Effects of cowpea (Vigna unguiculata) root mucilage on microbial community response and capacity for phenanthrene remediation

J O U RN A L OF E N V I RO N ME N TA L SC IE N CE S 3 3 (2 0 1 5) 4 5– 5 9

Available online at www.sciencedirect.com

ScienceDirect www.journals.elsevier.com/journal-of-environmental-sciences

Effects of cowpea (Vigna unguiculata) root mucilage on microbial community response and capacity for phenanthrene remediation Ran Sun1 , Richard W. Belcher 2 , Jianqiang Liang1 , Li Wang1 , Brian Thater 2 , David E. Crowley 2,⁎, Gehong Wei1,⁎ 1. State Key Laboratory of Crop Stress Biology in Arid Areas, College of Life Sciences, Northwest A&F University, Yangling, Shaanxi 712100, China. E-mail: [email protected] 2. Department of Environmental Sciences, University of California at Riverside, Riverside, CA 92521, USA

AR TIC LE I N FO

ABS TR ACT

Article history:

Biodegradation of polycyclic aromatic hydrocarbons (PAHs) is normally limited by their low

Received 25 September 2014

solubility and poor bioavailability. Prior research suggests that biosurfactants are synthesized as

Revised 23 October 2014

intermediates during the production of mucilage at the root tip. To date the effects of mucilage

Accepted 29 November 2014

on PAH degradation and microbial community response have not been directly examined. To

Available online 13 April 2015

address this question, our research compared 3 cowpea breeding lines (Vigna unguiculata) that differed in mucilage production for their effects on phenanthrene (PHE) degradation in soil. The

Keywords:

High Performance Liquid Chromatography results indicated that the highest PHE degradation

Bioremediation

rate was achieved in soils planted with mucilage producing cowpea line C1, inoculated with

Mucilage

Bradyrhizobium, leading to 91.6% PHE disappearance in 5 weeks. In root printing tests, strings

Phytoremediation

treated with mucilage and bacteria produced larger clearing zones than those produced on

PAH mineralization

mucilage treated strings with no bacteria or bacteria inoculated strings. Experiments with

Rhizosphere

14

C-PHE and purified mucilage in soil slurry confirmed that the root mucilage significantly

enhanced PHE mineralization (82.7%), which is 12% more than the control treatment without mucilage. The profiles of the PHE degraders generated by Denaturing gradient gel electrophoresis suggested that cowpea C1, producing a high amount of root mucilage, selectively enriched the PHE degrading bacteria population in rhizosphere. These findings indicate that root mucilage may play a significant role in enhancing PHE degradation and suggests that differences in mucilage production may be an important criterion for selection of the best plant species for use in phytoremediation of PAH contaminated soils. © 2015 The Research Center for Eco-Environmental Sciences, Chinese Academy of Sciences. Published by Elsevier B.V.

Introduction Biodegradation of polycyclic aromatic hydrocarbons (PAHs) in soil may be enhanced by cultivation of certain plant species that stimulate growth and activity of PAH degrading microorganisms.

This rhizosphere effect is still not clearly understood. Among the different substances that are deposited into the rhizosphere, certain fatty acids, particularly linoleic acid, have shown to greatly increase the biodegradation of phenanthrene (PHE) (Yi and Crowley, 2007). Possible sources of linoleic acid and other

⁎ Corresponding authors. E-mail: [email protected] (David E. Crowley), [email protected] (Gehong Wei).

http://dx.doi.org/10.1016/j.jes.2014.11.013 1001-0742/© 2015 The Research Center for Eco-Environmental Sciences, Chinese Academy of Sciences. Published by Elsevier B.V.

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substances that may influence PAH biodegradation include phospholipids that are deposited from decomposing roots, bacterial extracellular polysaccharides (EPS) and root mucilage. Extensive studies indicated that bacterial EPS stimulate the degradation of PAH by increasing their bioavailability (Jia et al., 2011; Sheng et al., 2010). Likewise, the surfactant properties of root mucilage were evaluated with respect to their ability to lower the surface tension of water and improve water availability to plants (Read et al., 2003). To the best of our knowledge, the role of root mucilage has not been examined in relation to PAH biodegradation. Given the surfactant properties of mucilage and its possible effects on the growth of PAH degrading bacteria, the deposition of mucilage in the rhizosphere can be postulated to have several effects on PAH degrader activity that would affect the rate of PAH degradation in contaminated soils. The composition of root mucilage was investigated for several different plant species, which indicated that root mucilage consist of a complex mixture of substances that could directly or indirectly affect PAH degradation. Mucilage composition varies for different plant species, but generally contains both monosaccharides and polysaccharides that can be used as sole carbon sources by various strains of rhizosphere bacteria that commonly degrade PAHs, such as Rhizobium leguminosarum, Pseudomonas sp. and Burkholderia cepacia (Knee et al., 2001). Root mucilage provides an abundant and rich carbon source for microbial growth in the rhizosphere plant, and affects soil microbial community structure and species composition (Benizri et al., 2007). The differences in the growth rates of bacteria on these substances lead to different process rates, shifts in community structure, or changes in both community structure and process rates, depending on the particular function. For example, Mounier et al. (2004) revealed that the addition of root mucilage stimulated the activity of the denitrifying community, but had only a minor impact on its diversity. López-Gutiérrez et al. (2005) reported that root mucilage added to atrazine contaminated soil temporarily increased the diversity of the atrazine degrading community, but had a negative effect on atrazine mineralization. It is expected that variations in the amount of mucilage production would have an indirect effect on root health by protecting the plant roots from pathogens, decreasing stress caused by desiccation, decreasing mechanical damage to the elongating roots, and protecting them from heavy metal toxicity by retention of metal cations in the rhizosphere (Oades, 1978; Horst et al., 1982; Morel et al., 1986). Differences in the level of mucilage, as well as the chemical composition of the mucilage play a critical role in microbial community structure and activity that would affect organic contaminant remediation. Our research examined the hypothesis that mucilage may increase the biodegradation of PHE and change the PHE degrading bacterial composition that resides in the rhizosphere. To this end, we took advantage of novel F2 breeding lines of the model plant species, cowpea (Vigna unguiculata), that are genetically identical, but that vary in their level of root mucilage production. While working with legume, we inoculated Bradyrhizobium yuanmingense R3 for the nitrogen fixation and growth enhancement. We developed the legume–rhizobia symbiosis model which can be used for PAH in-situ phytoremediation. Our experiments were conducted to examine the mineralization of 14C-PHE in the presence or absence of purified mucilage extracted from plant roots. We

then examined PHE degradation rates and the composition of the PHE degrading bacteria communities in soils planted with different cowpea lines that varied in mucilage production. This is the first study to explore the function of root mucilage in plant-microbe interactions during PAH phytoremediation.

