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ISRN Ecology
Volume 2012 (2012), Article ID 563191, 13 pages
http://dx.doi.org/10.5402/2012/563191
Research Article

Changes in Land Use System and Environmental Factors Affect Arbuscular Mycorrhizal Fungal Density and Diversity, and Enzyme Activities in Rhizospheric Soils of Acacia senegal (L.) Willd.

1Centre de Recherche de Bel-Air, Laboratoire Commun de Microbiologie, (IRD/ISRA/UCAD), BP 1386, 18524 Dakar, Senegal
2Département de Biologie Végétale, Faculté des Sciences et Techniques, Université Cheikh Anta Diop, BP 5005, Fann, Dakar, Senegal
3Centre Régional de Nkolbisson, IRAD: Institut de Recherche Agricole pour le Développement, P.O. Box 2067, Yaoundé, Cameroon
4Centre de recherche de Ouagadougou, Institut de Recherche pour le Développement (IRD), 01 BP 182 Ouagadougou, Burkina Faso
5Laboratoire Campus de Biotechnologies Végétales, Département de Biologie Végétale, Faculté des Sciences et Techniques, Université Cheikh Anta Diop, BP 5005, Fann, Dakar, Senegal
6Laboratoire des Symbioses Tropicales et Méditerranéennes, Cirad/IRD/Inra/Agro-M/UM2 Campus-international de Baillarguet, TA A-82 / J, 34398 Montpellier Cedex 5, France
7Laboratoire de Botanique et Biodiversité, Département de Biologie Végétale, Faculté des Sciences et Techniques, Université Cheikh Anta Diop, BP 5005, Fann, Dakar, Senegal
8CIRAD, Laboratoire des Symbioses Tropicales et Méditerranéennes, TA A-82 / J, 34398 Montpellier Cedex 5, France

Received 27 April 2012; Accepted 25 June 2012

Academic Editors: G. Brunialti and J. G. Zaller

Copyright © 2012 Fatou Ndoye et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract

The responses of the soil microbial community features associated to the legume tree Acacia senegal (L.) Willd. including both arbuscular mycorrhizal fungal (AMF) diversity and soil bacterial functions, were investigated under contrasting environmental conditions. Soil samples were collected during dry and rainy seasons in two contrasting rainfall sites of Senegal (Dahra and Goudiry, in arid and semiarid zone, resp.). Soils were taken from the rhizosphere of A. senegal both in plantation and natural stands in comparison to bulk soil. A multiple analysis revealed positive correlations between soil physicochemical properties, mycorrhizal potential and enzyme activities variables. The positive effects of A. senegal trees on soil mycorrhizal potential and enzyme activities indicates that in sahelian regions, AMF spore density and diversity as well as soil microbial functions can be influenced by land-use systems (plantation versus natural population of A. senegal) and environmental conditions such as moisture and soil nutrient contents. Our study underlines the importance of prior natural AMF screening for better combinations of A. senegal seedlings with AMF species to achieve optimum plant growth improvement, and for restoration and reforestation of degraded lands.

1. Introduction

Soil microorganisms and their enzymatic activities play key roles in the biochemical functioning of soils, including soil organic matter formation and degradation and nutrient cycling [1]. However, much less is known on the status of enzyme activities in semiarid regions as a function of land-use and management systems [2]. Additionally, the study of several enzyme activities together can provide information on the influence of soils, vegetation, and climatic factors on soil ecosystem functioning and quality [3]. This information would allow the selection of more sustainable and economically feasible cropping systems that guarantee the viability of agricultural activities in semiarid soils [2].

Arbuscular mycorrhizal fungi (AMF) are one of the most widespread and important components of the soil microbiota in natural and agricultural systems [4]. They form symbiotic associations with their host plants and improve their water and nutrient uptake like phosphorus (P), nitrogen (N), and micronutrients, and act as biocontrol agents against plant pathogens [5]. Furthermore, species composition and productivity of plant communities were shown to be conditioned by AMF species richness and diversity [6]. In Senegal, a few studies were done on the diversity of AMF [79]. For instance, Duponnois et al. [8] studied the diversity of AMF in soils from different aged fallows, and Manga et al. [9] studied that from rhizosphere of Acacia seyal. Although the mycorrhizal symbiosis is considered as a key factor to sustain vegetation cover in natural habitats, there are little information about AMF community assembly associated with Acacia senegal trees, particularly when considering land-use systems (natural population versus plantation) and the geographical rainfall gradient (north-south transect) in Senegal.

Likewise, the above- and below-ground environment is directly and indirectly influenced by trees. Distinct tree species affect chemical and microbial properties of soil to different extents [10]. Moreover, soil microbiological properties can be differently affected by communities of the same tree species depending on its management system (e.g., natural stands versus plantations) [11].

Acacia senegal (L.) Willd. is a multifunctional leguminous tree widely distributed in arid and semiarid zones of Africa and the Middle East. It has long been used for gum and fuelwood production and medicinal products [12]. This legume tree has a remarkable adaptability to drought and substantially contributes to soil fertility replenishment owing to its ability to symbiotically fix nitrogen. This tree is also known to host AMF symbionts and can sustainably improve soil mycorrhizal fpotential. Thus, A. senegal is used in agroforestry systems and reforestation programs [13]. Connecting vegetation characteristics and key soil community features could be useful to better understand natural forest ecosystems functioning and impact of forest conversion to plantation [14]. Thus, the objectives of this study are (i) to evaluate whether soil mycorrhizal infectivity and diversity as well as overall soil microbial community activities are influenced by land-use systems and environmental conditions and (ii) to investigate how soil enzyme activities, nutrient contents, and AMF community distribution are interrelated in our contrasted situations.

2. Materials and Methods

2.1. Study Sites and Samplings

The study sites (Dahra and Goudiry) were situated along a northern-southern region transect and concomitantly along a rainfall gradient in Senegal. Dahra, an arid zone, was located in the region of Louga (northern part of Senegal and at 300 km from Dakar, Lat. 15° 21 N; Long. 15° 29 W). Goudiry, a semiarid zone, was situated in the region of Tambacounda (southern part of Senegal and at 650 km from Dakar, Lat. 14° 11 N, Long. 12° 43 W). The average annual rainfall ranges from 300 to 400 mm and from 600 to 1000 mm, respectively, at Dahra and Goudiry. On each site, a natural population (naturally occurring plants) and a plantation of A. senegal (installed by humans by transplanting saplings of A. senegal) were selected. Each stand of A. senegal plantation is constituted of 4 plots and each plot comprised 25 trees. Stands of A. senegal natural population were constituted by trees of A. senegal irregularly distributed together with other species (Ziziphus spp., Combretum spp., Acacia seyal, etc.).

