthrie E.A., Walton B.T. Bioremediation in the rhizosphere // Environ. Sci. Technol. 1993. №27. pp. 2630-2636.
4) Aprill W., Sims R.C. Evaluation of the use of prairie grasses for stimulating polycyclic aromatic hydrocarbon treatment in soil // Chemosphere.1990. №20, pp. 253-265.
5) Banks M.K., Pekarek S., Rathbone K., Schwab P. Phytoremediation of petroleum contaminated soils: Field assessment. In: Leeson A., Alleman B.C. (Eds.), In Situ and On-Site Bioremediation (Vol. 3). Battelle Press, Columbus, OH, 1997. pp. 305-30S.
6) BGCh. Sichere Biotechnologie - Einstufung biologischer Arbeitsstoffe: Bakterien. Merkblatt B 006 (Berufsgenossenschaft der chemischen Industrie), Jedermann-Verlag, Heidelberg, 2002. 306 pp.
7) Boyle J.J., Shann J.R. Biodegradation of phenol, 2,4-DCP, 2,4-D, and 2,4,5-T in field-collected rhizosphere and nonrhizosphere soils // J. Environ. 1995. Qual. 24. pp. 7S2-7S5.
S) BuhlerM., Schindler J. Aliphatic hydrocarbons. In: Rehm H.-J., Reed G. (Eds.), Biotechnology: A Comprehensive Treatise in S Volumes. Biotransformations (Vol. 6a), Verlag Chemie, Weinheim, 19S4. pp.329-3S6.
9) Cheng W. Measurement of rhizosphere respiration and organic matter decomposition using natural 13C // Plant Soil. 1996. №183. pp. 263-26S.
10) Cunningham S.D, Shann J.R., Crowley D.E., Anderson T.A. Phytoremediation of contaminated water and soil. In: Kruger E.L., Anderson T.A., Coats J.R. (Eds.), Phytoremediation of Soil and Water Contaminants. ACS Symposium Series 664, American Chemical Society, Washington, DC, 1997. pp. 2-17.
11) Cunningham S.D., Anderson T.A., Schwab A.P., Hsu F.C. Phytoremediation of soils contaminated with organic pollutants // Adv. Agron. 1996. №56. pp. 55-114.
12) Das A.C., Debnath A., Mukherjee D. Effect of the herbicides oxadiazon and oxyfluorfen on phosphates solubilizing microorganisms and their persistence in rice fields // Chemosphere. 2003. №53. pp. 217-221.
13) DIN (Deutsches Institut fur Normung e.V.) Methode Nr. 3S409-1S, Bestimmung von Kohlenwasserstoffen in Wasser mit Hilfe eines IR-Olmessgerates. In: Deutsche Einheitsverfahren zur
Wasser-, Abwasser- und Schlammuntersuchung; Summarische Wirkungs- und StoffkenngroBen, Gruppe H., BeuthVerlag, Berlin. 1981.
14) Gunther T., Dornberger U., Fritsche W. Effects of ryegrass on biodegradation of hydrocarbons in soil // Chemosphere. 1996. №33. pp. 203-215.
15) Haby P.A., Crowley D.E. Biodegradation of 3-chlorobenzoate as affected by rhizodeposition and selected carbon substrates // J. Environ. Qual. 1996. №25. pp. 304-310.
16) Hatzinger P.B., Alexander M. Effect of aging of chemicals in soil on their biodegradability and extractability//Environ. Sci. Technol. 1995. №29. pp. 537-545.
17) Hirte W.F. Glycerin-Pepton-Agar, ein vorteilhafter Nahrboden fur boden-bakteriologische Arbeiten. Zbl. Bakt. II Abt. 114, 1961. pp. 141-146.
18) Hollender J., Althoff K., Mundt M., Dott W. Assessing the microbial activity of soil samples, its nutrient limitation and toxic effects of contaminants using a simple respiration test // Chemosphere. 2003. №53. pp. 269-275.
19) Huesemann M.H. Predictive model for estimating the extent of petroleum hydrocarbon biodegradation in contaminated soils // Environ. Sci. Technol. 1995. №29. pp. 7-18.
20) Hulster A., Muller J.F., Marschner H. Soil-plant transfer of polychlorinated dibenzo-p-dioxins and dibenzofurans to vegetables of the cucumber family (Cucurbitaceae) // Environ. Sci. Technol. 1994. №28. рр. 1110-1115.