1. Materials and methods 1.1. Plant and soil preparation Soil samples were collected from the soil under turf grass at Hancock Park in Los Angeles, CA, USA. The soil was collected in a location adjacent to the Rancho La Brea Tar Pits and has a long history of exposure to heavy oil that seeps into the soil matrix. The PHE concentration of original soil was 4.8 ± 1.3 mg/kg (n = 3). Air dried soil passed through a 2 mm sieve. Soil physicochemical properties were tested according to Cébron et al. (2009). The soil was a sandy clay loam soil, with pH 7.4 (soil:water of 1:5), organic carbon content 5.3%, inorganic carbon content 1.7%, and total nitrogen 0.34%. The soil was spiked with PHE (dissolved in acetone) to a final concentration of 100 mg/kg soil dry weight (Brinch et al., 2002). The control soil was treated identically, but with acetone solvent without PHE. Cowpea seeds were supplied by Dr. Timothy J. Close (Department of Botany and Plant Sciences, University of California, Riverside, USA). C1 (Line UCR 232): cowpea with high mucilage; C2 (Line: CB27): cowpea with low mucilage; C3 (Line: 63-33(1)): cowpea with no mucilage. The cowpea seeds were surface sterilized by immersion in 75% ethanol for 3 min, followed by stirring in 0.1% HgCl2 for 5 min, and finally rinsed 10 times in sterile distilled water. Surface sterilized cowpea seeds were pre-germinated at 28°C on moistened filter paper in sterile Petri dishes (Deng et al., 2011). B. yuanmingense R3 (GenBank accession number: JN602257) was isolated from surface sterilized nodules of cowpea (V. unguiculata) previously grown in the sample soil. This isolation was authenticated by showing infection and inoculation ability on 3 cowpea breeding lines based on our preliminary experiments. R3 has no ability for PHE degradation, which can eliminate the effect inoculums may have on disturbing the results of the PHE degradation. The seeds of the treatments receiving the Bradyrhizobium inoculums were inoculated with a cell suspension of B. yuanmingense R3. To prepare the cell suspension, R3 was inoculated into YMA liquid medium (10 g mannitol, 0.5 g K2HPO4, 0.2 g MgSO4·7H2O, 0.1 g NaCl, 3.0 g yeast extract, pH 7, per liter water) and the suspension was incubated at 28°C for 3 days. The cultures contained approximately 1.9 × 108 colony forming units per mL (CFU/mL). The inoculation followed the method of Teng et al. (2011).

1.2. Phytoremediation treatments The main experiment compared 10 treatments (Table 1). Each treatment was repeated with 7 replicates. All of the experiments were carried out for 35 days in a plant-growth chamber with 16 hr/8 hr (day/night) photoperiod at 70% humidity. The temperature was maintained at 28°C during the day and 20°C during the evening. Individual plant was planted in 164 mL

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Table 1 – Abbreviation of each treatment. Abbreviation

Treatment

HS (Control 1) HS + R (Control 2) SHS (Control 3) SHS + R (Control 4) SHS + C1 SHS + C2 SHS + C3 SHS + C1 + R SHS + C2 + R SHS + C3 + R

Soil Soil + Bradyrhizobium Spiked soil Spiked soil + Bradyrhizobium Spiked soil + cowpea1 Spiked soil + cowpea2 Spiked soil + cowpea3 Spiked soil + cowpea1 + Bradyrhizobium Spiked soil + cowpea2 + Bradyrhizobium Spiked soil + cowpea3 + Bradyrhizobium

HS: unspiked soil, R: phenanthrene spiked soil.

Bradyrhizobium

yuanmingense,

SHS:

Ray Leach Cone-tainers (Stuewe and Sons Inc., Tangent, Oregon, USA) filled to 2.5 cm below the container surface with PHE treated soil and prepared for each treatment as described in Table 1. All the pots were maintained at 60% water holding capacity. Thirty-five days after planting, the plants were removed from the containers and the roots were gently shaken to remove the bulk soil (BS). The remaining soil that adhered to the roots was considered rhizosphere soil (RS). Soil samples were collected for high pressure liquid chromatography (HPLC) and PCR-DGGE test. Seedlings that were 35 days old were sampled and analyzed for yield parameter test. The dry weight of the plant was determined after oven drying at 105°C for 24 hr. The leaf area was measured by using a portable area meter (LI-COR Model Li-3000, Lambda Instruments, Lincoln, Nebraska, USA). The chlorophyll content in leaves was estimated by using the chlorophyll meter (SPAD-502, Minolta Camera, Osaka, Japan).

1.3. Root mucilage collection Root mucilage samples were collected according to the methods described by Morel et al. (1986) and Mary et al. (1992). Fifty cowpea seeds of each breeding line were surface sterilized and allowed to germinate for 3 days. After the cotyledons emerged, the seeds were transferred to beakers for aeroponic culture (Pan et al., 2004; Cai et al., 2011). When the primary roots were 3 cm long, the root tips were sprayed with sterile water to release the mucilage, which was removed using a sterile glass capillary tube. Root mucilage was extracted every 12 hr for 15 days. Three replicates for each treatment were designed. The mucilage samples were combined, vortexed, and centrifuged for 30 min at 5000 ×g to remove cell debris. Mucilage was separated from the border cells by filtering through a 0.45-μm nylon filter (Nagahashi and Douds, 2004). The material was then lyophilized and stored at −80 C before use in further experiments. Root mucilage production of C1 and C2 was 3.4 ± 0.8 and 0.5 ± 0.3 mg dry weight mucilage/g dry roots separately. Root mucilage production of C3 was not detected.

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dish containing mineral salts agar (1.0 g KNO3, 0.38 g K2HPO4, 0.2 g MgSO4, 0.05 g FeCl3, and 15 g agar per liter water) with PHE as sole carbon source. The PHE was applied as a microcrystalline film by sublimation to produce a haze film on the agar surface (Alley and Brown, 2000). Biodegradation of PHE was observed by the formation of clearing zones on the PHE film. The radiuses of the clearing zones were measured and pictures were taken on days 3 and 7. To compare the effects of intact roots and mucilage on the growth of PHE degrading bacteria, sterilized cotton string segments treated with mucilage and rhizospheric bacterial suspensions were used as surrogates for plant roots. Control roots were also prepared by sterilization of root segments using 5% NaClO for 5 min, followed by 10 sterilized water rinses. Sterilized strings soaked with the rhizospheric bacteria solution (5 g root/mL) and fresh roots with rhizospheric bacteria were compared on the same PHE sublimation plates for 7 days (Alley and Brown, 2000). Sterilized strings were soaked, both with the cowpea C1 mucilage solution (1 mg/mL sterilized H2O) and the rhizospheric bacteria solution, then compared with a control treatment using autoclaved sterilized string segments soaked with mucilage solution.

1.5. Quantitative analyses of PHE disappearance The procedure used for extracting PHE from the soil was a modified version of EPA method 8100. Freeze dried soil (5 g) was collected at day 0 and day 35 for analyses of residual PHE. The initial concentration of PHE was measured within 1 hr after spiking. Prior to extraction, internal standard of 1 mL of 100 mg/L pyrene was used to quantify the PHE losses. The soils were then extracted twice by sonication in a solvent mixture for 30 min using 40 mL 1:1 (V/V) dichloromethane and acetone mixture (Yi and Crowley, 2007). The liquid phase was concentrated into a small volume (3–5 mL) using a rotary evaporator, dried under a gentle stream of nitrogen gas, and solvated 4 times with 0.5 mL hexane. The concentrated extracts were purified using Supelclean™ Lc-Florisil SPE Tubes (Sigma-Aldrich, USA) 6 mL (1 g) by EPA Method 3620B, with 20 mL of 7:3 (V/V) hexane and dichloromethane. After a final drying under nitrogen gas, the residues were dissolved in 2 mL of methanol. Quantification was performed using HPLC (Agilent 1100, Hewlett-Packard, Waldbronn, Germany) with a UV diode array detector at 245 nm. A 150 × 4.6 mm, 3.5 μm particle size Agilent ZORBAX Eclipse PAH column (Agilent Technologies Inc., Santa Clara, CA, USA) was used with a 9:1 (V/V) methanol/water gradient. The mobile phase flow rate was 1 mL/min and the injection volume was 25 μL. The recoveries of this method were in the range of 78.3%–112.6% and with a relative standard deviation (RSD) of less than 4.6%. The dissipation rate of PHE in soil was calculated by Eq. (1):

PHE% ¼ ðC 0 −C 35 Þ=C 0  100%

ð1Þ

1.4. Root printing Plant roots of three cowpea species were collected after 35 days using flame-sterilized forceps. The root of 1 cm length was cut and manually pressed onto agar medium in a Petri

where, PHE% is the dissipation rate of PHE, C0 (mg/kg) is the initial PHE concentration in soil, and C35 (mg/kg) is the remaining PHE concentration in soil at day 35.