On each plot of plantation or natural stand, 5 trees were randomly chosen, soil samples were collected in the rhizosphere of the selected trees from the 0–25 cm layer and also in bulk soils. The soil samples from each origin were then mixed to obtain a composite soil sample. In each site (Dahra and Goudiry), soils were taken in dry (February 2008) and rainy (August 2008) seasons. A total of 12 composite soil samples (3 types x 2 sites x 2 seasons) were obtained for all sites and seasons. The composite samples were sieved (2 mm) and stored at 4°C.

In each plot of plantation or natural stand, 5 trees were randomly chosen and fine roots of A. senegal plants were collected in rainy season and pooled to get a composite sample.

2.2. Physical and Chemical Characterization of Soils

The analysis of physicochemical characteristics of soils (Table 1) was performed at LAMA (Laboratoire des Moyens Analytiques, certified ISO 9001 version 2000, IRD, Dakar, Senegal). The soil was sandy and sandy silt, respectively, at Dahra and Goudiry. Soil total C, N, and P contents were higher in A. senegal natural population than in A. senegal plantation and bulk soils. Hence, these mineral contents were greater in Goudiry than in Dahra. In contrast, soil available P content was higher in soils from Dahra than those from Goudiry. Soil pH was slightly acid in both sites.

tab1
Table 1: Physicochemical characteristics of soils from Dahra and Goudiry (in Senegal).
2.3. Assessment of A. senegal Root Colonization Rate

Root systems of A. senegal trees were cleared in a 10% KOH solution for 1 h at 90°C and stained with Trypan blue (0.05%) for 30 min [15]. Root segments (1 cm) were mounted on a microscope slide and observed under a compound microscope. The proportion of root length colonized by AMF was estimated [16].

2.4. AM Fungal Spore Extraction and Enumeration

In each site, as well as in plantation and in natural population of A. senegal and bulk soil, spores were recovered from a 100 g subsample soil (with 10 replicates per soil type) by wet sieving, decanting, and sucrose gradient centrifugation methods [17]. Then, the supernatant was poured successively through 50, 100, 200, and 400  𝜇 m pore-size sieves and rinsed with running tap water. Spore density (total number of spores in 100 g of dry soil) was determined by counting spores with a normal appearance under a compound microscope (40X).

2.5. Determination of AM Fungal Species Composition
2.5.1. AM Fungal Trap Culture and Spore Isolation

Trap culture was achieved for 5 months under glasshouse conditions. For each composite soil, 250 g of it were placed into 2 kg pots, containing 1 kg of sterilized (121°C, 2 hours) sand. Zea mays (L.) was sown as a trap plant with a density of 5 seeds per pot, with 4 replicates per soil origin. Pots were watered every two days with deionized water. Once a month, pots were fertilized with 100 ml of Long Ashton’s nutritive solution [18]. After 5 months, spores of AMF were isolated from the trap culture by wet sieving and decanting method. On the basis of differences in spore morphology (color under transmitted light, shape, size, and wall ornamentation), various taxa were recognized [19].

2.5.2. DNA Extraction from Spores and Amplification

Ten spore types were identified and DNA was extracted from each spore type using the Purelink Plant Total DNA Purification Kit (Invitrogen, France) following the manufacturer’s instructions.

The large subunit (LSU) region of the nuclear rDNA was used as target region for the PCR experiment. DNA was amplified in a 25 𝜇 L reaction volume containing 5  𝜇 L of 5X PCR reaction buffer, 0.3  𝜇 L GoTaq polymerase (0.5 U), 2  𝜇 L dNTPs (2.5 mM), 0.625  𝜇 L each primer (20  𝜇 M), 0.5  𝜇 L BSA (10 mg/mL), with 3  𝜇 L DNA template. The first fungal DNA amplification was performed using the fungal primers LR1/NDL22 [20]. The PCR was carried out as follows: 5 min at 93°C, followed by 35 cycles of 1 min at 93°C, 1 min at 58°C, 1 min at 72°C, and a final elongation of 10 min at 72°C. A 1  𝜇 L aliquot of the first PCR product was directly used as template for the second PCR amplification using the specific primers FLR3/FLR4 [21] under conditions described above. The nested PCR products were analyzed in electrophoresis using a 1% agarose gel (Sigma, France) in a Tris-Acetate-EDTA buffer with a DNA size standard (Eurogentec SmartLadder). The amplified fragments of about 400 pb were excised under UV light and purified with the Purelink Gel Extraction Kit (Invitrogen, France) according to the manufacturer’s instructions.

2.5.3. Cloning, Sequencing, and Phylogenetic Analysis

Two-round PCR products of the two AMF spore types (FNSP2 and FNSP8) were subcloned into XL-2 blue using the pGEM-T Easy Vector (Promega/Catalys, Wallisellen, Switzerland) following the manufacturer’s instructions. Ten positive clones for each spore type were reamplified using the primers pair FLR3/FLR4 and the products were revealed on agarose gel. Bands obtained were excised and purified as described above.

The purified PCR products were sequenced using primers FLR3/FLR4. Sequencing reactions were analyzed on a 3730 XL (Applied Biosystems) 96 capillary sequencers using a BigDye 3.1 Sequencing Kit (Genoscreen, France). Sequences were corrected using ChromasPro v1.33 (Technelysium Pty) and sequence similarities were determined by using the BLAST sequence similarity search tool [22] provided by GenBank. Corrected sequences were submitted to GenBank and were allocated with accession numbers. Multiple alignments and maximum likelihood tree were performed using the programs ClustalX [23] and MEGA version 5 [24], respectively.

2.6. Enzymatic Activities Assessment

The activity of 4 enzymes (fluorescein diacetate, dehydrogenase, and acid and alkaline phosphatases) was measured on the rhizospheric soil samples. For each enzymatic determination, controls with twice-autoclaved soil samples were included for nonenzymatic decomposition of the soil solution. The concentrations of enzymatic hydrolysis products were determined by spectrophotometer and compared with a standard curve.

2.6.1. FDA Activity

Total soil microbial activity potential was measured through fluorescein diacetate (3, 6′-diacetylfluorescein) hydrolysis assay, according to Alef et al. [25]. After 1 h of incubation on a rotary shaker, the fluorescein released from FDA was measured in the supernatant at 𝜆 = 4 9 0  nm and expressed as 𝜇 g FDA/h/g of soil.

