Introduction of Faba bean in crop rotation: Impacts on soil chemical and biological characteristics

Introduction of Faba bean in crop rotation: Impacts on soil chemical and biological characteristics

Applied Soil Ecology 120 (2017) 219–228 Contents lists available at ScienceDirect Applied Soil Ecology journal homepage: www.elsevier.com/locate/aps...

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Applied Soil Ecology 120 (2017) 219–228

Contents lists available at ScienceDirect

Applied Soil Ecology journal homepage: www.elsevier.com/locate/apsoil

Introduction of Faba bean in crop rotation: Impacts on soil chemical and biological characteristics

MARK



Amira Aschia,b, , Michaël Aubertb, Wassila Riah-Angleta, Sylvie Nélieuc,d, Caroline Duboisa, Marthe Akpa-Vinceslasb, Isabelle Trinsoutrot-Gattina UniLaSalle – Campus Rouen, Agri’Terr Unit, CS 40118, F-76134 Mont-Saint-Aignan, France Normandie Univ, UNIROUEN, IRSTEA, ECODIV, 76000 Rouen, France c UMR ECOSYS, INRA, Agro Paris Tech, Université Paris-Saclay, 78026, Versailles, France d Plateforme Biochem-Env, UMR ECOSYS, INRA, AgroParisTech, Université Paris-Saclay, 78026, Versailles, France a

b

A R T I C L E I N F O

A B S T R A C T

Keywords: Faba bean Enzyme activities PLFA Microbial biomass Active communities

Agricultural practices such as crop rotation affect soil physical, chemical and biological properties. Legumes crop effect has been shown to provide several agro-ecological services as a cereal previous crop. The aim of the present field study was to estimate the middle term effects of introducing faba bean in crop rotation on the structure and function of soil microbial communities. Two experimental rotation systems were tested (i) WheatBeet-Faba Bean-Rape-Wheat (Leg+) and (ii) Wheat-Flax-Wheat-Beet-Wheat (Leg−). Soil samples were collected on tilled plots at 0–10 cm depth on July 2013 under wheat. Soil microbial biomass and soil enzymatic activities (β-glucosidase, cellulase, urease and arylamidase activities) were assessed. Soil microbial diversity was evaluated with two complementary approaches: Phospholipid fatty acid profiling (PLFA) and the metabolic capabilities of the microbial community (Biolog Ecoplates). Soil organic carbon and total nitrogen were significantly and respectively 1.5 and 1.3 times higher in faba bean's rotation compared to free faba bean rotation. Soil microbial biomass did not differ significantly between the two rotations. In general, Leg+ rotation resulted in the greatest carbon mineralization and β-glucosidase and arylamidase activities. The analysis of Biolog data and PLFA profiling indicated that the rotation including faba bean has modified microbial populations and induced differences in the catabolic capability of soil microbial communities. Our results suggested that changing rotation crop by introducing faba bean two years before wheat modifies the surrounding habitat of microbial communities by providing available carbon and nitrogen as well as suitable soil pH. This new habitat could impact the structure of microbial communities and their functions. Leg + rotation seems to be a suitable practice promoting microbial activities in agricultural plots.

1. Introduction In Europe, numerous innovative practices have emerged to reduce the impact of agriculture on climate and environment changes with a focus of scientists on the way crop rotation can be designed. Indeed, a well designed crop rotation can contribute to weed control and decrease in diseases and pest attacks (Bagayonko et al., 1992) leading to a reduction in the use of pesticides. Furthermore, plant species present in crop rotation influence the soil water holding capacity by reducing soil erosion (Kollas et al., 2015). Diversification of crop along rotations plays an important role in the amount and quality of organic matter entering the soil (Raphael et al., 2016). According to the species identity, the mineralization of preceding crop residues can release important quantity of nutrients, which maintain soil fertility for the



following one and create suitable habitats for soil biota (Askegaard and Eriksen, 2007; Sauvadet et al., 2016). In this context, embedding grain legume in crop rotation has been shown to provide multiple environmental, agricultural and economical benefits. In fact, these plants have the ability to symbiotically fix atmospheric N2 through their association with Rhizobium bacteria, leading to potential decrease in the use of inorganic N amendments and thus reduce fossil energy consumption for plant production (Hardarson et al., 1991; Lopez-Bellido et al., 2006; Turpin et al., 2002). Furthermore, some legumes have a greater ability to mobilize P from less labile P forms than cereals (Kamh et al., 1999; Nuruzzaman et al., 2005a, 2005b). Finally, legumes act as an important source of protein for human and livestock feed and improve gross margins of full crop rotation (Jensen et al., 2010; Khan et al., 2010; Kopke and Nemecek, 2010; Preissel et al., 2015). In the other hand,

Corresponding author at: UniLasalle-Campus Rouen, Agri'Terr Unit, 3 rue du tronquet, 76130 Mont-Saint-Aignan, France. E-mail address: [email protected] (A. Aschi).

http://dx.doi.org/10.1016/j.apsoil.2017.08.003 Received 1 November 2016; Received in revised form 3 August 2017; Accepted 5 August 2017 0929-1393/ © 2017 Elsevier B.V. All rights reserved.

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Wheat-Beet-Wheat (Leg−), (ii) innovative rotation including legumes: Wheat-Beet-Faba Bean-Rape-Wheat (Leg+). Sampling was performed in the final wheat crop of each rotation, corresponding to spring wheat. The spring wheat plots had been tilled in March and composite sample was collected in July, in the middle of the growth season, by combining five soil cores (0–10 cm) under spring wheat cover in order to homogenize rhizospheric properties.