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1.6. 14C-PHE mineralization experiments 14

C-PHE (100 μCi in 2 mL methanol) was used to examine the rate of mineralization in the presence and absence of mucilage. Mineralization experiments compared the rates of PHE mineralization by microflora indigenous to the plant experiments soil, with that achieved by a pure strain of a PHE degrader (Delftia tsuruhatensis D17, GenBank accession number: JN590248). Strain D17 can degrade PHE effectively according to our unpublished research before. The soils were air dried, passed through a 2 mm sieve, and spiked with a mixture of radio labeled and non-radio labeled PHE using 0.01 μCi/g soil 14C-PHE in a final concentration of 12C-PHE at 100 mg/kg, following the method of Doick and Semple (2003). Respirometers consisting of 125 mL conical flasks with rubber stoppers and sealed with teflon were prepared to contain 10 g soil and 30 mL minimal salt solution (1.0 g KNO3, 0.38 g K2HPO4, 0.2 g MgSO4, 0.05 g FeCl3 per liter water) yielding a 1:3 (V/V) of soil:liquid suspension. Respirometry of the soil slurries was conducted for 20 days, in triplicate. Respired CO2 was trapped using 1 mol/L NaOH (1 mL) in 7 mL glass scintillation vials inside each respirometer. Cowpea C1's Root mucilage was added to the treatment with a concentration of 1 mg/mL, which is the critical micelle concentration in the preliminary test. During the experiment, we examined mucilage effects on a known PHE degrader; cells of Delftia tsuruhatensis D17 were cultured to midlog phase and a cell suspension was prepared to optical density (OD600 = 1.0), from which 500 μL was added to each flask. The flasks were incubated at 28°C and 100 r/min. At selected time intervals, the CO2 traps were quickly removed and replaced. The 14C activity was determined by liquid scintillation counting (Canberra Packard Tri-Carb 2250CA liquid scintillation analyzer, USA) with external source calibration for conversion of cpm to dpm.

CGGGAACGTATTCACCG-3′) with GC-clamps (CGC CCG CCG CGC CCC GCG CCC GGC CC GCC GCC CCC GCC CC) (Eurofins MWG Operon) attached to the forward primer (Muyzer et al., 1993). These primers target a 454 bp sequence of the 16S rRNA V6, 7, and 8 regions. Bacterial 16S rRNA genes were amplified by touch-down PCR using the following program: 95°C for 9 min, followed by 19 cycles at 94°C for 1 min, 65°C (decrease 0.5°C per cycle) for 1 min, and 72°C for 3 min, followed by 9 cycles of 94°C for 1 min, 55°C for 1 min, and 72°C for 3 min, with a final extension at 72°C for 7 min. The size of the PCR product was confirmed by electrophoresis in 1% agarose gel and staining with ethidium bromide for 5 min.

1.9. Denaturing gradient gel electrophoresis analyses Denaturing gradient gel electrophoresis (DGGE) was conducted using a DCode™ Universal Mutation Detection System (Bio-Rad, Hercules, CA, USA). Samples of PCR product were loaded onto polyacrylamide gel with linear denaturing gradient (50% to 70% denaturant; 80% denaturant is 5.6 mol/L urea and 32% (V/V) formamide). Electrophoresis was performed at 100 V and 55°C for 720 min. Gels were stained for 20 min using SYBR Gold (1:1000 dilutions, Invitrogen, Carlsbad, CA, USA) and photographed using a Bio Rad Gel Doc 2000 Gel Documentation System (Bio-Rad, Hercules, CA, USA). Two replicates showed the same DGGE gel pattern in preliminary experiments. One replicate was randomly selected to conduct further DGGE analyses. The Shannon index and peak intensity was determined based on analyses of the photograph using the Bio Rad Quantity One software (Bio-Rad, Hercules, CA, USA). The Shannon index (H) and Evenness (EH) of the bacterial community were calculated by Eqs. (2) and (3) (Hill et al., 2003; Hu et al., 2007). H¼−

Xs

P i¼1 i

lnP i

ð2Þ

1.7. Enrichment culture

EH ¼ H= lnS

Five gram samples of RS and BS soil were collected from each treatment, and suspended in 200 mL of MS medium. Two replicates for each sample were designed. PHE (100 mg/L) was added as enrichment substrate and the suspension was incubated with shaking at 180 r/min at 28°C in the dark. Seven days later, a 20 mL aliquot was transferred to 180 mL of a fresh MS medium with the same PHE concentration as above and incubated under the same conditions. This procedure was repeated three times. The 200 mL aliquot of the third enrichment culture was centrifuged for 30 min at 5000 ×g to collect bacterial cells produced from the enrichment cultures.

where, H is the value of the Shannon index, Pi is the ratio of the specific band intensity to the total intensity of all bands in a lane and s is the number of bands in the sample (Shannon, 1948). The Richness (S) of the bacterial community was determined from the number of bands in each lane (Dong and Reddy, 2010).

1.8. DNA extraction and PCR amplification Total DNA was extracted from the RS and BS bacterial cell pellet using a MasterPure™ DNA Purification Kit (Epicentre, Madison, WI, USA). PCR was performed using the following reaction mixtures: 12.5 μL of 2 × Go to Taq Green Master Mix, 0.5 μL of each primer (10 μmol/L), 0.5 μL (5–15 ng) of template DNA, and 11 μL of nucleases-free water to give a final volume of 25 μL. The primers used for PCR were the EuBf933 (5′-GCACAAGCGGTGGAGCATGTGG-3′) and EuBr1387 (5′-GCC

ð3Þ

1.10. Microbial species identification Predominant DGGE bands (1–22) were excised and transferred to a clean eppendorf tube, washed with Milli-Q water, and incubated in 50 μL of Milli-Q water at 4°C overnight. After centrifugation at 11,000 ×g for 60 sec, the supernatant was transferred to a new tube, from which 2 μL was used as the template for a second round of PCR-DGGE analyses to check the band position and purity (Zhang et al., 2009). PCR products from the second round of DGGE were purified using the Wizard PCR prep DNA purification system (Promega, Madison, WI, USA) and cloned using a pGEM-T Easy Vector Systems Kit (Promega, Madison, WI, USA). The 16S rRNA genes were sequenced with an ABI 3730 DNA Analyzer and the sequences were submitted to the GenBank database (http://www.ncbi.nlm.nih.gov) with the following accession numbers: from JN565946 to JN565967.