21) Joner E.J., Corgie S.C., Amellal N., Leyval C. Nutritional constraints to degradation of polycyclic aromatic hydrocarbons in a simulated rhizosphere // Soil Biol. Biochem. 2002. №34. рр. 859-864.
22) Kastner M., Breuer-Jammali M., Mahro B. Enumeration and characterization of the soil microflora from hydrocarbon-contaminated soil sites able to mineralize polycyclic aromatic hydrocarbons (PAH) // Appl. Microbiol. Biotechnol. 1994. №41. рр. 267-273.
23) Koren O., Knezevic V., Ron E.Z., Rosenberg E. Petroleum pollution bioremediation using water-insoluble uric acid as the nitrogen source // Appl. Environ. Microbiol. 2003. №69. рр. 6337-6339.
24) Kuiper I., Kravchenko L.V., Bloemberg G.V., Lugtenberg B.J.J. Pseudomonas putida Strain PCL 1444, selected for efficient root colonization and naphthalene degradation, effectively utilizes root exudate components // Mol. Plant Microbe In. 2002. №15. рр. 734-741.
25) Kuzyakov Y. Review: Factors affecting rhizosphere priming effects // J. Plant Nutr. Soil Sci. 2002. №165. рр. 382-396.
26) Kuzyakov Y., Ehrensberger H., Stahr K. Rhizodeposition of Lolium perenne on two nitrogen levels and enhance of humus decomposition // Soil Biol. Biochem. 2001. №33. рр. 61-74.
27) Li X., Feng Y., Sawatsky N. Importance of soil-water relations in assessing the endpoint of bioremediated soils // Plant growth. Plant Soil. 1997. №192. рр. 219-226.
28) Liste H.-H., Alexander M. Rapid screening of plants promoting phenanthrene degradation // J. Environ. Qual. 1999. №28. рр. 1376-1377.
29) Liste H.-H., Alexander M. Plant-promoted pyrene degradation in soil // Chemosphere 2000. №40. рр. 7-10.
30) Liste H.-H., Alexander M. Accumulation of phenanthrene and pyrene in rhizosphere soil // Chemosphere. 2000. №40. рр. 11-14.
31) Liste H.-H., Rivera A.Q., Tang J. Availability of aged toluene and decane for biodegradation in soil // Water Air Soil Poll. 2002. №133. рр. 227-234.
32) McCutcheon S.C., Jerald L., Schnoor J.L. Phytoremediation: Transformation and Control of Contaminants. John Wiley and Sons, Hoboken, NJ, 2003. 987 pp.
33) Qiu X., Leland T.W., Shah S.I., Sorensen D.L., Kendall E.W. Field study: Grass remediation for clay soil contaminated with polycyclic aromatic hydrocarbons. In: Kruger E.L., Anderson T.A., Coats J.R. (Eds.), Phytoremediation of Soil and Water Contaminants. ACS Symposium Series 664, American Chemical Society, Washington, DC, 1997. pp. 186-199.
34) Reddy B.R., Sethunathan N.J. Mineralization of parathion in the rhizosphere of rice and pearl millet//Agric. Food Chem. 1983. №31. рр. 1379-1381.
35) Richter K. Ackergras. In: Breunig W., Martin B., Wojahn E. (Eds.), Pflanzenproduktion (Futterproduktion). VEB Deutscher Landwirtschaftsverlag, Berlin, 1986. pp. 79-85.
36) Salanitro J.P., Dorn P.B., Huesemann M.H., Moore K.O., Rhodes I.A., Jackson L.M.R., Vipond T.E., Western M.M., Wisniewski H.L. Crude oil hydrocarbon bioremediation and soil ecotoxicity assessment //Environ. Sci. Technol. 1997. №31. рр. 1769-1776.
37) Sawatsky N., Li X. Importance of soil-water relations in assessing the endpoint of bioremediated soils: II. Water-repellency in hydrocarbon contaminated soils // Plant Soil. 1997. №192. рр. 227-236.
38) Schimel J.P., Jackson L.E., Firestone M.K. Spatial and temporal effects on plant microbial competition for inorganic nitrogen in a California annual grassland // Soil Biol. Biochem. 1989. №21. рр. 1059-1066.
39) Schlegel H.G. General Microbiology. 7th Edition (Hardcover), Cambridge University Press, Cambridge, 1993. 843 pp.
40) Schwab A.P., Banks M.K. Biologically mediated dissipation of polyaromatic hydrocarbons in the root zone. In: Anderson T.A., Coats J.R. (Eds.), Bioremediation through Rhizosphere Technology. ACS Symposium Series 563, American Chemical Society, Washington, DC, 1994. pp. 132-141.