2.6.2. Dehydrogenase Activity

Dehydrogenase activity was measured following the method of Skujiņs [26]. After 24 h of soil incubation in a triphenyl tetrazolium chloride (TTC) solution and a Tris-HCl buffer in darkness, followed by a second incubation at 37°C for 2 h on a rotary shaker, the extracted formazan was estimated at 𝜆 = 5 4 6  nm. Its concentration was calculated from a standard curve and dehydrogenase activity was expressed as 𝜇 g formazan/h/g of soil.

2.6.3. Acid and Alkaline Phosphatase Activities

Acid and alkaline phosphatase activities were assayed using a colorimetric determination of p-nitrophenol released when soil was incubated (during 1 h at 37°C) with p-nitrophenyl phosphate as substrate (pNPP, 5 mM) in pH 6 and pH 11 buffers, respectively [27]. The amount of p-nitrophenol released was determined by reading the optical density at 𝜆 = 4 0 0  nm and expressed as 𝜇 g pNPP/h/g of soil.

2.7. Statistical Analyses

Data were subjected to a one-way analysis of variance (ANOVA) using the SPSS software version 13. Mean values were compared using the Student-Newman-Keuls range test ( 𝑃 < 0 . 0 5 ). Percentage data of root colonization were arcsine transformed prior to analysis. Three-factor analysis of variance was performed using the Student-Newman-Keuls range test ( 𝑃 < 0 . 0 5 , 0 . 0 0 1 , a n d 0 . 0 0 0 1 ) of the XLSTAT version 13, in order to determine the main effects of sites, zones, and seasons and their interactions on soil enzyme activities.

The biochemical variables were grouped into 3 classes: (1) soil physicochemical properties including total organic C, N, P, and available P, soil pH and physical properties; (2) soil mycorrhizal parameters including AMF spore density and richness and root colonization rates; (3) enzymes variables including FDA hydrolysis, dehydrogenase, and acid and alkaline phosphatases. STATIS method was conducted on the soil physicochemical variables, as well as on the vegetation cover and soil microbial properties, to determine how these variables were interrelated. The STATIS method is introduced by Escoufier [28] and is used to analyze multiple data tables, each with information from the same set of individuals. It identifies what tables are alike, provides a summary table of all and describes the differences and similarities between the tables in relation to this summary table. These differences and similarities between said tables are analyzed by means of a structure called the compromise. The main steps of the STATIS method are interstructure, compromise, and intrastructure. A standardized principal component analysis (PCA) was used to balance the influence of different parameters. The implementation of this method was performed in the R environment using the software ADE4.

3. Results

3.1. Mycorrhizal Colonization of A. senegal Roots and Soil Spore Density

In Dahra, the natural occurrence of arbuscular mycorrhizas in A. senegal roots was not significantly different between natural population and plantation. However, in Goudiry, colonization rate was significantly lower in the natural stand compared to the plantation that presented value of root colonization not significantly different from those of the Dahra site (Figure 1).

563191.fig.001
Figure 1: Mycorrhizal colonization of roots from A. senegal plantations and natural stands in Dahra and Goudiry (in Senegal). For each site, bars with the same letter are not significantly different according to the Student-Newman-Keuls test ( 𝑃 < 0 . 0 5 ).

The density of AMF spores varied highly among the treatments from 290.71 spores per 100 g of soil (in bulk soil of Goudiry) to 2673.29 spores per 100 g of soil (in soil collected from A. senegal plantation in Dahra). In both sites, AMF spore abundance was significantly higher in soil samples from A. senegal plantations than those from natural populations and bulk soils (Figure 2).

563191.fig.002
Figure 2: AM fungal spore density in soils from Dahra and Goudiry (in Senegal). For each site, bars with the same letter are not significantly different according to the Student-Newman-Keuls test ( 𝑃 < 0 . 0 5 ).
3.2. AMF Species Richness and Community Composition

Ten AM fungal morphotypes belonging to 4 genera (Glomus, Scutellospora, Acaulospora, and Gigaspora) were isolated and identified as well in the rhizosphere of A. senegal in plantations and natural populations as in bulk soils (Table 2, Figure 3). In Dahra, 5 and 8 morphotypes representing 4 genera were found in A. senegal natural population and plantation, respectively, compared to 6 morphotypes within 3 genera in bulk soil (Glomus, Gigaspora, and Acaulospora). In Goudiry, rhizosphere of A. senegal plantation and natural population had the same number of AMF species (4) representing 2 genera (Glomus and Gigaspora). Finally, the bulk soil displayed 4 morphotypes within 3 genera (Glomus, Gigaspora, and Acaulospora). Three AMF spore morphotypes (FNSP1, FNSP2, and FNSP4) were found in all sites, while FNSP2 and FNSP9 were detected only at Dahra site. The AMF spore types FNSP5, FNSP7, and FNSP8 were only observed at the A. senegal plantation of Dahra and FNSP10 only at the bulk soil of Goudiry.

tab2
Table 2: AMF species richness in soils from plantations, natural stands of Acacia senegal, and bulk soils in Dahra and Goudiry (Senegal).
fig3
Figure 3: Spores of arbuscular mycorrhizal fungi isolated in soils from Dahra and Goudiry sites after trap culture. (a) Glomus sp. isolate FNSP1 in PVLG reagent, (b) Scutellospora sp. aff gregaria isolate FNSP5, (c) Glomus sp. isolate FNSP6, (d) Glomus globiferum isolate FNSP2 in PVLG + Melzer reagent, (e) Glomus globiferum isolate FNSP3 in PVLG + Melzer reagent, (f) Gigaspora gigantea isolate FNSP4 in PVLG reagent, (g) Glomus intraradices isolate FNSP7 in PVLG + Melzer reagent, (h) Acaulospora tuberculata isolate FNSP8 in PVLG + Melzer reagent, (i) Acaulospora longula isolate FNSP9 in PVLG + Melzer reagent, (j) Acaulospora longula isolate FNSP10 in PVLG + Melzer reagent.

BLAST search results of large subunit (LSU) sequences obtained from these AMF spores isolated from trap culture confirmed that the species belonged to the Glomeromycota. Results of the phylogenetic analysis of the LSU sequences (Figure 4) indicated the presence of 4 main groups: the first, Acaulospora group included FNSP9 and FNSP10 species which presented 92% of similarity with Acaulospora longula and the species FNSP8 which had 100% of homology with Acaulospora tuberculata. The species FNSP4 belonged to the Gigaspora group and had 100% of similarity with Gigaspora gigantea. The third, Glomus group B included FNSP2 and FNSP3 species which presented 100% of homology with Glomus globiferum. The last, Glomus group A encompassed the species FNSP1 and FNSP7 which presented 82% and 98% of homology with Glomus sp. isolate HE577801 and Glomus intraradices, respectively.