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1.11. Statistical analyses

2.2. Root printing

Analyses of one-way or two-way analysis of variance (ANOVA) tests followed by a Ducan's test were performed with IBM SPSS Statistics 19 to assess significant differences among treatments, and the significance level was P < 0.05. The differences in growth parameters of each cowpea species were compared by one-way ANOVA test. The two-way ANOVA test was employed to determine any differences in growth parameters and PHE degradation rate, with three cowpea species and presence/absence of Bradyrhizobium inoculation as the two factors. If the ANOVA results were significant, Ducan's multiple comparison method was used to determine the difference. Pearson's correlations were assessed between PHE dissipation rate (RPHE) and biological parameters (IBM SPSS Statistics 19). BLAST tools were used to evaluate the similarity of sequences obtained from the DGGE bands with other 16S rRNA gene sequences in the NCBI database. DNA MAN was used to compare multiple sequences, and the Neighbor-Joining method in the MEGA version 5.0 software packages was used for cluster analyses and the construction of phylogenetic trees.

A root printing technique was used to reveal the presence and distribution of PHE degrading bacteria by the formation of clearing zones on agar medium containing a thin film of PHE. The radius of clearing zone was measured and data is shown in Table 3. This method revealed that no clearing zones formed around the sterilized roots, indicating that the disappearance of PHE was biologically driven and did not involve simple solubilization by surfactant materials contained in lysates or exudates from the root tissues (Fig. 1a). Root segments with live bacteria and sterile cotton strings inoculated with RS bacteria both formed clearing zones on the PHE assay medium (Fig. 1b and c). Non-sterile roots with associated PHE degrading bacteria produced larger clearing zones than cotton strings inoculated with bacteria after day 3 and 7 observation, suggesting residual root derived carbon stimulated growth linked degradation of PHE on the agar medium. Likewise, strings treated with mucilage and bacteria (Fig. 1f) produced larger clearing zones than those produced on mucilage treated strings with no bacteria (Fig. 1d) or bacteria inoculated strings (Fig. 1e). The radius of clearing zones formed by plant roots C1, C2 and C3 did not show any significant difference on day 3. However, plant root C1 appeared to form clearing zone significantly bigger than C2 and C3 based on the radius data on day 7, while the result between C2 and C3 was not significant (Table 3).

2. Results 2.1. Plant growth Several plants' parameters were monitored after 35 day growth (Table 2). Cowpea C1 showed the significant higher value than cowpea C2 and C3 on shoot height, root length, leaf area, number of nodules, fresh root-biomass and dry root-biomass accumulation after 35 day growth in the treatments, with or without B. yuanmingense R3 inoculation. However, cowpea C2 and C3 did not show any significant difference between each other on most parameters, except leaf chlorophyll content, dry root biomass and fresh shoot biomass. From the perspective of biomass accumulation, cowpea C1 indicated a higher tolerance to PHE than cowpea C2 and C3. With addition of B. yuanmingense R3, the number of nodules (220%), nodule weight (142%), and the plant biomass (27.5%) of cowpea C1 increased significantly. However, cowpea C2 and C3 did not show any significant difference on most parameters, except leaf chlorophyll content and root length. Root fresh biomass of C2, number of nodules of C3 showed an increase due to the B. yuanmingense R3 inoculation. However, the shoot height, leaf area, and the shoot dry biomass of all three cowpea lines did not show significant change based on the R3 addition. The result indicated that the B. yuanmingense R3 impacts the underground parts more than the above-ground parts. The two-way ANOVA results indicated that the effects of cowpea species were significant at P < 0.001 level for all the plant parameters except the nodule weight were significant at P < 0.01 level. The Bradyrhizobium inoculation was significant at P < 0.01 level for all the plant parameters except for shoot height and leaf area. The interactions between cowpea species and Bradyrhizobium inoculation was also significant for the number of nodule (P < 0.001) and fresh shoot biomass (P < 0.05).

2.3. Comparison of PHE removal from soil by cowpea lines differing in mucilage production Three different cowpea cultivars C1, C2 and C3 were tested for their ability to enhance the dissipation of PHE from contaminated soil. After 35 days, There was no difference between the two control treatments that examined PHE disappearance in soil without plants, or in unplanted soil inoculated with B. yuanmingense (48.4% and 51.2%, respectively) (Table 4). In the treatments evaluating the effects of the 3 cowpea lines, inoculation with B. yuanmingense R3 resulted in greater disappearance of PHE as compared to non-inoculated plants. The greatest disappearance of PHE occurred in the treatment using the cowpea line C1 that produced high mucilage. In the treatments with the R3 inoculation, PHE dissipation rate for soils planted with the cowpea lines C1, C2 and C3 was 91.6%, 76.8% and 77.2% respectively. Soils planted with cowpea lines C2 and C3 appeared to remove equivalent amounts of PHE as the difference between the two treatments was not statistically significant. According to two-way ANOVA, the effects of both cowpea species and Bradyrhizobium inoculation on PHE degrading rates were significant (for cowpea F = 18.271, P < 0.001; for Bradyrhizobium F = 28.641, P < 0.001) but not the interaction effects between them (for interaction F = 0.374, P > 0.05).

2.4. Impact of cowpea mucilage on 14C-PHE mineralization 14

C-PHE mineralization was determined by radiorespirometry analyses. The mineralization rate of 14C-PHE in the soil slurry treatment with mucilage (HS + RM) was greater than the treatment without mucilage added (Fig. 2). Maximum 14CO2 recovery with root mucilage (82.7%) was 12% higher than in

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Table 2 – Plant yield parameters of each treatment. Treatment SHS SHS SHS 1 F SHS SHS SHS 1 F

Shoot height (cm)

Root length (cm)

Leaf area (cm2)

Chlorophyll content of leaf (SPAD)

Number of nodules

12.5 ± 0.38 a 8.5 ± 0.29 b 8.7 ± 0.59 b 37.779 ⁎⁎⁎ 12.9 ± 0.68 a 10.2 ± 0.73 b 9.9 ± 0.67 b 11.719 ⁎⁎

18.3 ± 1.17 a 15.1 ± 0.28 b 15.4 ± 0.66 b 6.872 ⁎⁎ 18.5 ± 0.54 a 17.1 ± 0.42 b 16.5 ± 0.84 b 7.000 ⁎⁎

119.5 ± 5.07 a 60.0 ± 6.10 b 57.2 ± 8.06 b 29.157 ⁎⁎ 96.0 ± 3.52 a 67.5 ± 6.28 b 53.5 ± 4.07 b 20.673 ⁎⁎

45.1 ± 2.03 a 32.2 ± 1.47b 30.4 ± 1.29 b 24.230 ⁎⁎⁎ 47.0 ± 2.19 a 45.3 ± 3.35 a 37.4 ± 2.20 b 3.806 ⁎

10.0 ± 1.15 a 8.9 ± 0.63 b 4.3 ± 0.81 b 11.355 ⁎⁎ 32.0 ± 2.70 a 11.3 ± 1.36 b 10.3 ± 1.02 b 42.246 ⁎⁎⁎

13.167 ⁎⁎⁎ 16.710 ⁎⁎⁎ 0.690

48.627 ⁎⁎⁎ 1.989 3.775

15.647 ⁎⁎⁎ 16.671 ⁎⁎⁎ 3.250

53.352 ⁎⁎⁎ 110.059 ⁎⁎⁎ 19.362 ⁎⁎⁎

+ C1 + C2 + C3 + C1 + R + C2 + R + C3 + R

F value of two-way ANOVA Cowpea 37.489 ⁎⁎⁎ Bradyrhizobium 6.675 Interaction 0.088 Treatment

SHS SHS SHS 1 F SHS SHS SHS 1 F

Nodule weight (g)