41) Schwab A.P., Banks M.K. Phytoremediation of petroleum-contaminated soils: A field study. In: Adriano D.C., Bollag J.-M., Frankenberger Jr. W.T., Sims R.C. (Eds.), Bioremediation of Contaminated Soils. Agronomy Monograph 37, American Society of Agronomy, Madison, WI, 1999. pp. 783-795.
42) Shimp J.F., Tracy J.C., Davis L.C., Lee E., Huang W., Erickson L.E., Schnoor J.L. Beneficial effects of plants in the remediation of soil and groundwater contaminated with organic materials // Environ. Sci. Technol. 1993. №23. рр. 41-77.
43) Sims G.K. Biological degradation of soil. In: Lal R., Stewart B.A. (Eds.), Soil Degradation. Advances in Soil Science 11, Springer-Verlag, New York, 1990. pp. 289-330.
44) Smibert R.M., Krieg N.R. Phenotypic characterization. In: Gerhardt P., Murry R.G.E., Wood W.A., Krieg N.R. (Eds.) Methods for General and Molecular Bacteriology. American Society for Microbiology, Washington, DC, 1994. pp. 607-615.
45) Suslow T.V., Schroth M.N., Isaka M. Application of a rapid method for gram differentiation of plant pathogenic and saprophytic bacteria without staining // Phytopathology. 1982. №72. рр. 917-918.
46) UM-BBD (The University of Minnesota Biocatalysis/Biodegradation Database), 2003. Microbial biocatalytic reactions and biodegradation pathways primarily for xenobiotic, chemical compounds. At: <http://umbbd.ahc.umn.edu/>, Retrieval 2003-12-05.
47) Uren N.C. Types, amounts, and possible functions of compounds released into the rhizosphere by soil-grown plants. In: Pinton R., Varanini Z., Nannipieri P. (Eds.), The Rhizosphere: Biochemistry and Organic Substances at the Soil-Plant Interface. Marcel Dekker, New York, 2001. pp. 19-40.
48) US EPA (U.S. Environmental Protection Agency), 1998. Method 610, Polynuclear aromatic hydrocarbons. In: Methods for Organic Chemical Analysis of Municipal and Industrial Wastewater. <http://www.epa.gov/waterscience/methods/guide/methods.html>.
49) Van Bruggen A.H.C., Semenov A.M. In search of biological indicators for soil health and disease suppression. In: Zeiss, M.R. (Ed.), Soil Health: Managing the Biotic Component of Soil Quality // Appl. Soil Ecol. 2000. (Special Issue) 15. pp. 13-24.
50) Wallnofer P.R., Engelhardt G. Aromatic and heterocyclic structures. In: Rehm H.-J., Reed G. (Eds.), Biotechnology: A Comprehensive Treatise in 8 Volumes. Biotransformations (Vol. 6a), Verlag Chemie, Weinheim, 1984. pp. 277-328.
51) Watkins J.W., Sorensen D.L., Sims R.C. Volatilization and mineralization of naphthalene in soil-grass microcosms. In: Anderson T.A., Coats J.R. (Eds.), Bioremediation through Rhizosphere Technology. ACS Symposium Series 563, American Chemical Society, Washington, DC, 1994. pp. 123-131.
52) Westergaard K., Muller A.K., Christensen S., Bloem J., Sorensen S.J. Effects of tylosin as a disturbance on the soil microbial community // Soil Biol. Biochem. 2001. №33. рр. 2061-2071.
53) Westlake K., Archer D.B., Boone D.R. Diversity of cellulolytic bacteria in landfill // J. Appl. Bacteriol. 1995. №79. рр. 73-78.
54) White J.C. Phytoremediation of weathered pp ’-DDE residues in soil // Internat. J. Phytoremed. 2000. №2. рр. 133-144.
55) Wilson S.C., Jones K.C. Bioremediation of soil contaminated with polynuclear aromatic hydrocarbons (PAHs): A review // Environ. Pollut. 1993. №81. рр. 229-249.
56) Wiltse C.C., Rooney W.L., Chen Z., Schwab A.P., Banks M.K. Greenhouse evaluation of agronomic and crude oil-phytoremediation potential among alfalfa genotypes // J. Environ. Qual. 1998. №27. рр.169-173.
57) Wootton M.A., Kremer R.J., Keaster A.J. Effects of carbofuran and the corn rhizosphere on growth of soil microorganisms //Bull. Environ. Contam. Toxicol. 1993. №50. рр. 49-56.