563191.fig.004
Figure 4: Maximum likelihood tree of partial LSU rDNA sequences from AMF spores from trap culture under soil samples collected in different origins in Senegal. Known AMF sequences from the NCBI database are included. Bootstrap values were estimated from 100 replicates and Mortierella polycephala was used as outgroup.

Morphological and anatomical identification indicated that the morphotype FNSP5 belonged to Scutellospora gregaria and the morphotype FNSP6 to Glomus sp.

3.3. Effect of Land Use and Moisture on Soil Enzyme Activities
3.3.1. Soils Collected in Dry Season

In Dahra, FDA hydrolysis activity was not significantly different between soils from the rhizosphere of A. senegal plantation, natural population, and bulk soil. By contrast, in Goudiry, soils from A. senegal natural population and bulk soil had the highest FDA hydrolysis activity. Furthermore, dehydrogenase activity was higher in soil from A. senegal natural population in Dahra, whereas in Goudiry, the greatest activity was found in soil from A. senegal plantation. In Dahra, the activity of acid and alkaline phosphatases was significantly greater in soil from natural population, whereas in Goudiry, the best responses were observed in soils from rhizosphere of A. senegal natural population and bulk soil (Table 3).

tab3
Table 3: Enzyme activities of soils from Dahra and Goudiry sampled during the dry season.
3.3.2. Soils Collected in Rainy Season

Table 4 presents the results obtained on the activity of soil enzymes in rainy season. FDA, dehydrogenase, and acid phosphatase activities were significantly higher in soils from A. senegal natural population and lower in those from bulk soils, in both sites. In Dahra, the highest alkaline phosphatase activity was observed in soil from A. senegal plantation and the lowest in bulk soil. In Goudiry, the greatest alkaline phosphatase activity was recorded in A. senegal natural population and the lowest in that from A. senegal plantation.

tab4
Table 4: Enzyme activities of soils from Dahra and Goudiry sampled during the rainy season.
3.4. Effects of Factors and their Interactions on Soil Enzyme Activities

The effects of different factors and their interactions on soil enzyme activities are summarized in Table 5. The effects of factors (sites, zones, and seasons) were highly significant ( 𝑃 < 0 . 0 0 0 1 ) on the activities of all enzymes except that of sites for acid phosphatase activity. The dual ((sites x zones), (sites x seasons), and (zones x seasons)), and triple (sites x zones x seasons) interactions were highly significant ( 𝑃 < 0 . 0 0 0 1 ) for FDA, dehydrogenase, and alkaline phosphatase activities. For acid phosphatase activity, the effects of (sites x seasons) and (zones x seasons) were highly significant and those of (sites x zones) and (sites x zones x seasons) significant at 𝑃 < 0 . 0 5 .

tab5
Table 5: Significance level (F-values) of effects of different factors and their interactions on soil enzyme activities based on three-way analysis of variance (ANOVA).
3.5. Relations between All Datasets

The plan of the first two axes of the interstructure (Figure 5(a)) explained 92.43% of the total variability and showed the isolation of mycorrhizal parameters from the physicochemical and enzymes tables. The results of the intrastructure gave a matrix vector correlation between the 3 tables. They showed a strong positive correlation between soil physicochemical properties and enzyme activities datasets (coefficient of variation R V = 0 . 7 7 ). In contrast, mycorrhizal parameters values were weakly correlated with soil physicochemical characteristics (coefficient of variation R V = 0 . 3 8 ) and enzyme activities variables (coefficient of variation R V = 0 . 3 3 ).

fig5
Figure 5: Interstructure and compromise from STATIS analysis: Interstructure and compromise (a) and intrastructure (b) of variables and sites. DPL: Dahra plantation of Acacia senegal, DNP: Dahra natural population of Acacia senegal, DBS: Dahra bulk soil, GPL: Goudiry plantation of Acacia senegal, GNP: Goudiry natural population of Acacia senegal, and GBS: Goudiry bulk soil.

For intrastructure of variables, the plan 1-2 of the compromise explained 73.92% of the total variability (Figure 5(b)). The first source of variability between groups of variables appeared on the axis 1 and concerned the physicochemical characteristics of soils and enzyme activities parameters. Soils rich in clay and silt were also rich in total C, N, and P and in base elements. In contrast, soils rich in sand and available P were rich in AMF spore density and species number. For intrastructure of sites, the analysis of the Figure 5(b) showed a classification of sites into 3 main groups. The first group was formed by the 3 soils from Dahra (DPL, DNP, and DBS), which were sandy and characterized by high amount of available P, AMF density and diversity, and low enzyme activities. The group 2 was composed by the soil from A. senegal plantation of Goudiry (GPL) rich in clay, silt, and Na and characterized by high dehydrogenase activity. Finally, the group 3 encompassed the soil from A. senegal natural population and bulk soil of Goudiry (GNP and GBS) which were rich in clay and silt with high amounts of total C, N, and P, correlated to high FDA and phosphatases activities.

4. Discussion

The present study clearly underlined that AMF density and diversity, and soil enzyme activities were strongly influenced by land-use systems, soil nutrient contents, and moisture. It also indicated that interrelations exist between the above parameters along our northern-southern transect in Senegal.