Shoot fresh biomass (g)

0.07 ± 0.01 0.07 ± 0.01 0.05 ± 0.01 2.192 0.17 ± 0.02 a 0.09 ± 0.01 b 0.06 ± 0.02 b 8.797 ⁎

3.4 ± 0.40 a 3.1 ± 0.28 a 2.7 ± 0.13 b 8.958 ⁎⁎ 4.4 ± 0.21 a 3.1 ± 0.24 b 2.9 ± 0.30 b 24.065 ⁎⁎⁎

30.712 ⁎⁎⁎ 24.146 ⁎⁎⁎ 4.496 ⁎

+ C1 + C2 + C3 + C1 + R + C2 + R + C3 + R

F value of two-way ANOVA Cowpea 10.920 ⁎⁎ Bradyrhizobium 32.490 ⁎⁎⁎ Interaction 3.240

Root fresh biomass (g)

Shoot dry biomass (g)

Root dry biomass (g)

0.6 ± 0.03 a 0.4 ± 0.05 b 0.2 ± 0.05 b 13.443 ⁎⁎ 0.9 ± 0.03 a 0.6 ± 0.08 b 0.4 ± 0.02 b 32.612 ⁎⁎⁎

1.3 ± 0.05 a 1.1 ± 0.09 ab 0.8 ± 0.08 b 7.237 ⁎ 1.4 ± 0.07 a 1.2 ± 0.07 b 1.0 ± 0.12 b 8.790 ⁎⁎

0.3 ± 0.02 a 0.2 ± 0.03 b 0.1 ± 0.02 b 18.297 ⁎⁎ 0.5 ± 0.05 a 0.3 ± 0.04 b 0.2 ± 0.01 c 14.668 ⁎⁎

42.312 ⁎⁎⁎ 30.604 ⁎⁎⁎ 1.389

15.601 ⁎⁎⁎ 5.259 ⁎ 0.158

31.244 ⁎⁎⁎ 34.711 ⁎⁎⁎ 0.643

1

F: F value of one-way ANOVA. For each parameter, values sharing the same letter are not significantly different (Duncan's Test, P < 0.05). Data are presented as mean ± standard error, n = 3. SPAD: soil and plant analyzer development. ⁎⁎⁎ P < 0.001. ⁎⁎ P < 0.01. ⁎ P < 0.05.

the treatment without mucilage (70.7%). Sterilized control treatments confirmed that the PHE was not degraded. Therefore, any evolved 14CO2 was assumed to be the result of microbiological activity. The effect of root mucilage on PHE degradation by the pure bacterium strain Delftia tsuruhatensis D17 was tested. The presence of root mucilages lowered the mineralization rate for

the first 6 days, then 14C-PHE mineralization was significantly enhanced (P < 0.01) by the presence of root mucilage. The mineralization rate of D17 + RM treatment was 62.5%, which was 8.8% higher than the D17 treatment without added mucilage (53.7%) at the end of the mineralization experiment.

Table 3 – Radius of clearing zone formed on PHE sublimation plates.

The structural diversity of the PHE-degrading microbial community was examined by Shannon index (H), Evenness (EH) and Richness (S) (Table 5). The SHS + C1 + R rhizosphere samples with the highest Richness (S = 13) and Shannon index (H = 2.50) showed the most complex profiles, indicating the presence of a great variety of PAH-degrading bacterial species. Among the BS treatments, SHS + C1 showed the highest Richness (S = 13) and Shannon index (H = 2.48). The Shannon index from the C1 treatments was markedly higher than the other two plants (C2, C3) in the same process, which suggests that the three cowpea breeding lines selected different bacterial communities. Among the treatments with Bradyrhizobium R3 added, the scores of S and H for PHE degrading bacteria in rhizosphere were higher than bulk soil treatments. The Shannon index from rhizosphere treatments

Treatment

Radius of clearing zone (mm) 3 days

Sterilized root Plant root C1 Plant root C2 Plant root C3 String + bacteria String + bacteria + mucilage

0 4.3 3.3 3.7 2.3 3.7

± ± ± ± ± ±

0.0 0.3 0.3 0.3 0.3 0.3

7 days c a a a b a

0 8.0 5.7 6.7 4.7 8.7

± ± ± ± ± ±

0.0 d 0.6 a 0.3 bc 0.3 b 0.3 c 0.3 a

Data are presented as mean ± standard error, n = 3. For each parameter, values sharing the same letters are not significantly different (Ducan's Test, P < 0.05).

2.5. DGGE analyses of the PHE degrading bacterial community

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Fig. 1 – Phenanthrene sublimation plates showing clearing zones around cowpea C1 roots or cotton string (root surrogates) with or without root mucilage: (a) sterilized root segment after 7 days; (b) cotton string inoculated with rhizosphere bacteria suspension without mucilage after 3 days (left), cowpea C1 plant root print after 3 days (right); (c) cotton string inoculated with rhizosphere bacteria suspension without mucilage after 7 days (left), cowpea C1 plant root print after 7 days (right); (d) sterilized cotton strings treated with root mucilage and left on plate for 7 days; (e) cotton string treated with bacteria and left on plate for 7 days; and (f) cotton strings inoculated with rhizosphere bacteria and treated with mucilage, 7 days bacterial growth.

with Bradyrhizobium R3 inoculation was higher than the treatments without inoculation, which suggests that the rhizosphere communities were more diverse due to the addition of Bradyrhizobium R3. However, there was no apparent effect of Bradyrhizobium R3 inoculation on the communities in the bulk soil. For the control treatments without plants, the scores of S and H were increased after the soil spiked with PHE.

DGGE profiles of the PHE degrading bacterial communities from each treatment are shown in Fig. 3. The identities of bacterial species that were represented by individual DNA OTUs in the DGGE gels were determined by excising the bands and DNA sequencing. The phylogenetic distribution of prominent 16S rRNA gene sequences from samples of different treatments is shown in Table 6. In C1 + R treatment, the predominant bacteria species were Labrysmonachus (band 12),

Table 4 – Extractable concentrations of the PHE at day 0 and day 35 in the spiked soil. Treatment

C0 (mg/kg)

C35 (mg/kg)

SHS SHS + R SHS + C1 SHS + C2 SHS + C3 SHS + C1 + R SHS + C2 + R SHS + C3 + R Cowpea Bradyrhizobium Interaction

103.3 ± 4.2 99.2 ± 3.2 98.7 ± 2.9 103.0 ± 4.4 98.4 ± 5.2 98.3 ± 3.2 97.8 ± 3.8 102.3 ± 1.9 F = 18.271 ⁎⁎⁎ F = 28.641 ⁎⁎⁎

53.2 ± 3.4 48.3 ± 2.1 21.1 ± 2.0 32.9 ± 2.4 32.8 ± 2.1 8.3 ± 1.4 22.5 ± 2.0 23.2 ± 2.4 P = 0.000 P = 0.000 P = 0.695

F = 0.374

Dissipation rate (%) 48.4 51.2 78.6 68.1 66.3 91.6 76.8 77.2

± ± ± ± ± ± ± ±

3.1 d 2.7 d 1.8 b 1.8 c 3.7 c 1.2 a 2.8 b 2.8 b

C0: just after spiking, C35: 35 days after spiking. Data are presented as mean ± standard error, n = 3. For each parameter, values sharing the same letter are not significantly different (Duncan's Test, P < 0.05). ⁎⁎⁎ P < 0.001.