rotation including legumes at least for one time, can influence soil microbial communities directly (Bunemann et al., 2004; Voisin and Gastal, 2015) and indirectly through their effects on the quantity, the quality, and the distribution of soil organic matter in upper-soil horizons. Such systems tend to have higher microbial biomass and activities (Moore et al., 2000). Melero et al. (2012) also showed a greater amount of soil organic carbon, total N and β-glycosidase activity in legumes-wheat rotation under no tillage system than in continuous wheat and fallow-wheat rotation. Lupwayi et al. (1999) demonstrated that wheat plots preceded by legume crops had a higher microbial biomass and a lower qCO2 than wheat proceeded by summer fallow. Moreover, the presence of legumes in crop rotation, through its implication in nitrogen fixation, promote the activity of nitrogenase enzyme and the release of hydrogen gas (H2), which induces the multiplication of some specific microorganisms in soil (microorganisms which use H2 as energy sources) (Dong et al., 2003). Nevertheless, if the beneficial effects of introducing legumes as wheat’s previous crop have been clearly highlighted, a serious knowledge gap remains concerning its medium-term effects on soil characteristics (biotic and abiotic) during crop rotation i.e. are the benefits of legumes likely to be sustained beyondone crop? This lack of knowledge limits the agricultural community to diversify the technical solutions, such as identifying a panel of possibilities to built crop rotations around the presence of legumes, for the sustainable management of soil fertility. The present study was thus conducted to examine the persistence of faba bean impacts on wheat cultivation when faba bean was grown two years before wheat. We hypothesized that the crop rotation including faba bean could affect soil physicochemical properties and microbial communities by changing the diversity of these communities and their enzymes activities. To reach these objectives, we used a single-site agricultural assay designed for the evaluation of agronomic and ecological effects of faba bean position along crop rotation. It consisted in replicate comparisons of rotations with and without faba bean placed two years before a wheat culture within which soil was sampled for chemical analysis evaluation microbial communities diversity and soil functioning. We assessed both total organic carbon and nitrogen and the amount of active carbon. Microbial biomass was used to estimate the abundance of soil microorganisms and enquire us about modifications due to soil management. Molecular approach, using RNA and DNA co-extraction and real-time PCR, provided information about active bacterial and fungal community under the two studied rotation. Phospholipids fatty acid (PLFA) profiling were chosen to estimate the relative diversity of microbial community in sampled soil. Functions of microbial community were evaluated by enzymes measurement and soil carbon mineralization. The activities of enzymes involved in C and N cycle were studied, some of which have been previously shown to rapidly respond to soil perturbations and modifications of agricultural practice (Lebrun et al., 2012). Finally, the community-level physiological profile was employed to estimate bacterial functional diversity.

2.2. Measurement of soil properties Topsoil samples were air-dried and sieved at 2 mm for soil chemical and physical analyses. Soil pH in H2O and in 1 M KCl was measured with a glass electrode in 1:2.5 suspension (NF ISO 10390, 2005), ΔpH (ΔpH = pHH2O − pHKCl) was determined. For a given soil type, this index is positively correlated with base saturation (Baize, 2000). The cation exchange capacity (CEC) was determined using cobalt hexamine trichloride according to NF ISO 23470 (2007). Moisture content was recorded after drying at 105 °C for 48 h. Five grams of air dried soil were used to determine soil textural. Soil samples were firstly treated by HCl solution to remove soil carbonate then H2O2 33% was added to eliminate soil organic matter and finally hexametaphosphate solution was used to break up the soil particles. References concentration was done according to NFX-31-107. Particle size distributions (%) were measured by laser diffraction (Malvern Mastersizer 2000, UK) according to manufacture protocol. Total carbon (TC) and total nitrogen (TN) were quantified by a dry combustion method using an automatic analyzer (Flash2000 Thermo Scientific). The Organic Carbon (OC) was measured with TOC-Analyzer (Shimadzu TOC- SSM 5000). Permanganate Oxidizable Carbon (eq. Active carbon, POXC) was extracted from 2.5 g of air dried soil. Quantity of carbon oxidized by MnO4 was measured as described by Culman et al. (2012) using spectrophotometer (Varian Cary 50 Scan) at 550 nm. 2.3. Structure of microbial community 2.3.1. Microbial biomass carbon The microbial biomass carbon (MBC) in soil was determined by the chloroform fumigation-extraction method described by Vance et al. (1987) with some modifications. Briefly, sixty grams of field-moist soil were separated into two beakers. The first one was fumigated with ethanol-free chloroform for 24 h in dark conditions and the second was used as a control (same conditions without CHCl3 fumigation). The extraction of Soluble Organic Carbon was performed from fumigated and non-fumigated soil samples with K2SO4 (0.5 M). In the extracts, carbon content was measured with a total organic C analyzer (Shimadzu TOC-VCSH). MBC was calculated as the difference in C content in fumigated and non-fumigated sample without using a correction coefficient. 2.3.2. DNA/RNA co-extraction and quantitative reverse transcription qPCR RNA and DNA were co-extracted from 2 g soil using the Total RNA isolation Kit® and the DNA Elution Accessory Kit® for Soil, respectively (MO BIO-USA). Extracts were stored at −80 °C for RNA and −20 °C for DNA until use. To remove contaminating DNA from RNA preparation, TURBO DNA-free™ kit (Applied Biosystems, France) was used and RNA extracts were further purified with the RNeasy® MinElute Cleanup kit (Qiagen Gmbh, Germany) according to the manufacturer’s protocol using silica spin columns. Purified nucleic acid extracts were eluted in a final volume of 15 μL with DEPC-treated water and stored at −80 °C. The quality of RNA was assessed by Experion™ RNA StdSens Analysis Kit (BIO-RAD) according to manufacturer‘s recommendation (LarocheAjzenberg et al., 2012). Purified dsDNA and RNA were quantified by fluorimetry. dsDNA quantification was operated by using the Fluorescent DNA quantitation Kit (Hoechst 33258, Biorad) and dsDNA extracts were stored at −20 °C (Gangneux et al., 2011). RNA was quantified using Quant-iT™

2. Materials and methods 2.1. Site, rotations and soil sampling The study site was located in north western France and belongs to the Institute of Plant Arvalis in Rots (Calvados, Normandy) (N49°12′05.74”, W 0°27′13.08”). The mean annual precipitation is 712 mm spread over 7 months from October to April and the mean annual temperature is 10.5 °C. The region has a maritime climate with low thermal amplitude during the year. The experimental site had a silt loam texture and was chosen to evaluate agronomic and ecological effects of the position of faba bean in crop rotation. The soil was classified as a Calcisol (FAO classification). The experimental design consisted in 8 plots (12 × 90 m) of two rotations with four replicates. Two rotation systems were examined with different position of leguminous in crop succession: (i) common rotation in Normandy area: Wheat-Flax220

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Table 1 Enzymes and substrates used for enzyme activity assays. Recommended name