4.1. Effects of Tree Plantation and Soil Moisture on AM Fungal Spore Abundance and Diversity and on Root Colonization

Arbuscular mycorrhizal fungi (AMF) play an essential role in ecosystem functioning by influencing nutrient fluxes and organism interactions [29]. A set of reports has underlined the effect of vegetation on AMF formation, density, and diversity [30]. In this present work, the total number of spores per 100 g of dry soil was higher in rhizospheric soil from A. senegal than in bulk soil for all studied sites. This was in accordance with results of Karthikeyan and Selvaraj [31] on rhizospheric and nonrhizospheric soils of Ipomoea pes-caprae and Phyla nodiflora and also with observations made by Namenasrullah and coworkers [32] on wheat and maize crops under soils from different localities. These results suggested that AMF spore abundance varies depending on soil physicochemical properties and moisture as well as on vegetation cover. The presence of vegetation cover improves soil microbial biomass and activity by facilitating a vast network of AM hyphae and spores interconnecting the roots of the plant cover [33]. It was reported that lower plant density and vegetation supply less carbohydrates to the soil than needed by AMF [34], significantly reducing spores number and thus limiting AM infection. Importantly, soil texture may influence spore density [35]. Moreover, it can be inferred from our results that AMF spore density was greater in plantation than in natural population of A. senegal. Variations observed in soil physicochemical properties and plant density may encourage such AMF spores distribution [8, 36]. This present study reveals that AMF spore density and diversity were greater in Dahra (dry region) than in Goudiry (wet region). In wet soil, water availability would increase fungal mycelium growth for root colonization, leading to a decrease on spore germination [5]. The study also illustrates the diversity of AMF spores under soils from different land uses and climatic zones. Ten AMF morphotypes belonging to 4 genera (Glomus, Acaulospora, Scutellospora, and Gigaspora) were detected in the soils. These results were comparable to the diversity of AMF described in Senegal [7, 8] and in other countries [37]. In contrast, Opik et al. [38] described 34 different AMF taxa in a single habitat (a boreal herb-rich coniferous forest). Among the 10 AMF spores described here, the FNSP5 species closed to Scutellospora gregaria and the FNSP7 species closed to Glomus intraradices were previously described in Senegal by Duponnois et al. [8] and Manga et al. [9], respectively. The genus Glomus is the most abundant in these investigated sites and in many ecologically different environments from natural stands to managed agroecosystems in Senegal [7, 9] and other countries [39, 40]. Its dominance in arid and semiarid regions might be due to their remarkable adaptation to drastic conditions such as drought and salinity [41]. Our findings indicated that AMF species richness varied between sites (Dahra, Goudiry) and sampling areas (plantation, natural sand, and bulk soil). AMF species richness was more important in Dahra than in Goudiry. Klironomos and Hart [42] described large between- and within-site variations in the composition of AMF communities. Physical and chemical properties of soil and climatic factors such as soil moisture might explain differences observed between Dahra and Goudiry sites [43, 44]. The effect of land uses (plantation versus natural population) on the diversity of AMF was noticed in the site of Dahra, but not in that of Goudiry. Opik and coworkers [38] found in a boreal herb-rich coniferous forest that management practices had no adverse impact on the AMF biodiversity. These results suggested that the relationship between management practice and AMF richness is complex and may be positive [45] or negative [46]. Thus, AMF spore density and diversity can vary depending on edapho-climatic conditions including rainfall, soil texture, and management practices [30].

In order to ensure plants efficient growth, especially under semiarid regions which are often characterized by low levels of P availability in soil [47], it has been suggested to boost the native soil mycorrhizal potential through the culture of highly mycotrophic plants or through the controlled mycorrhization [48]. However, as it was noticed by Kumar et al. [49] on Sida cardifolia in different locations, variations were observed in the rates of AM colonization between root sampling sites. These variations could be attributed to factors such as soil nutrient content and fertility, soil moisture, changes in microhabitat, and acclimatization of a particular AMF genus/species to a particular location [49].

4.2. Spatial and Seasonal Variations of Soil Enzyme Activities

Soil enzyme activities in a forest floor are influenced by several variables such as nutrient availability and input of leaf litter [50]. Results from this work confirmed previous reports indicating that enzyme activities could vary with land-use systems and environmental factors [51, 52]. In Dahra, soil enzyme activities were higher in the rhizosphere of A. senegal and lower in bulk soil, whereas in Goudiry, the highest values (except dehydrogenase activity) were obtained in the rhizosphere of A. senegal natural population. These findings could be related to the differences in soil nutrient contents between soil under A. senegal and bulk soil. Importantly, increased enzyme activities could be attributed to increased organic matter and litter quality and quantity as well as improved soil physical parameters [53]. Particularly, legumes (by their rhizodeposition rich in amino acids and soluble sugars) help in solubilizing insoluble P in soil, improving the soil physical environment, and raising soil microbial activity [54].

When a between-sites comparison was performed, we noted that enzyme activities were greater in Goudiry than in Dahra. For instance, soils from Goudiry contained low available P but the highest total P and phosphatases activity. Similar results have been obtained by Gianfreda et al. [55] and recently by Wang et al. [56]. It is known that when available P is deficient in soil, soil biota can stimulate the production of extracellular phosphatases to increase the supply of inorganic P in soil [56]. As the two sites differ in their ecological parameters including soil moisture and soil fertility, it could be assumed that these parameters, which constitute key factors controlling the growth and survival of soil microbes [57], might have encouraged such spatial variation in enzyme activities.

The results indicated that soil enzyme activities studied here (except dehydrogenase activity) were greater in rainy season than in dry season. This enhancement could be attributed to an increase in water availability. Soil drying is known to reduce microbial activity and mineralization of organic C, N, and P [58] to decrease microbial mobility [59] and to restrict substrate and nutrient availability [60]. Interestingly, Fall et al. [61] in their study implemented in Senegal recorded a stimulation of microbial biomass in rhizosphere of A. senegal during the wet season. Finally, changes in the chemical composition of litter during its decomposition might also contribute to seasonality in enzyme activities [62].

4.3. Relationships between All Datasets

Results reported here showed that a set of relationships exists between soil enzyme activities, mycorrhizal variables, and physicochemical properties. Soil physicochemical properties are strongly related to the activities of studied enzymes ( R V = 0 . 7 7 ). Enzyme activities are in certain soils (GNP, GBS) positively correlated with total C and total N, as previously found by Acosta-Martinez et al. [2]. The positive correlation between enzyme activities and total N may be related to the low organic matter content of semiarid soils. However, a weak correlation between soil physicochemical properties and mycorrhizal datasets was noticed ( R V = 0 . 3 3 ). Several studies have indicated that soil nutrient content mainly phosphorus may impact AMF [7]. For instance, Grant et al. [63] has found that when the available P increases in soil, the amount of P also increases in the plant, and carbon drain on the plant by the AMF symbiosis becomes nonbeneficial to the plant. Our findings converged towards this affirmation when we considered soil samples from Goudiry, which had low number of AM spores concurrently with high level of total N, P, and C contents and phosphatase activity (which may further result in greater mobilization of P for plant nutrition). Nevertheless, differences in AMF spore density and species richness were observed in some cases and were not directly related to soil physicochemical characteristics. Other factors such as abiotic conditions (soil moisture and temperature, temporal and spatial variation in organic nutrient availability, etc.) and biotic constraints including the level of microbial degradation of enzyme molecules in soil might further be taken into consideration to fully address the question.