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Mineralizsation of 14C phenanthrene (%)

90

82.7%

80 70.7%

70

62.5%

60

53.7%

50 HS+RM HS Control D17 D17+RM

40 30 20 10 0

0.09% 0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 Assay time (day)

Fig. 2 – Mineralization 14C-PHE in soil spiked with 100 mg/kg PHE by indigenous PHE degraders or by a strain of PHE degrading bacteria, Delftia tsuruhatensis D17, in soil with or without added root mucilage (1 mg/mL). Letters indicate statistically significant differences between the treatments at last sample date (P < 0.01, Duncan's Test). The error bars indicate the standard error of the means (n = 3).

Mesorhizobium sp. (band 14), Ochrobactrum sp. (band 7), Sphingomonas sp. (band 10), Pseudomonas putida (band 9), Methylovorus sp. (band 8) and Geobacillus pallidus (band 19). Even though B. yuanmingense R3 was inoculated into these treatments, there was no band of Bradyrhizobium sp. detected in the DGGE profile. This revealed that B. yuanmingense R3 was not PHE degrading bacteria. Ochrobactrum sp. (band 7), Sphingomonas sp. (band 10) and Mesorhizobium sp. (band 14) were all detected in the rhizospheric soil from C1 + R, C2 + R and C3 + R treatments. Taxonomic analyses indicated that the clones were divided into 5 groups, α-Proteobacteria, β-Proteobacteria, γ-Proteobacteria, Firmicutes, and Bacteroidetes (Fig. 4), and the percentages of each group in different treatments are illustrated in Fig. 5. Most of the Table 5 – Bacterial diversity indices are based on DGGE analyses of 16S rRNA gene fragments.

Bulk soil

Rhizosphere

Control

Treatment

S

EH

H

SHS + C1 SHS + C2 SHS + C3 SHS + C1 SHS + C2 SHS + C3 SHS + C1 SHS + C2 SHS + C3 SHS + C1 SHS + C2 SHS + C3 HS HS + R SHS SHS + R

13 5 2 6 2 3 9 5 5 13 6 9 2 3 7 5

0.97 0.90 0.92 0.95 0.98 0.82 0.89 0.98 0.92 0.97 0.89 0.96 0.99 0.98 0.99 0.88

2.48 1.45 0.64 1.71 0.68 0.90 1.96 1.57 1.48 2.50 1.60 2.11 0.69 1.08 1.93 1.41

+R +R +R

+R +R +R

S: species Richness, EH: Shannon's Evenness, H: Shannon index.

treatments consisted of bacteria from more than one phylum except Control HS + R and (BS) SHS + C3. These two treatments were dominated by α-Proteobacteria. Compared with the composition of bulk soil bacterial community among the three cowpea line treatments, cowpea C1 treatment revealed four phyla (α-Proteobacteria; β-Proteobacteria; γ-Proteobacteria; Bacteroidetes) with or without Bradyrhizobium innoculation, which is the most among the cowpea lines which were treated equally. Bacteria from β-Proteobacteria were unique in the soil planted with cowpea C1, however, after the Bradyrhizobium inoculation, bacteria from β-Proteobacteria were introduced in the C2 and C3 rhizosphere soil treatment. Bacteria from Bacteroidetes phyla were detected in the bulk soil of C1 treatment; however, bacteria from this group disappeared in C1 rhizospheric soil treatment. There were no bacteria from Firmicutes phyla in any C1 treatment. In C1 rhizosphere treatment, the proportion of α-Proteobacteria and γ-Proteobacteria slightly decreased and β-Proteobacteria increased due to the addition of Bradyrhizobium. The band peak intensity of DGGE profiles from three cowpea lines treatments was compared and the complex results indicated the change of a composition and quantity of the PHE-degrading bacterial community (Fig. 6). The positions (Rf) and peaks (Int), corresponded to individual microbial identities and the band intensities. Fig. 6a reflects that there are 14 identifiable peaks (intensity > 20), 8 peaks from C1 treatment (red line), 2 peaks from C2 treatment (green line) and 4 peaks from C3 treatment (yellow line). However, with the inoculation of Bradyrhizobium, Fig. 6b indicates there are 19 identifiable peaks (intensity > 20), 10 peaks from C1 treatment (red line), 6 peaks from C2 treatment (green line) and 4 peaks from C3 treatment (yellow line). This indicated that the inoculation of Bradyrhizobium also played a role in changing PHE-degrading bacterial community structure in rhizosphere. Some peaks overlapped and illustrated that even though the PHE-degrading bacteria varied greatly in the rhizosphere of three cowpea lines, they harbored the same PHE-degrading bacteria species. There is one overlapping group in Fig. 6a and four overlapping groups in Fig. 6b. Cowpea C1 showed more peaks than C2 and C3, which indicates there are more PHE-degrading bacteria species in C1 rhizosphere. The peak intensity results illustrated that PHE-degrading bacterial community structure in treatments of different cowpea lines varied greatly. In conclusion, the Shannon index and PCR-DGGE profile provide valuable information about major population or community shifts in general. The bacterial community changed, not only in regard to species diversity (indicated by Shannon index) and Richness (represented by number of OTUs), but also in its structure (showed by phylogenetic analyses).

2.6. Phenanthrene degradation and biological parameters The Pearson's correlation coefficient obtained between PHE dissipation rates and biological parameters is shown in Table 7, including botany parameters according to growth forms and microbiology parameters according to DGGE profiles. Significant correlations (P < 0.05) were found between most of the botany parameters and PHE dissipation rates except the leaf

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Bulk soil

Rhizosphere soil

Control

area. Strong significant correlations (P < 0.01) were detected between PHE dissipation rates and root biomass (both dry and fresh). The results indicated that root biomass is one of the most important botany factors that impact PHE dissipation. With regard to microbiology parameters, the rhizosphere Shannon index and Richness correlated with PHE dissipation significantly (P < 0.05). However, bulk soil Shannon index and Richness correlated with PHE dissipation negatively, thus indicating that microbial communities in rhizosphere contribute more to the PHE dissipation than microbial communities in bulk soil.

3. Discussion

Fig. 3 – 16S rRNA gene profiles of PHE degrading bacteria generated from different treatments. Lanes 1–6: bulk soil samples from treatment SHS + C1, SHS + C2, SHS + C3, SHS + C1 + R, SHS + C2 + R and SHS + C3 + R; lanes 7–12: rhizosphere soil samples from treatment SHS + C1, SHS + C2, SHS + C3, SHS + C1 + R, SHS + C2 + R and SHS + C3 + R; lane 13: unspiked soil control without plant; lane 14: unspiked soil control inoculated with Bradyrhizobium without plant; lane 15: PHE spiked soil control without plant; lane 16: PHE spiked soil control inoculated with Bradyrhizobium without plant; lane M: DNA ladder. Bands 1–22 were cloned and sequenced for identification of bacterial species.