Abbreviations

EC number

Assays conditions

References

Substrate

Optimum pH

N cycle enzymatic activities Arylamidase ARYL N Urease URE

3.4.11.2 3.5.1.5

L-Leucine β-Naphthylamide (8 mM) Urea (0.1 M)

8.0 Soil pH

Acosta-Martinez and Tabatabai (2000) Kandeler and Gerber (1988)

C cycle enzymatic activities Cellulase CEL β-glucosidase GLU

3.2.1.4 3.2.1.21

4-NP-β-D-cellobioside (10 mM) p-NP-β-D-glucopyranoside (50 mM)

6.0 6.0

Trap et al. (2012) Trap et al. (2012)

RiboGreen® RNA Assay Kit, RNA extracts were stored at −80 °C. The cDNA was synthesized from purified RNA using High capacity cDNA reverse transcription kit (Kit Applied Biosystems)(Laroche-Ajzenberg et al., 2012). Fungal and bacterial communities were estimated on both dsDNA and cDNA by qPCR using universal primer sets designed for each type of microorganism. For total and active fungal communities, 18S rDNA and 18S rRNA gene qPCR amplifications were carried out in a total volume of 25 μL with 1 x qPCR Master Mix (SYBR Green I, Applied Bioystem), 0.5 μM of FU18S1 and Nu-SSU-1536 primers (Borneman and Hartin, 2000), 0.5 mg mL−1 of BSA (NEB) and 10 ng of total DNA and or cDNA. Standard curves were obtained using serial dilutions of linearized plasmids containing cloned Fusarium graminearum 18S rRNA genes (assuming that the F. graminearum genome is representative of soil fungi genomes). After an initial denaturation and enzyme-activation step of 15 min at 95 °C, 40 cycles of PCR were performed in Light Cycler 480 as follows: 40 s at 95 °C, 45 s at 64 °C and 30 s at 72 °C. The qPCR efficiency ranged from 93% to 98%. 16S rDNA and 16S rRNA amplifications were carried out in the same conditions as 18S rDNA and rRNA PCR except for primers [63f 5′CAGGCCTAACACATGCAAGTC-3′ (Marchesi et al., 1998) and BU16S4 5′-CTGCTGCCTCCCGTAGG-3′ derived from 341F (Muyzer et al., 1993)] and amplification protocol (40 s at 95 °C, 45 s at 64 °C and 30 s at 72 °C). The 314 bp 16S PCR product exceeds the recommendations for SYBR_Green qPCR analysis; however the efficiency of the qPCR ranged from 98% to 102%.

split-less mode (1 μL, injector temperature 250 °C) equipped with a BPX70 column (60 m, 0.25 mm i.d., 0.25 mm df., SGE), and helium as a carrier gas. The oven temperature was programmed at 80 °C for 1 min, followed by a first ramp at 25 °C/min to 160 °C with a 3 min hold, then a second ramp at 3° C/min to 180 °C with a 9 min hold, and a final ramp at 2° C/min to 220 °C, which was held for 7 min. Fatty acids were commonly denoted as the total number of C atoms, the number of double bonds, followed by the position of the double bond from the methyl end of the molecule. The FAME identification and quantification were performed using as standards: 37 component FAME mix from Supelco, methyl nonadecanoate (19:0, used as internal standard) from Fluka and Br1 Mix, methyl-13-methyl tetradecanoate (i15:0), methyl 15-methyl hexadecanoate (i17:0), methyl vaccinate (18:1w7c), methyl cis-9,10-methyleneoctadecanoate (cy19:0) from Larodan, Solna, Sweden. The PLFAs i14:0, i15:0, a15:0, i16:0, i17:0 and a17:0 were chosen to represent Gram-positive bacteria while 16:1ω7c, 17:1ω7cand cy19:0 have been used as an indicator of Gram-negative bacteria (Wu et al., 2009; Bowles et al., 2014). The PLFAs 18:1ω9c and 18:2ω6c were designated as representatives of fungi (Acosta-Martinez et al., 2007; Shahzad et al., 2012). The following ratios of fatty acid relative abundance were also calculated: fungi to total bacteria, Gram positive bacteria to Gram negative bacteria, iso/anteiso [(i15:0 + i17:0)/(a15:0 + a17:0)]; saturated/monounsaturated [(14:0 + 15:0 + 16:0 + 18:0 + 20:0)/(16:1ω7c + 18:1ω9c + 18:1ω7c + 20:1ω9c)]. These ratios have been used as an index of environmental stress (Mckinley et al., 2005; Petersen and Klug, 1994). 2.4. Functions of microbial communities

2.3.3. Phospholipids fatty acid analysis (PLFA) Fresh soil samples were homogenized, sieved at 2 mm and frozen at −80 °C until PLFA analysis. After freeze-drying, PLFA were extracted using a modified method of Bligh and Dyer (1959) (Frostegard et al., 1991; Shahzad et al., 2012). Fatty acids were extracted from 2 g of soil by a single-phase mixture of chloroform-methanol-pH 4 citrate buffer (1:2:0.8, v/v/v) shaken at 300 rpm for 1.5 h and centrifuged 15 min at 1500 rpm. The supernatant was retained and the soil was re-extracted as before. Phase splitting in the combined supernatants was obtained by adding citrate buffer and chloroform (overnight separation). The CHCl3 layer was dried under N2 at ambient temperature, re-dissolved with chloroform and purified on silica cartridges (chromabond® 3 mL/ 500 mg SiOH). After discarding chloroform and acetone rinses, the methanolic fraction containing phospholipids was evaporated under N2. Before analysis by gas chromatography-mass spectrometry (GC–MS), PLFA had to be transformed into their less polar fatty acid methyl ester (FAME) derivatives. Instead of the usual method by alkaline methanolysis using KOH in methanol/toluene (Dowling et al., 1986), the derivatization has been performed on-line in GC injector by tri methyl sulfonium hydroxide (TMSH) (Gomez-Brandon et al., 2008). The extract was dissolved in 500 μL methyl tert-butyl ether. A 130 μL fraction was inserted in an injection vial with 50 μL TMSH and allowed to react 30 min at ambient temperature. 20 μL of a 1 μg/mL solution of methyl nonadecanoate (FAME 19:0) was then added as an internal standard. FAMEs were analyzed with GC/MS (4000 GC/MS, Varian) in