5. Conclusion

The present study suggests a positive effect of A. senegal trees on soil mycorrhizal potential and enzyme activities. It also indicates that in sahelian regions, AM fungal spore density and diversity as well as soil microbial functions can be influenced by land-use systems (plantation versus natural population of A. senegal) and environmental conditions such as moisture and soil nutrient contents. Strong relationships between enzyme activities and soil physicochemical properties have been noticed. This underlines the importance of prior natural AMF screening for better combinations of A. senegal seedlings with AMF species to achieve optimum plant growth improvement and environment protection.

Acknowledgments

This research was funded by the ACACIAGUM/INCO/STREP project (N° 032233). The authors are grateful to the French Embassy in Senegal (SCAC) and UNESCO for fellowships and to Oumar Sadio for statistical analyses.

References

  1. E. Kandeler, C. Kampichler, and O. Horak, “Influence of heavy metals on the functional diversity of soil microbial communities,” Biology and Fertility of Soils, vol. 23, no. 3, pp. 299–306, 1996. View at Publisher · View at Google Scholar · View at Scopus
  2. V. Acosta-Martínez, T. M. Zobeck, T. E. Gill, and A. C. Kennedy, “Enzyme activities and microbial community structure in semiarid agricultural soils,” Biology and Fertility of Soils, vol. 38, no. 4, pp. 216–227, 2003. View at Publisher · View at Google Scholar · View at Scopus
  3. E. Benitez, R. Nogales, M. Campos, and F. Ruano, “Biochemical variability of olive-orchard soils under different management systems,” Applied Soil Ecology, vol. 32, no. 2, pp. 221–231, 2006. View at Publisher · View at Google Scholar · View at Scopus
  4. R. D. Finlay, “Ecological aspects of mycorrhizal symbiosis: with special emphasis on the functional diversity of interactions involving the extraradical mycelium,” Journal of Experimental Botany, vol. 59, no. 5, pp. 1115–1126, 2008. View at Publisher · View at Google Scholar · View at Scopus
  5. S. E. Smith and D. J. Read, Mycorrhizal Symbiosis, Academic Press, London, UK, 3rd edition, 2008.
  6. M. G. A. Van Der Heijden, J. N. Klironomos, M. Ursic et al., “Mycorrhizal fungal diversity determines plant biodiversity, ecosystem variability and productivity,” Nature, vol. 396, no. 6706, pp. 69–72, 1998. View at Publisher · View at Google Scholar · View at Scopus
  7. T. A. Diop, M. Guèye, B. L. Dreyfus, C. Plenchette, and D. G. Strullu, “Indigenous arbuscular mycorrhizal fungi associated with Acacia albida Del. in different areas of Senegal,” Applied and Environmental Microbiology, vol. 60, no. 9, pp. 3433–3436, 1994. View at Scopus
  8. R. Duponnois, C. Plenchette, J. Thioulouse, and P. Cadet, “The mycorrhizal soil infectivity and arbuscular mycorrhizal fungal spore communities in soils of different aged fallows in Senegal,” Applied Soil Ecology, vol. 17, no. 3, pp. 239–251, 2001. View at Publisher · View at Google Scholar · View at Scopus
  9. A. Manga, T. A. Diop, D. E. van Tuinen, and M. Neyra, “Molecular diversity of arbuscular mycorrhizal fungi associated with Acacia seyal in a semiarid zone of Senegal,” Sécheresse, vol. 18, no. 2, pp. 129–133, 2007.
  10. Y. Zhang, L. D. Guo, and R. J. Liu, “Survey of arbuscular mycorrhizal fungi in deforested and natural forest land in the subtropical region of Dujiangyan, Southwest China,” Plant and Soil, vol. 261, no. 1-2, pp. 257–263, 2004. View at Publisher · View at Google Scholar · View at Scopus
  11. O. A. Oseni, M. M. Ekperigin, A. A. Akindahunsi, and G. Oboh, “Studies of physiochemical and microbial properties of soils from rainforest and plantation in Ondo state, Nigeria,” African Journal of Agricultural Research, vol. 2, no. 11, pp. 605–609, 2007.
  12. H. J. Von Maydell, Trees and Shrubs of the Sahel: their Characteristics and Uses, Deutsche Gesellschaft fuer Technische Zusammenarbeit (GTZ), Eschborn, Germany, 1986.
  13. M. E. Ballal, Yield trends of gum arabic from Acacia senegal as related to some environmental and managerial factor [Ph.D. thesis], Faculty of Forestry, University of Khartoum, Khartoum, Sudan, 2002.
  14. S. Piriyaprin, V. Sunanthapongsout, P. Limlong, C. Leavngv-utiviroj, and N. Pasda, “Study on soil microbial biodiversity in rhizophore of vetiver grass in degrading soil,” in Proceedings of the 17th World Congress of Soil Science, pp. 14–21, 2002.
  15. J. M. Phillips and D. S. Hayman, “Improved procedures for clearing roots and staining parasitic and vesicular-arbuscular mycorrhizal fungi for rapid assessment of infection,” Transactions of the British Mycological Society, vol. 55, pp. 158–161, 1970.
  16. A. Trouvelot, J. L. Kough, and V. Gianinazzi-Pearson, “Mesure du taux de mycorhization VA d'un système radiculaire. Recherche de méthodes d'estimation ayant une signification fonctionnelle,” in Mycorrhizae: Physiology and Genetics, V. Gianinazzi-Pearson and S. Gianinazzi, Eds., pp. 217–221, INRA, Paris, France, 1986.
  17. J. W. Gerdemann and T. H. Nicolson, “Spores of mycorrhizal endogone species extracted from soil by wet sieving and decanting,” Transactions of the British Mycological Society, vol. 46, pp. 235–244, 1963.
  18. V. Furlan, “Techniques et procédures pour la culture des champignons endomycorhiziens,” Note Technique 65, Université Laval, Québec, Canada, 1981.
  19. M. Brundrett, N. Bougher, B. Dell, T. Grove, and N. Malajczuk, Working with Mycorrhizas in Forestry and Agriculture, Pirie Printers, Canberra, Australia, 1996.