Root mucilage has several important functions that include facilitating root penetration, soil aggregate formation, and alleviation of heavy metal toxicity, but has not been previously investigated with regard to its potential role in PAH degradation. In this research, we evaluated the effects of cowpea root mucilage using purified mucilage and conducted experiments with 3 cowpea breeding lines that differed in mucilage production. Plant experiments confirmed that cowpea C1 with high root mucilage production stimulates the disappearance of PHE significantly when compared with the other two cowpea lines (Table 4). Use of an aged contaminated soil resulted in relatively fast disappearance rates, with approximately half of the added PHE disappearing in the unplanted control soils after 5 weeks, irrespective of inoculation with Bradyrhizobium (48.4% and 51.2%, respectively). This indicated that PHE-degrading bacteria existed in the aged contaminated soil intrinsically. The presence of cowpea C1 plants enhanced the rate of disappearance another 40.4%. The loss of PHE was about 10% lower in the treatments with cowpea lines C2 and C3. However, we cannot assert that root mucilage was the main reason that leads to the PHE

Table 6 – Sequence results from band excised from Fig. 3. DGGE band 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22

Accession number

Blast search results and accession number

Score ⁎

PAH

PGPRb

Biosurfactant

JN565946 JN565947 JN565948 JN565949 JN565950 JN565951 JN565952 JN565953 JN565954 JN565955 JN565956 JN565957 JN565958 JN565959 JN565960 JN565961 JN565962 JN565963 JN565964 JN565965 JN565966 JN565967

Labrys monachus (NR025581) Labrys monachus (NR025581) Methylobacterium sp. (HQ220096) Bacteroidetes bacterium (AY337599) Filobacillus sp. (DQ664537) Xanthomonas sp. (AF513452) Ochrobactrum sp. (HQ670703) Methylovorus sp. (HQ380796) Pseudomonas putida (HQ622810) Sphingomonas sp. (GQ273963) Uncultured bacterium clone (GQ037609) Labrys monachus (NR025581) Stenotrophomonas rhizophila (JF700462) Mesorhizobium sp. (EF100516) Methylobacterium sp. (JF753450) Azospirillum sp. (AB049112) Methylobacterium sp. (HQ220096) Azospirillum sp. (AB049112) Geobacillus pallidus (HQ324908) Serratia marcescens (JF719838) Azospirillum lipoferum (DQ787330) Azospirillum sp. (AY118223)

97% 97% 99% 99% 98% 100% 100% 99% 99% 98% 98% 98% 98% 99% 99% 98% 99% 99% 99% 98% 99% 99%

+c + + + + + + + + + NR + + + + + + + + + + +

NRd NR NR NR NR + NR + + NR NR NR NR + NR + NR + + + + +

NR NR NR NR NR NR NR NR + NR NR NR NR NR NR NR NR NR + NR NR NR

PGPR: Plant Growth Promoting Rhizobacteria; +: positive reports; NR: no reports. ⁎ Score values indicate the percentage of 16S rDNA which is similar to bacteria that are referenced in the BLAST database by these accession numbers.

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Fig. 4 – Phylogenetic analyses of twenty two 16S rRNA gene bands obtained from the RS bacteria and BS bacteria sequenced for bacterial identification. The branching pattern was generated by the neighbor-joining method. The topology shown was obtained using 1000 bootstrap replication. The numbers at the nodes indicate the levels of bootstrap support percentage based on 1000 resamplings.

degradation, since the plant yield parameters were different among the three cowpea breeding lines. We can only conclude that cowpea C1 is superior in PHE phytoremediation by showing higher biomass accumulation and higher PHE degradation rate among the three cowpea breeding lines. To further explore the direct function of root mucilage on PHE degradation, we applied root printing technique, which proved to be a simple method to visually assess PHE degradation. Root mucilage increased the PHE clear zones for both plant roots and cotton strings that were used as root surrogates (Fig. 1). The formation of clearing zones were dependent on the presence of an active microflora and was greatest for root prints produced from live root segments, suggesting that both the presence of

mucilage and other substances released from the roots promoted PHE degradation. 14 C-PHE mineralization test provides a quantitative way to analyze the effect of root mucilage on PHE degradation directly. The differences in the mineralization rate of PHE in soil microcosms amended with mucilage generally corresponded to the observations from the plant study. The mineralization curves generated for the PHE degrader Delfia tsurohatensis D17 (Fig. 2) were also influenced by the presence of mucilage. Degradation rates were exceptionally fast with the majority of the PHE degrading in the first two days. Mucilage initially slowed the degradation, but later resulted in a greater total amount of mineralization. Strain D17 can use purified

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α-Proteobacteria

β-Proteobacteria

γ-Proteobacteria

40%

60%

Firmicutes

Bacteroidetes

(Control)SHS+R (Control)SHS (Control)HS+R (Control)HS (RS)SHS+C3+R (RS)SHS+C2+R (RS)SHS+C1+R (RS)SHS+C3 (RS)SHS+C2 (RS)SHS+C1 (BS)SHS+C3+R (BS)SHS+C2+R (BS)SHS+C1+R (BS)SHS+C3 (BS)SHS+C2 (BS)SHS+C1 0%

20%

80%

Percentage of phylogenetic groups of bacterial communities

Fig. 5 – Phylogenetic groups of bacterial communities in each treatment. BS: bulk soil; RS: rhizosphere soil.

Fig. 6 – Band peak intensity (Int) of DGGE profiles from different cowpea line treatments. (a) band peak intensity of treatments (RS) SHS + C1, (RS) SHS + C2 and (RS) SHS + C3; (b) band peak intensity of treatments (RS) SHS + C1 + R, (RS) SHS + C2 + R and (RS) SHS + C3 + R.

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Table 7 – Pearson's correlations between PHE dissipation rate (PHE%) and biology parameters according to plant growth forms and DGGE bacterial diversity indices. Pearson's correlations Botany parameter

Microbiology parameter

Parameter

Root length Nodule number Nodule weight Root fresh biomass Root dry biomass Shoot height Leaf area Chlorophyll content of leaf Shoot fresh biomass Shoot dry biomass Rhizosphere Shannon index Rhizosphere richness Rhizosphere evenness Bulk soil Shannon index Bulk soil richness Bulk soil evenness

PHE% R

P

0.890 ⁎ 0.915 ⁎ 0.871 ⁎ 0.933 ⁎⁎ 0.943 ⁎⁎ 0.891 ⁎ 0.589 0.856 ⁎

0.018 0.011 0.024 0.007 0.005 0.017 0.219 0.030

0.893 ⁎ 0.819 ⁎ 0.915 ⁎

0.017 0.046 0.011

0.946 ⁎ 0.117 0.423 0.320 0.254

0.004 0.825 0.403 0.536 0.627

R: Pearson's coefficient of correlation, P: significance (2-tailed). P < 0.05. ** P < 0.01. *

mucilage for carbon source (data not shown), which may explain why the mineralization rate of treatment D17 + RM was lower than the treatment D17 during the first 6 days. Mucilage therefore, has the potential to both enhance the growth of degrading populations and stimulate PHE degradation. It can be speculated that the mechanisms of root mucilage that create the increase in PHE degradation may be due to two reasons. First, cowpea C1 secretes complex root mucilage that may provide a significant carbon source for microbes that colonize in the rhizosphere. Knee et al. (2001) pointed out that purified pea mucilage was used as the sole carbon source for the growth of several pea rhizospheric bacteria, including R. leguminosarum, B. cepacia and Pseudomonas fluorescens. Many Burkholderia and Pseudomonas spp. can degrade PAHs (Huang et al., 2008; Juhasz et al., 1997; Pathak et al., 2009), which may explain why cowpea C1 promoted a higher degradation rate than the other C2 and C3 breeding lines. Second, root mucilage may increase PAH bioavailability by serving as a biosurfactant. In our previous test, the cowpea's mucilage can lower the water surface tension to 48 mN/m at critical micelle concentration of 1 mg/mL (data not shown), which means cowpea's root mucilage may play a biosurfactant function to help increase bioavailability of PAH. It is a promising field for the future research. Plant root mucilages contain surfactants that may alter the interaction of bioavailability of hydrophobic substances, formation of water films, and the rates of microbial growth. Read and Gregory (1997) measured the surface tension of both maize and lupin mucilage was reduced to 48 mN/m at total solute concentrations of 0.7 mg/mL, which indicated the presence of surfactant. It has been suggested that polar glycolipids are synthesized as intermediates during the production of mucilage at the root tip (Green and Northcote, 1979). Phosphatidylcholine (lecithin), which is chemically similar to the phospholipid surfactants was identified in maize, lupin and