2.4.1. Enzyme activities The choice of enzymes in this study was based on their involvement in biogeochemical cycles especially in carbon and nitrogen cycle. The activities of four enzymes were measured using methods described in Table 1. Cellulase, β-glucosidase, arylamidase and urease were examined on < 2 mm field-moist sample at their optimal pH values. Microbial activity analyses were performed in 4 composite soil samples per rotation (obtained after pooling). 2.4.2. Soil carbon mineralization To estimate Carbon mineralization, 20 mL of NaOH (1 M) was placed in 1L hermetic reception with thirty grams of fresh shaved-soil and left for 72 h. The amount of CO2eC produced during incubation was trapped in NaOH and determined by measuring conductivity with pH meter (METTLER TOLEDO Seven Multi™). 2.5. Potential metabolic diversity Biolog Ecoplates 96-well (Biolog Inc., USA) were used to assess the carbon source utilization patterns of the soil bacterial communities (Garland and Mills, 1991). Biolog Ecoplates were inoculated according to Calbrix et al. (2005): 5 g of fresh Leg+ and Control soils were resuspended by a mechanical shaking process in 45 mL of a sterile 0.85% NaCl solution. This suspension was centrifuged at 1000 rpm for 5 min. 221

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The supernatant was then diluted in sterile NaCl solution (0.85%) and placed (150 μL) in each of the 96 wells in the Biolog Ecoplates in order to reach a cell density of 1500 cultivable bacteria per well. Aliquots of the supernatant were spread on R2A plates to check that every Biolog Ecoplate had been inoculated with the same bacterial density. The plates were incubated at 20 °C, and the optical density at 590 nm (OD590) was measured (Emax reader, Molecular Device, Sunnyvale, CA) at 0, 24 and 48 h. The profiles were observed after 48 h for Ecoplates incubations. Optical density readings were corrected by subtraction of the blank control value and negative values were set to zero. Substrate richness (positive well numbers) and the average well color development AWCD (calculated as the average optical density across all wells per plate) were estimated and used as potential metabolic diversity of microbial communities. For the Biolog Ecoplates analysis, a well is considered as positive when the OD590nmi − OD590control > 0.25. The functional diversity from the Biolog Ecoplates was evaluated by enumeration of the positive wells and by calculating the Average Well Color Development (AWCD) of all of the carbon sources for each sample. The AWCD was calculated as follows: AWDC = x(ODi − ODA1)/95 with the ODi = OD590nm in each well and the ODA1 = OD590nm in the control well A1.

Table 3 Structure of soil microbial communities in the two studied treatments: Leg+ (New rotation including faba bean two years before sampling date) and Leg− (Classic rotation without leguminous during 5 years). Different letters in the same line indicate significant differences (n = 4, p < 0.05). Treatments Structure of microbial communities

Leg+

Leg−

Total dsDNA (μg g−1 soilDW) Microbial biomass carbon −1 (mg C kg−1 soilC·kgsoilDW) Total microbial PLFA (nmol g−1 soilDW)

5.69 ± 0.26a 326.19 ± 43a

4.11 ± 1.71a 305.94 ± 28a

15.80 ± 3.14a

16.09 ± 3.80a

2.54 ± 0.10a

1.00 ± 0.40b

10.69 ± 1.24a

8.56 ± 5.43a

4.34 ± 7.19a

2.49 ± 0.49a

1.40 ± 0.43a

1.37 ± 0.29a

14.47 ± 3.00a 0.76 ± 0.17a 0.053 ± 0.007a 1.38 ± 0.12b 0.99 ± 0.04a 2.19 ± 0.19b

14.87 ± 3.57a 0.74 ± 0.20a 0.049 ± 0.002a 1.63 ± 0.08a 0.96 ± 0.03a 2.62 ± 0.11a

Total communities Bacterial dsDNA (copy number of 16S 10 genes g−1 soilDW)(×10 ) Fungal dsDNA (copy number of 18S 7 genes g−1 soilDW)(×10 ) Active communities Bacterial cDNA (copy number of 16S 12 genes g−1 soilDW)(×10 ) Fungal cDNA (copy number of 18S 10 genes·g−1 soilDW)(×10 ) Total bacterial PLFA (nmol g−1 soil) Total fungal PLFA (nmol g−1 soil) Fungal to bacterial ratio Gram(+) to Gram(−) ratio Iso to anteiso ratio Sat to monounsat ratio

2.6. Statistical analysis Normality and homogeneity of variances were checked according to the Wilk-Shapiro test and the Bartlett test, respectively. Data of soil physicochemical properties and microbial community structure were analyzed through standard analyses of variance (ANOVA). The differences between the studied rotations were considered significant for a value of p < 0.05. In addition, to assess the effects of faba bean position in the crop rotation on potential microbial activities, principal component Analysis (PCA) were performed with the results obtained from the degradation of Biolog's substrates. Coordinates of each point were extracted and tested to highlight significant differences between the two studied rotations. Furthermore, spearman correlation coefficients were calculated between soil physicochemical proprieties and structural and functional parameters of soil microbial communities. All the statistical tests were carried out using R software (R Development Core Team, 2009).

dry soil) than in the rotation including faba bean (12.10 ± 0.69 gkg−1 air dry soil). Furthermore, active carbon (POXC) and total carbon content (TC) were respectively 1.09 and 1.5 times lower in control (Leg−). The wheat of Leg+ rotation exhibited 1.3 times higher of total N content. There was also a significant effect of Leg+ rotation on soil pHwater and pHKCl. Leg+ rotation presented higher pH values in comparison to control rotation (Leg−). ΔpH was statistically different between the two rotations (0.63 ± 0.04 for Leg+ and 0.94 ± 0.02 for Leg− rotation). The CEC exhibited the same trend than pH and was higher in Leg+ rotation.