  20. D. E. van Tuinen, E. Jacquot, B. Zhao, A. Gollotte, and V. Gianinazzi-Pearson, “Characterization of root colonization profiles by a microcosm community of arbuscular mycorrhizal fungi using 25s rDNA-targeted nested PCR,” Molecular Ecology, vol. 7, no. 7, pp. 879–887, 1998. View at Publisher · View at Google Scholar · View at Scopus
  21. A. Gollotte, D. E. van Tuinen, and D. Atkinson, “Diversity of arbuscular mycorrhizal fungi colonising roots of the grass species Agrostis capillaris and Lolium perenne in a field experiment,” Mycorrhiza, vol. 14, no. 2, pp. 111–117, 2004. View at Publisher · View at Google Scholar · View at Scopus
  22. S. F. Altschul, T. L. Madden, A. A. Schäffer et al., “Gapped BLAST and PSI-BLAST: a new generation of protein database search programs,” Nucleic Acids Research, vol. 25, no. 17, pp. 3389–3402, 1997. View at Publisher · View at Google Scholar · View at Scopus
  23. J. D. Thompson, T. J. Gibson, F. Plewniak, F. Jeanmougin, and D. G. Higgins, “The CLUSTAL X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools,” Nucleic Acids Research, vol. 25, no. 24, pp. 4876–4882, 1997. View at Publisher · View at Google Scholar · View at Scopus
  24. K. Tamura, D. Peterson, N. Peterson, G. Stecher, M. Nei, and S. Kumar, “MEGA5: molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods,” Molecular Biology and Evolution, vol. 28, no. 10, pp. 2721–2739, 2011.
  25. K. Alef, “Estimation of hydrolysis of fluorescein diacetate,” in Methods in Applied Soil Microbiology and Biochemistry, K. Alef and P. Nannipieri, Eds., pp. 232–233, Academic Press, London, UK, 1998.
  26. J. Skujiņs, “Extracellular enzymes in soil,” CRC Critical Reviews in Microbiology, vol. 4, no. 4, pp. 383–421, 1976. View at Scopus
  27. M. A. Tabatabai and J. M. Bremner, “Use of p-nitrophenyl phosphate for assay of soil phosphatase activity,” Soil Biology and Biochemistry, vol. 1, no. 4, pp. 301–307, 1969. View at Scopus
  28. Y. Escoufier, “L'analyse des tableaux de contingence simples et multiples,” Metron, vol. 40, pp. 53–77, 1982.
  29. M. G. A. van der Heijden, S. Verkade, and S. J. de Bruin, “Mycorrhizal fungi reduce the negative effects of nitrogen enrichment on plant community structure in dune grassland,” Global Change Biology, vol. 14, no. 11, pp. 2626–2635, 2008. View at Publisher · View at Google Scholar · View at Scopus
  30. S. L. Stürmer and J. O. Siqueira, “Species richness and spore abundance of arbuscular mycorrhizal fungi across distinct land uses in Western Brazilian Amazon,” Mycorrhiza, vol. 21, no. 4, pp. 255–267, 2011. View at Publisher · View at Google Scholar · View at Scopus
  31. C. Karthikeyan and T. Selvaraj, “Diversity of arbuscular mycorrhizal fungi (AMF) on the coastal saline soils of the West coast of Kerala, Southern India,” World Journal of Agricultural Science, vol. 5, pp. 803–809, 2009.
  32. Namenasrullah, M. Sharif, K. Rubina, and T. Burni, “Occurrence and distribution of arbuscular mycorrhizal fungi in wheat and maize crops of Malakand division of North West Frontier Province,” Pakistan Journal of Botany, vol. 42, no. 2, pp. 1301–1312, 2010. View at Scopus
  33. J. D. Graves, N. K. Watkins, A. H. Fitter, D. Robinson, and C. Scrimgeour, “Intraspecific transfer of carbon between plants linked by a common mycorrhizal network,” Plant and Soil, vol. 192, no. 2, pp. 153–159, 1997. View at Publisher · View at Google Scholar · View at Scopus
  34. A. Michelsen, N. Lisanework, and I. Friis, “Impacts of tree plantations in the Ethiopian highland on soil fertility, shoot and root growth, nutrient utilisation and mycorrhizal colonisation,” Forest Ecology and Management, vol. 61, no. 3-4, pp. 299–324, 1993. View at Scopus
  35. A. H. Meyer, A. Botha, A. J. Valentine, E. Acher, and P. J. E. Louw, “The occurrence and infectivity of arbuscular mycorrhizal fungi in inoculated and uninoculated rhizosphere soils of two-year-old commercial grapevines,” South African Journal for Enology and Viticulture, vol. 26, no. 2, pp. 90–94, 2005.
  36. L. M. Carvalho, P. M. Correia, R. J. Ryel, and M. A. Martins-Loução, “Spatial variability of arbuscular mycorrhizal fungal spores in two natural plant communities,” Plant and Soil, vol. 251, no. 2, pp. 227–236, 2003. View at Publisher · View at Google Scholar · View at Scopus
  37. Y. Abbas, M. Ducousso, M. Abourouh, R. Azcón, and R. Duponnois, “Diversity of arbuscular mycorrhizal fungi in Tetraclinis articulata (Vahl) Masters woodlands in Morocco,” Annals of Forest Science, vol. 63, no. 3, pp. 285–291, 2006. View at Publisher · View at Google Scholar · View at Scopus
  38. M. Öpik, M. Moora, M. Zobel et al., “High diversity of arbuscular mycorrhizal fungi in a boreal herb-rich coniferous forest,” New Phytologist, vol. 179, no. 3, pp. 867–876, 2008. View at Publisher · View at Google Scholar · View at Scopus
  39. M. Öpik, M. Moora, J. Liira, and M. Zobel, “Composition of root-colonizing arbuscular mycorrhizal fungal communities in different ecosystems around the globe,” Journal of Ecology, vol. 94, no. 4, pp. 778–790, 2006. View at Publisher · View at Google Scholar · View at Scopus
  40. I. Hijri, Z. Sýkorová, F. Oehl et al., “Communities of arbuscular mycorrhizal fungi in arable soils are not necessarily low in diversity,” Molecular Ecology, vol. 15, no. 8, pp. 2277–2289, 2006. View at Publisher · View at Google Scholar · View at Scopus
  41. J. Błaszkowski, M. Tadych, and T. Madej, “Arbuscular mycorrhizal fungi (Glomales, Zygomycota) of the Błedowska Desert, Poland,” Acta Societatis Botanicorum Poloniae, vol. 71, no. 1, pp. 71–85, 2002. View at Scopus
  42. J. N. Klironomos and M. M. Hart, “Colonization of roots by arbuscular mycorrhizal fungi using different sources of inoculum,” Mycorrhiza, vol. 12, no. 4, pp. 181–184, 2002. View at Publisher · View at Google Scholar · View at Scopus