wheat root mucilage (Read et al., 2003). These glycolipids and other phospholipids associated with plant cell membranes would be expected to show marked surface activity (Bordoloi and Konwar, 2009). However, the effect of root mucilage simply as a surfactant has not been considered. It is possible that other root-produced substances with surfactant properties may also affect the PAH desorption. In this way, it will be easier for the microorganisms to access PAH and speed-up the rate of PAH mineralization. Plants induce changes in microbial structure and activity in the rhizosphere (Cébron et al., 2011). Since mucilage affects soil properties in the rhizosphere and thereby modifies the environment in which soil microbes' function, there may well be effects on microbial composition and activity. Based on the DGGE data (Fig. 3), the PHE degrading bacterial community that was enriched by root mucilage in the C1 treatment, had greater species Richness and Shannon index than soils from the C2 and C3 plant treatments where the plants do not produce abundant root mucilage. Root mucilage production thus appeared to promote bacterial growth in the rhizosphere (Knee et al., 2001) and to selectively enrich PHE degraders. Identifications of the predominant bacteria were accomplished by analyses of the DNA sequences cut from individual bands in the DGGE gels. Although, not examined for the individual species identified here, prior research has shown that isolates of these same speciescan degrade PAHs. These include species identified from the DGGE results such as P. putida, Labrys monachus, Methylobacterium sp., Spingomonas sp., Xanthomonas sp., Bacteroidetes, Filobacilus sp., Serratia marcescens, Ochrobactrum sp., Azospirillum sp., G. pallidus, and Stenotrophomonas rhizophila (Viñas et al., 2005; Schröder and Collins, 2011; Tao et al., 2007; Park and Crowley, 2006; Yuliani et al., 2012; Yang et al., 1994). Some of these species are also known to produce biosurfactants, including P. putida, G. pallidus, and Azospirillum sp. (Kruijt et al., 2009; Bonilla et al., 2005). Some of these species also have known plant growth promotion functions, including P. putida, Methylovorus sp., Xanthomonas sp., G. pallidus, S. marcescens and Azospirillum lipoferum (Bhattacharyya and Jha, 2011; Malik et al., 1997; Tsavkelova et al., 2006; Gravel et al., 2007). The DGGE results (Fig. 4) indicated that besides the PHE degrading bacteria, the PHE degrading consortia also consisted of mixtures of bacterial species that were not able to degrade PHE, but that might have other functions to benefit bioremediation of PHE, such as biosurfactant production, plant growth promotion (PGPR), and nitrogen fixation. While this study focused on culturable PHE degraders, the results suggest that PHE degradation in the rhizosphere commonly involves various species of bacteria by encouraging healthy plant growth and thus enhancing microbial activity to achieve PHE degradation. We developed a legume–rhizobia symbiosis model with cowpea plants and B. yuanmingense R3 for PHE phytoremediation. R3 cannot degrade PHE directly. However, prior research has shown that rhizobial microorganisms can stimulate the degradation of PAHs, and increase the population densities of PAH degraders in the plant rhizosphere (Teng et al., 2011; Johnson et al., 2005). Among numerous rhizobial microorganisms, several bacterial species in the genus of Rhizobium, Sinorhizobium, Mesorhizobium and Agrobacterium are able to grow on low molecular weight PAH as a carbon source (Yessica et al., 2013; Keum et al., 2005; Golubev et al., 2008; Jiménez et al., 2011; Sun et al., 2010). Similar results were also found in our DGGE data,

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Mesorhizobium sp. (band 14) was detected as PHE degrading bacteria in the rhizospheric soil of treatments C1 + R, C2 + R, and C3 + R. The presence of nodules can also stimulate the selective enrichment of organic hydrocarbon degrader organisms on the nodule surfaces. Dashti et al. (2009) found that the surfaces of root nodules on broad beans (Vicia faba) and lupine (Lupinus albus) were colonized with bacterial consortia that utilized oil and fixed nitrogen simultaneously. The inoculation of R3 not only improved plant growth, and increased PHE degradation rate in planted soils, but also changed the composition of PAHdegrading bacteria community. The role of plants in degrading organic contaminants during the bioremediation process is dependent on plant species, in particular, its root system and root exudates. In our study, botany parameters, especially the root biomass indicated strong significant correlations with PHE dissipation. Large root biomass accumulation will increase nutrient uptake in order to maintain plant health, to overcome stress and led to more effective PHE remediation. Corgie et al. (2004) found that PHE degradation declined with more distance from plant roots. Our Pearson correlation data also proved that microbial communities in rhizosphere contribute more to the PHE dissipation than microbial communities in bulk soil. A previous study reported that the root exudates and PAH degrader quantity were important factors that influenced PAH dissipation (Yi and Crowley, 2007). Since the root mucilage production cannot be measured during the process of plant experiments, we cannot analyze the correlations between root mucilage production and PHE dissipation. However, according to the Pearson's correlation analyses, we can conclude that the PHE phytoremediation is a complex process. The botany parameters (such as plant species, root exudates) and microbiology parameters (diversities of microbial community) will combine to influence the PHE dissipation in situ. To our knowledge, there have been few reports on the possible function of root mucilage on organic contaminant degradation, and papers about leguminous plants' root mucilage are scarce. Therefore, more research should be done about the function of root mucilage on PAH bioremediation. Root mucilage may benefit PAH rhizoremediation by (1) providing carbon source and energy for PAH degraders, (2) increasing PAH bioavailability by serving as a biosurfactant, and (3) stimulating growth of PGPR and biosurfactant producing bacteria to achieve PAH degradation together with PAH degraders.

4. Conclusions In conclusion, all three cowpea lines displayed a positive effect on PHE dissipation. Cowpea C1 with high root mucilage production was the most efficient plant in PHE phytoremediation by revealing higher biomass accumulation, higher Shannon index and higher PHE degradation rate than cowpea C2 and C3. Plant biomass and diversity of rhizosphere PHE degrading microbial showed a positive correlation with PHE dissipation, indicating that many important factors contribute together to influence PHE dissipation. Root mucilage enhanced 14C-PHE mineralization significantly both in soil slurry treatment and pure D17 strain treatment. The exact mechanisms involved in this process by how root mucilage enhances PHE remediation are not revealed,

57

and further work is required to further elucidate the processes involved. This study indicated that root mucilage originated from plant roots may play an important role in enhancing PHE degradation in soil and variation in root mucilage production may be a useful criterion in selecting plants for phytoremediation of contaminated soils.

Acknowledgments This work was supported by the National High Technology Research and Development Program (863) of China (No. 2012AA100402) and Cheung Kong Scholars Programme and the National Natural Science Foundation of China (Nos. 31125007 and 31370142). The authors are grateful to Dr. Philip Roberts and Dr. Wellington Muchero for providing seeds of the cowpea breeding lines; Dr. Milko Jorquera for conducting the DGGE analyses and Dr. Ron Mitchell for his review of the manuscript.

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