3.2. Structure of microbial communities in the two rotation system 3. Results

3.2.1. Abundance The quantification of microbial biomass carbon measured by fumigation-extraction method and the total DNA extracted from soil did not differ statistically between the final wheat of the two rotations Leg− and Leg+ (Table 3). Indeed, the two rotations exhibited on average 316 mg C per kg of dry soil for microbial biomass carbon and 4.47 μg g−1 of dry soil for total DNA, respectively. Furthermore, the quantification of the total bacterial and fungal communities by qPCR analysis showed that the microbial community was dominated by bacteria in both rotations. Bacterial biomass was significantly higher in Leg+ (2.54 × 1010 ± 0.1 × 1010 copy of 16S rDNA genes g−1) than in Leg− rotation (1 × 1010 ± 0.6 × 1010 copy of 16S rDNA genes g−1) and was positively correlated to soil organic carbon (r = 0.71, p < 0.05), total nitrogen content (r = 0.74, p < 0.05) and soil pH (r = 0.76, p < 0.05) (Table 4). However, fungal DNA did not differ statistically between the two studied rotations and was not correlated to any soil physicochemical parameters measured (Table 4). The quantification of active bacterial and fungal communities was assessed by the measurement of cDNA (Table 3). The results showed that the abundance of active communities were higher than those of total communities. Active bacterial biomass was higher than active fungal biomass but the two communities did not statistically differ between studied rotations and they did not exhibit any correlations with measured soil properties. Concerning the quantification of total PLFA (Table 3), the final

3.1. Soil characteristics The rotation including faba bean two years before wheat showed greater values of soil chemical properties (Table 2). All the tested parameters were systematically higher in the final wheat of Leg+ compared to the final wheat of Leg− rotation. The amount of organic carbon (OC) was lower in the Leg− rotation (8.11 ± 0.23 gkg−1air Table 2 Results of soil physicochemical properties in the two studied treatments: Leg+ (New rotation including faba bean two years before sampling date) and Leg− (Classic rotation without leguminous during 5 years). Different letters (a and b) in the same line indicate significant differences (n = 4, p < 0.05). Treatments

pHwater pHKCl ΔpH CEC (cmol+ kg−1) Total C (g kg−1 airdriedsoil) Total N (g kg−1 airdriedsoil) Organic C (g kg−1 airdriedsoil) POXC (mg kg−1 airdriedsoil)

Leg+

Leg−

7.80 ± 0.05a 7.17 ± 0.07a 0.63 ± 0.04b 18.91 ± 0.43a 14.45 ± 0.41a 1.46 ± 0.03a 12.10 ± 0.69a 632.57 ± 5.82a

6.10 ± 0.10b 5.16 ± 0.07b 0.94 ± 0.02a 11.72 ± 0.05b 10.11 ± 0.23b 1.09 ± 0.05b 8.11 ± 0.23b 576.73 ± 8.51b

222

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Table 4 Spearman’s correlations between physicochemical parameters and structural and functional parameters. Significant correlations are in bold ((> 0.65), n = 4), and significant adjusted p value are indicated as *: p < 0.05; **: p < 0.01; ***: p < 0.001; n.s.: p > 0.05. Total C

Organic C

Total DNA Total Microbial PLFA Microbial biomass carbon

0.08 −0.21 −0.04

n.s.

Total communities Bacterial dsDNA Fungal dsDNA

0.69 0.19

n.s.

Active communities Bacterial cDNA Fungal cDNA Total bacterial PLFA Total fungal PLFA Fungal to bacterial ratio Gram(+) to Gram(−) ratio Iso to anteiso ratio Sat to monounsat ratio

−0.24 0.07 −0.21 0.07 0.55 −0.67 0.26 −0.59

n.s.

C cycle enzyme β-glucosidase Cellulase

0.43 0.00

n.s.

N cycle enzyme Arylamidase Urease Mineralized carbon

0.76 0.21 0.83

*

n.s. n.s.

n.s.

n.s. n.s. n.s. n.s. n.s. n.s. n.s.

n.s.

n.s. *

Active C

0.31 −0.24 −0.17

n.s.

0.71 0.45

*

−0.28 0.26 −0.24 0.02 0.36 −0.67 0.45 −0.59

n.s.

0.36 −0.33

n.s.

0.62 0.45 0.88

n.s.

n.s. n.s.

n.s.

n.s. n.s. n.s. n.s. n.s. n.s. n.s.

n.s.

n.s. **

Total N

0.23 0.31 0.31

n.s.

0.69 0.17

n.s.

−0.47 −0.33 0.31 0.24 0.19 −0.95 0.09 −0.81

n.s.

0.78 0.14

*

0.86 0.48 0.67

**

wheat of Leg+ rotation showed values ranging from 11.71 to 19.31 nmol g−1soil while the final wheat of Leg− rotation exhibited values ranging from 11.16 to 20.10 nmol g−1 soil. The two rotations were not statistically different (Table 3). Finally, including faba bean in crop rotation did not have significant influence on the abundance of soil microbial communities except for bacterial communities (dsDNA) expressed as 16S genes copy number.

n.s. n.s.

n.s.

n.s. n.s. n.s. n.s. *** n.s. *

n.s.

n.s. n.s.

pH

0.18 0.00 0.14

n.s.

0.74 0.14

*

−0.67 −0.26 0.00 0.33 0.57 −0.81 0.00 −0.74

n.s.

0.71 0.04

*

0.90 0.24 0.69

**

n.s. n.s.

n.s.

n.s. n.s. n.s. n.s. * n.s. *

n.s.

n.s. n.s.

CEC

water

0.01 −0.07 0.17

n.s.

0.76 0.12

*

−0.23 0.12 −0.07 0.24 0.57 −0.74 0.09 −0.59

n.s.

0.55 0.21

n.s.

0.83 0.07 0.71

*

n.s. n.s.

n.s.

n.s. n.s. n.s. n.s. * n.s. n.s.

n.s.

n.s. *

0.32 0.03 0.55

n.s.

0.83 0.21

**

−0.54 −0.18 0.03 0.32 0.39 −0.85 −0.01 −0.85

n.s.

0.81 0.38

*

0.96 −0.01 0.61

***

n.s. n.s.

n.s.

n.s. n.s. n.s. n.s. ** n.s. **

n.s.

n.s. n.s.