  43. E. Lumini, A. Orgiazzi, R. Borriello, P. Bonfante, and V. Bianciotto, “Disclosing arbuscular mycorrhizal fungal biodiversity in soil through a land-use gradient using a pyrosequencing approach,” Environmental Microbiology, vol. 12, no. 8, pp. 2165–2179, 2010. View at Publisher · View at Google Scholar · View at Scopus
  44. P. L. Leal, S. L. Stürmer, and J. O. Siqueira, “Occurrence and diversity of arbuscular mycorrhizal fungi in trap cultures from soils under different land use systems in the amazon, Brazil,” Brazilian Journal of Microbiology, vol. 40, no. 1, pp. 111–121, 2009. View at Scopus
  45. M. Vallino, N. Massa, E. Lumini, V. Bianciotto, G. Berta, and P. Bonfante, “Assessment of arbuscular mycorrhizal fungal diversity in roots of Solidago gigantea growing in a polluted soil in Northern Italy,” Environmental Microbiology, vol. 8, no. 6, pp. 971–983, 2006. View at Publisher · View at Google Scholar · View at Scopus
  46. L. Whitfield, A. J. Richards, and D. L. Rimmer, “Relationships between soil heavy metal concentration and mycorrhizal colonisation in Thymus polytrichus in Northern England,” Mycorrhiza, vol. 14, no. 1, pp. 55–62, 2004. View at Publisher · View at Google Scholar · View at Scopus
  47. V. Estaún, R. Savé, and C. Biel, “AM inoculation as a biological tool to improve plant revegetation of a disturbed soil with Rosmarinus officinalis under semi-arid conditions,” Applied Soil Ecology, vol. 6, no. 3, pp. 223–229, 1997. View at Scopus
  48. A. Sanon, F. Ndoye, E. Baudoin, Y. Prin, A. Galiana, and R. Duponnois, “Management of the mycorrhizal soil infectivity to improve reforestation programs’ achievements in Sahelian ecosystems,” in Current Research, Technology and Education Topics in Applied Microbiology and Microbial Biotechnology, A. Méndez-Vilas, Ed., vol. 1, pp. 230–238, 2010.
  49. K. V. C. Kumar, K. R. Chandrashekar, and R. Lakshmipathy, “Variation in arbuscular mycorrhizal fungi and phosphatase activity associated with Sida cardifoliain Karnataka,” World Journal of Agricultural Science, vol. 4, no. 6, pp. 770–774, 2008.
  50. L. Kong, Y. B. Wang, L. N. Zhao, and Z. H. Chen, “Enzyme and root activities in surface-flow constructed wetlands,” Chemosphere, vol. 76, no. 5, pp. 601–608, 2009. View at Publisher · View at Google Scholar · View at Scopus
  51. L. P. Qiu, J. Liu, Y. Q. Wang, H. M. Sun, and W. X. He, “Research on relationship between soil enzyme activities and soil fertility,” Plant Nutrition and Fertility Science, vol. 10, pp. 277–280, 2004.
  52. Z. J. Shi, Y. Lu, Z. G. Xu, and S. L. Fu, “Enzyme activities of urban soils under different land use in the Shenzhen city, China,” Plant, Soil and Environment, vol. 54, no. 8, pp. 341–346, 2008. View at Scopus
  53. B. A. Caldwell, “Enzyme activities as a component of soil biodiversity: a review,” Pedobiologia, vol. 49, no. 6, pp. 637–644, 2005. View at Publisher · View at Google Scholar · View at Scopus
  54. A. Ghosh, M. Bhardwaj, T. Satyanarayana, M. Khurana, S. Mayilraj, and R. K. Jain, “Bacillus lehensis sp. nov., an alkalitolerant bacterium isolated from soil,” International Journal of Systematic and Evolutionary Microbiology, vol. 57, no. 2, pp. 238–242, 2007. View at Publisher · View at Google Scholar · View at Scopus
  55. L. Gianfreda, R. M. Antonietta, A. Piotrowska, G. Palumbo, and C. Colombo, “Soil enzyme activities as affected by anthropogenic alterations: intensive agricultural practices and organic pollution,” Science of the Total Environment, vol. 341, no. 1–3, pp. 265–279, 2005. View at Publisher · View at Google Scholar · View at Scopus
  56. J. B. Wang, Z. H. Chen, L. J. Chen, A. N. Zhu, and Z. J. Wu, “Surface soil phosphorus and phosphatase activities affected by tillage and crop residue input amounts,” Plant, Soil and Environment, vol. 57, no. 6, pp. 251–257, 2011. View at Scopus
  57. E. Gömöryová, “Small-scale variation of microbial activities in a forest soil under a beech (Fagus sylvatica L.) stand,” Polish Journal of Ecology, vol. 52, no. 3, pp. 311–321, 2004. View at Scopus
  58. M. Pulleman and A. Tietema, “Microbial C and N transformations during drying and rewetting of coniferous forest floor material,” Soil Biology and Biochemistry, vol. 31, no. 2, pp. 275–285, 1999. View at Publisher · View at Google Scholar · View at Scopus
  59. D. M. Grifin, “Water potential as a selective factor in the microbial ecology of soil,” in Water Potential Relations in Soil Microbiology, J. F. Parr, W. R. Gardner, and L. F. Elliott, Eds., pp. 141–151, Soil Science Society and America, Madison, Wis, USA, 1981.
  60. L. E. Sommers, C. M. Gilmour, R. E. Wildung, and S. M. Beck, “The effect of water potential on decomposition processes in soil,” in Water Potential Relation in Soil Microbiology, J. F. Parr, W. R. Gardner, and L. F. Elliot, Eds., pp. 97–117, Soil Science Society of America, Madison, Wis, USA, 1981.
  61. D. Fall, D. Diouf, A. M. Zoubeirou, N. Bakhoum, A. Faye, and S. N. Sall, “Effect of distance and depth on microbial biomass and mineral nitrogen content under Acacia senegal (L.) Willd. trees,” Journal of Environmental Management, vol. 95, pp. S260–S264, 2011. View at Publisher · View at Google Scholar · View at Scopus
  62. K. Styla and A. Sawicka, “Seasonal changes in biochemical and microbiological activity of soil against the background of differentiated irrigation in an apple tree orchard after replantation,” Agronomic Research, vol. 7, no. 1, pp. 113–124, 2009.
  63. C. Grant, S. Bittman, M. Montreal, C. Plenchette, and C. Morel, “Soil and fertilizer phosphorus: effects on plant P supply and mycorrhizal development,” Canadian Journal of Plant Science, vol. 85, no. 1, pp. 3–14, 2005. View at Scopus