Arylamidase activity was significantly 2.2 times higher in Leg+ rotation than in the control rotation. Urease activity was not significantly different between the two rotations. The activity of β-glucosidase and cellulase also responded differently to the presence of faba bean in crop rotation. The β-glucosidase activity seemed to be more sensitive to the presence of faba bean and was 1.3 times higher in Leg+ rotation than in Leg− rotation, whereas, the analysis of cellulase activity revealed no significant difference. Furthermore, some correlations were made in order to examine the relationships between enzyme activities and soil nitrogen and carbon contents. Arylamidase activity was positively related to the total nitrogen content (r = 0.90, p < 0.01), total carbon content (r = 0.76, p < 0.05), active carbon (r = 0.86, p < 0.01) and soil pH (r = 0.83, p < 0.05). The β-glucosidase activity was positively correlated to active carbon (r = 0.78, p < 0.05), total nitrogen content (r = 0.71, p < 0.05) and soil CEC (r = 0.81, p < 0.05). No correlations were showed between urease and cellulase activities and soil nitrogen and carbon contents (Table 4). The result of soil carbon mineralization rate expressed as the total C released per day per gram of organic carbon is presented in Fig. 2. This rate was significantly different between the two rotations. In addition, the cumulative amount of mineralized C was statistically higher in Leg+ rotation compared to control rotation. The mean value of this mineralized C was about 99.3 mg of CeCO2 kg−1 dry soil per day in Leg+ rotation and 29.7 mg of CeCO2 kg−1 dry soil per day in Leg− rotation (data not showed). The amount of carbon released was positively correlated to soil pH (r = 0.71, p < 0.05) and organic carbon content (r = 0.88, p < 0.01).

3.2.2. Diversity Microbial community structure estimated by PLFA markers is shown in Table 3. The analyses of variance performed on total bacterial and fungal PLFA biomarkers indicated a significant difference between these two groups in each rotation with higher concentration of bacterial PLFA (mean value 14.66 ± 3 nmol g−1 soilDW) than fungal PLFA (mean value 0.75 ± 0.17 nmol g−1 g−1 soilDW) (statistical test not shown). Although, there were no statistical differences in total bacteria and total fungi between Leg+ rotation and Leg− rotation. The ratio of Gram positive to Gram negative bacteria showed a significant difference between the two rotations. The Leg− rotation exhibited 1.1 times higher Gram (+) to Gram (−) ratio than in Leg+ rotation indicating that the soil collected from Leg+ rotation contained relatively more Gram-negative bacteria than Gram-positive bacteria comparing to control rotation (Leg−). This ratio was negatively correlated to active carbon, total nitrogen content, soil pH and soil CEC (Table 4). Moreover, the iso/anteiso ratio and saturated/monounsaturated ratio, which have been used as an index of environmental stress (abiotic, nutritional), did not showed the same trend between the two rotations. Saturated/ monounsaturated ratio exhibited higher values in Leg− rotation than in Leg+ rotation and they were negatively correlated to active carbon, total nitrogen and soil CEC. At the opposite, the iso/anteiso ratio did not differ statistically between rotations.

3.4. Potential metabolic diversity under faba bean rotation To reduce the dimensionality of the potential metabolic diversity data set, a PCA was performed to compare the two rotations (Fig. 3. A). The first axis (PC1) and the second axis (PC2) explained 38.66% and 21.62% of the total variability, respectively. PCA clearly illustrated how the rotation including faba bean had significantly influence the potential metabolic diversity of microbial community based on the second axis (PC 2, p < 0.05). The comparison of the potential metabolic diversity of the two

3.3. Functions of microbial community in the two rotation system 3.3.1. Enzyme activities and soil carbon mineralization Among the four tested enzymes, two were involved in nitrogen cycle (arylamidase and urease) and the two others were involved in carbon cycle (cellulase and β-glucosidase). All these enzymes did not respond in a similar way to the presence of faba bean in the rotation (Fig. 1). 223

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Fig. 1. Enzyme activities in the Leg+ (New rotation including faba bean two years before sampling date) and Leg− (Classic rotation without leguminous since 5 years) treatments. Different letters indicate significant differences between treatments (n = 4, p < 0.05).

leguminous. It mobilizes more soil phosphorus (Nuruzzaman et al., 2005), increases yields of succeeding crops and reduces problems caused by weeds and pathogens (Kopek and Nemecek, 2010). In this review, authors concluded that the biological nitrogen fixation can provide not only assimilated nitrogen to faba bean crop, but also to the whole crop rotation. Moreover, Pare et al. (1993) demonstrated that faba bean sown three years earlier increase corn dry matter and ear nitrogen accumulation compared to monoculture. Positive middle effect of faba bean on wheat crop was shown. This legume increased significantly the yield by 21% in the first and 12% in the second year compared to N fertilized continuous cereals (Wright, 1990). Then, these advantages make it one of the most interesting species for understanding the medium term effects of introducing legumes on microbial communities of agricultural soils. Including legumes in crop rotation seems to be suitable practices for maintaining soil microbial communities and farmers productivity Fig. 2. Carbon mineralization rate under the two rotation systems. Different letters indicate significant differences between treatments (n = 4, p < 0.05).

4.1. Leg+ rotation vs Leg− rotation 4.1.1. Effects on soil physicochemical proprieties In the present study, significant difference in the soil pH was detected between the final wheat of both rotations with and without legumes. Our result was probably due to the incorporation of faba bean residues to the soil two years before sampling date. Butterly et al. (2010) report that chemical properties of the residues can influence soil pH because of their alkalinity and N contents. Legumes can initially cause the acidification of soil but adding residues can offset this effect by the decomposition of organic anions and organic nitrogen (Yan et al., 1995). Cation Exchange Capacity (CEC) is an important soil property in describing nutrient availability for plant growth and it is closely linked to soil pH. The positive co-variation of the CEC and the pH in the agricultural soils is widely documented which join our results (Helling et al., 1964; Manrique et al., 1991; Bailey et al., 2008).

rotations based on the AWCD of the six groups of substrates is presented in Fig. 3B. The six groups included carbohydrates, carboxylic acids, amino acids, complex carbon sources, amines and carbon phosphate. Among them, only two groups of substrate showed significant differences between the two rotations with Leg+ presented higher AWCD for complex carbon sources, while the control rotation presented the higher AWCD for amino acids. 4. Discussion Faba bean (Vicia faba L.) is grown under a wide range of climatic conditions from temperate to subtropical regions. The introduction of this plant in crop rotation offers many advantages over other 224

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Fig. 3. Potential metabolic diversity in response to the presence of faba bean in rotation sequence, (A) Results of the principal component analyses (correlation circle) performed on the Biolog Ecoplate profiles from Leg+ (New rotation including faba bean two years before sampling date) and Leg− (Classic rotation without leguminous since 5 years) treatments after 48 h of incubation. The first two components explained 60% of the total variance. (B) Percent of total carbon source utilization response based on the AWCD (Average Well Color Development) in Leg+ and Leg− for the different guilds-amines (A), amino acids (AA), carboxylic acids (CA), complex carbon sources (CCS), carbon phosphate (CP) and carbohydrates (Carb). Different letters show significant differences within each substrates source (n = 4, p < 0.05). A1: Water/A2: β-Methyl-D-Glucoside/A3:DGalactonic Acid γ-Lactone/A4: L-Arginine/B1: Pyruvic acid methyl ester/B2: D-Xylose/B3: DGalacturonic acid/B4: L-Asparagine/C1: Tween 40/ C2: i-Erythritol/C3: 2-Hydroxy benzoic acid/C4: LPhenylalanine/D1: Tween 80/D2: D-Mannitol/D3: 4Hydroxy benzoic acid/D4: L-Serine/E1: αCyclodextrin/E2: N-Acetyl-D-Glucosamine/E3: γHydroxybutyric acid/E4: L-Threonine/F1: Glycogen/ F2: D-Glucosaminic acid/F3: Itaconic acid/F4: acid/G1: D-Cellobiose/G2: Glycyl-L-glutamic Glucose-1-Phosphate/G3: α-Ketobutyric acid/G4: Phenylethyl-amine/H1: α-D-Lactose/H2: D, L-αGlycerol Phosphate/H3: D-Malic acid/H4: Putrescine

Grandy et al., 2002), perhaps due to legume effects on microbial communities, the production of polysaccharides, and aggregate stabilization (Haynes and Beare, 1996; Grandy and Robertson, 2007). Thus, the differences in soil organic C between the two rotations may probably come from the cumulative effects of the disappearance of a culture generating low residue amounts (e.g. flax) by a culture generating high residue amounts (faba bean). Nitrogen is considered as the most limiting nutrient in many agricultural plant production systems. Many studies have further highlighted the influence of legume on nitrogen pools in the soils (Evans et al., 2001; Peoples et al., 2001; Olesen et al., 2009). In this study, soil with the rotation including faba bean showed a higher amount of total nitrogen. Faba bean is known for its positive soil N balance comparing to other legumes (Hauggaard-Nielsen et al., 2009). It has a greater efficiency of nitrogen fixation (Herridge et al., 2008), which allowed it to accumulate more nitrogen. Indeed, the introduction of some legumes in the crop rotation, such as vetch, resulted in higher amounts of soil organic matter and soil N concentration in comparison with continuous cereal cropping and cereal fallow rotation (Liu et al., 2006). Moreover, in our experiment a previous crop in Leg+ rotation was rape while in Leg− rotation was beet. The literature reported that total N was not affected when a previous crop is rape or beet because their residues bring similar amount of nitrogen to the soil (+ 20 uN/ha) (Barbot et al., 2004). Nevertheless, the amount of dry matter is more important in faba bean resulting in a higher C/N ratio than other legumes crop residues (chickpea, soya) (Hassan et al., 2012). This higher C/N ratio indicate a slower decomposition kinetics of this plant (Trinsoutrot et al., 2000), that could have an effect on both structure and functions of soil microbial communities.

Soil organic carbon is considered as a good indicator of soil health because of its influence on all aspects of soil fertility. In crop rotation, among factors that influence soil organic carbon, quality and quantity of plant residues restituted to the soil are of major concern (Berg and McClaugherty, 2008). Thus, our result shows that introducing faba bean in crop rotation enhances the amount of organic carbon. In fact, the difference in soil organic carbon between the start of the experiment (2009) and the date of sampling (2013) showed an increase of 1.06 g/ kg. The organic content of the original soil was quite low at the beginning of the experiment, and it may have been a sufficient return of crop residues to allow this increasing trend in soil organic matter in the 5 last years. Hernanz et al. (2009) demonstrated such a pronounced effect on organic carbon with an increase about 1.6 gkg−1 five years after the introduction of pea-vetch in crop rotation. In Hernanz et al. (2009) study, shifting from cereal-follow rotation to wheat-pea/vetch rotation had a similar effect on soil organic carbon as shifting from conventional tillage to no tillage. The active carbon content (POXC), which is considered as an available carbon fraction for microbes, was positively affected by the presence of faba bean incorporation. Similar findings were observed by Sainju et al. (2007) after six years of cerealpea rotation. Moreover, in our experiment, POXC was correlated to soil organic carbon, CEC, soil pH and total nitrogen as previously observed by Tatzber et al. (2015) in various agronomic conditions (cropping systems, soil tillage, organic inputs). However, our study working at the rotation scale, the difference in soil C pools (POXC, OC, and TC) between the two rotations may have a cause other than the presence of legumes in crop rotation. In fact, control rotation included three cereals, beet and flax crop whose residues are exported leading to lower carbon stock in the soil. At the opposite, rotation including faba bean provided rich residues, which are incorporated to the soil enhancing organic matter content. Carbon additions from leguminous cover crops are relatively high compared to cereal crops (Drinkwater et al., 1998;

4.1.2. Effects on the structure of microbial communities 4.1.2.1. Abundance. Total microbial biomass was assessed according to 225

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the plant residues can influence the microbial community structure and catabolic diversity (Hamer and Makeschin, 2009). Then, in our case, the presence of faba bean seems to have selected bacterial communities able to degrade these complex compounds more than those which are able to degrade the amino acids compound. In addition, the difference in residues quality and quantity as well as root growth may influence the structure of soil microbial communities and then their potential metabolic diversity (Calbrix et al., 2005).

three approaches, total microbial biomass carbon, total DNA and total PLFA, which exhibited similar trends. Indeed, neither of these measures showed a significant difference between the two rotations although they describe different parts of the soil microbial biomass. Total DNA includes living and dead cells while microbial biomass and PLFA are commonly thought to only target intact living cells (Gangneux et al., 2011). Furthermore, our observations are in opposite with different correlations found between MBC and PLFA (White and Macnaughton, 1997) or between MBC and total DNA (Gangneux et al., 2011). In the present study, microbial biomass carbon and total DNA-RNA co-extracted seem to be slightly lower than value observed by Lebrun et al. (2012) and Laroche-Ajzenberg et al. (2012) who had studied these parameters in silt loam soil. Low water content and high temperature in the topsoil at the sampling date in both rotations might have influence our results (Schloter et al., 2003). These observations are in agreement with those of Franchini et al. (2007) who show that microbial biomass C seemed to be more sensitive to sampling period and soil management than to crop rotation. Additionally, Viana et al. (2011) demonstrate that total PLFA is significantly correlated to soil water content at sampling date and Blume et al. (2002) observed a variation of total PLFA between winter and summer sampling period in silt loam soil which support our previous conclusions. DNA analysis allows specific quantification of bacteria and fungi. In the present study, our results achieved an agreed outcome in agricultural soils namely the domination of bacteria in cultivated plots (Fierer et al., 2005; Janssen, 2006). This is due to the high sensitivity of fungi to soil disturbance. Agricultural machinery causes mechanical destruction of hyphae and reduces significantly the number of propagules in a soil horizon varying in depth according practices (Cookson et al., 2008; Helgason et al., 2009; Spedding et al., 2004). Furthermore, the positive correlation observed between total bacterial DNA and organic carbon and total nitrogen supported the results of some authors who showed that the amount of soil organic carbon and total nitrogen content influence the abundance of microbial groups (AsumingBrempong et al., 2008; Fierer et al., 2007). In our case, the higher amount of OC in Leg+ rotation increases the amount of bacterial DNA. In fact, the increasing of crop residues and its higher value of active carbon and organic carbon could be expected to increase bacteria community abundance in faba bean rotation (Li et al., 2015).

4.1.3. Effect on soil functions Enzymes activities related to C and N cycles were investigated as indicators of soil organic matter modification in response to the presence and the position of faba bean in rotation crops. The potential enzyme activities were within the range of values usually observed in silty loam soil (Lebrun et al., 2012). Our findings demonstrated that only β-glucosidase and arylamidase activities were positively affected by the presence of faba bean. In addition, these enzymes activities were correlated to active carbon and total nitrogen content, thus confirming their involvement in the mineralization of soil organic matter (Melero et al., 2009). Then, our observation can be attributed to the high organic carbon and total nitrogen contents incorporated to the soil through faba bean residues. Dodor and Tabatabai, (2002) have demonstrate that crops diversification induces a greater C addition and increases both C-cycle and N-cycle enzyme activities. Furthermore, the higher POXC content and carbon mineralization rate can be associated to the higher β-glucosidase activity which is involved in catalyzing the hydrolysis and biodegradation of plant debris (Ajwa and Tabatabai, 1994). On the other hand, cellulase and urease activities, which are widely used in the evolution of soil quality changes due to soil management (Nayak et al., 2007; Mikanova et al., 2009; Mangalassery et al., 2015), seemed to be not influenced by the presence of faba bean. For the rotation including this legume, the possible beneficial effects of faba bean may have been suppressed as a result of a lower moisture and higher topsoil temperature at sampling date. In fact, some studies reported that urease and cellulase activities are regulated by various factors such as climate or sampling dates (Krajewska, 2009; Lebrun et al., 2012) contrary to β-glucosidase activity which is particularly stable and has a low seasonal variability (Turner et al., 2002). 5. Conclusion

4.1.2.2. Diversity. PLFA profiling enable to assess a level of microbial diversity. Planting legume seems to induce an increase of Gramnegative bacteria. According to Bunemann et al. (2004), including legumes in corn crop rotation increase relative PLFA abundance of Gram-negative bacteria. In fact, rhizobium associated to leguminous are a Gram-negative bacteria and it supports basic pH. Abundance of this bacterial genus increases on legume grown and persists in soil even several years after (Voisin and Gastal, 2015). The lower value of saturated/monounsaturated fatty acids ratio in the final wheat of Leg+ rotation can probably be explained by the higher soil organic carbon content. Bossio et al. (1998) have shown that different farming regimes can influence PLFA profiling. These authors have demonstrated that a higher input of available carbon substrates to soil microorganisms induce changes in specific fatty acid relative abundances in soil communities such as monounsaturated fatty acids.

Our findings highlighted that the combination Faba bean-rape-wheat rotation seems to be, in our conditions, a suitable crop rotation to improve soil physicochemical and biological state. Then, this practice can be used as a part of an overall strategy to ameliorate soil fertility and preserve microorganisms. To confirm these positive effects of crop rotation with faba bean on soil microbial communities, further studies are needed in other agricultural contexts (i.e. different soil management). This result, associated with a higher or equivalent gross margin for farmers and higher environmental benefits for society of legume crop rotations (Reckling et al., 2015), confirm that the introduction of legume crop could contribute to the development of agroecology in European cropping systems. Acknowledgments This study was supported by the Region Haute-Normandie (France) through the GRR-TERA (Territory-Environment-Risk-Agronomy) and particularly its research network VASI (Vegetal-Agronomy-SoilInnovation). A. Aschi was supported by a regional grant also provided by the Region Haute-Normandie. PLFA analyses were achieved on the platform Biochem-Env, a service of the “Investment d’Avenir” infrastructure Ana EE-France, overseen by the French National Research Agency (ANR) (ANR-11-INBS-0001). Rachid Benabdallah (INRA ECOSYS) is acknowledged for its technical support. The authors acknowledge ARVALIS-vegetal institute-Rots for giving them an open access to his agricultural assay.

4.1.2.3. Potential metabolic diversity. The potential metabolic diversity of soil microbial communities evaluated by the Biolog Ecoplate showed significant differences between the soils of the two rotations. This trend indicates a difference in the functional structure of bacterial communities. Crop rotation including faba bean presented a greater rate of complex carbon sources utilization by soil bacterial communities than control rotation. In fact, Gregorich et al. (2001) showed that the organic matter in soils under rotations with legumes would present higher content of aromatic carbonate forms which would resist to biological degradation. Furthermore, the biochemical composition of 226

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