Open Access

Arbuscular mycorrhizal fungal community assembly in the Brazilian tropical seasonal dry forest

  • Tancredo Augusto Feitosa de Souza1Email author and
  • Helena Freitas1
Ecological Processes20176:2

https://doi.org/10.1186/s13717-017-0072-x

Received: 9 August 2016

Accepted: 6 January 2017

Published: 10 January 2017

Abstract

Introduction

Here, we compare the arbuscular mycorrhizal fungal (AMF) community composition in soils from the root zone of the exotic invasive species Prosopis juliflora (EXO soils) and soils from the root zone of the native species Mimosa tenuiflora (NAT soils) from five locations in the Brazilian tropical seasonal dry forest, Paraíba, Brazil, using morphological analyses.

Results

AMF community composition in EXO and NAT soils were dissimilar. Available phosphorus, diversity index, spore abundance, and species richness were the main factors differing between the EXO and NAT soils. In general, the most dominant order present in the soils were Glomerales (44.8%) and Gigasporales (41.4%). The most abundant AMF genus in all studied soils was Funneliformis.

Conclusions

Differences in AMF community composition were associated with (1) differences in the dominant plant species (P. juliflora vs. M. tenuiflora) and (2) changes in soil chemical factors (soil, pH, total organic carbon, total nitrogen, and available P) in EXO soils. These results contribute to a deeper view of the AMF communities in exotic soils and open new perspectives for ecological processes involving AMF species and exotic plant species in the Brazilian tropical seasonal dry forest.

Keywords

Glomeromycota AMF community AMF diversity Native plant species Exotic plant species Caatinga

Introduction

The Brazilian tropical seasonal dry forest, also referred to as “Caatinga”, was described by Santos et al. (2011) as a “type of desert vegetation, which consists of small, thorny trees that shed their leaves seasonally in interior northeastern Brazil.” This ecoregion consists of a broad mosaic vegetation type that covers 850,000 km2 (nearly 10% of Brazil’s territory) in Brazilian Northeast. It has a unique biota that contains over 1000 endemic plant species, but unfortunately, it is poorly conserved, with only 1% of its territory in protection conservation areas (Alves et al. 2009; Andrade et al. 2009). Drought, intensive agriculture, excessive grazing, and biological invasion are recognized as the major threats with significative negative environmental impacts on plant diversity (Pegado et al. 2006; Andrade et al. 2008; Andrade et al. 2009).

Despite evidences of invasive exotic plant species introduction in the Brazilian tropical seasonal dry forest dating 516 years before present, when the Portuguese arrived in Brazil in 1500, Pegado and co-workers (2006) reported that invasive exotic plant species (e.g., Prosopis juliflora (Sw.) DC.) were introduced during 1942 as a result of a governmental program during that period. This governmental program was proposed to help the regional farmers during dry period providing them with a “tree of life” that would be used as an alternative to fodder and shelter for livestock. But, in fact, this “tree of life” became the most common troublesome invasive exotic plant species in the Brazilian tropical seasonal dry forests affecting native plant community composition and ecological processes in the invaded areas since 1942 (Alves et al. 2009; Andrade et al. 2009; Souza et al. 2016a).

Many studies were done to shown how invasion by P. juliflora alter the native plant community (e.g., Pegado et al. 2006; Andrade et al. 2008; Andrade et al. 2009). These works showed negative impacts of P. juliflora on native plant diversity and plant community structure. But, it is still unclear if the biological invasion in Brazilian Northeast has negative impacts on arbuscular mycorrhizal fungal (AMF) diversity. Invasive plant species need AMF because these soil microorganisms increase nutrient acquisition, growth, and vitality of their hosts, independently if they are exotic or native plant species. In semi-arid conditions, as is in our case, AMF are also crucial for the protection of their hosts against abiotic (drought, salinity) stresses and have been found to determine plant community composition and function in other studies (Reinhart and Callaway 2006; Smith and Read 2008; Hodge and Storer 2014; Shah et al. 2009).

The consensus is that the biological invasion alters the dynamics of plant communities and soil characteristics (e.g., soil pH and nutrient availability), which may influence AMF community structure and functioning by three different mechanisms according to studies from elsewhere (Parkash and Aggarwal 2009; Rodríguez-Echeverría et al. 2009; Zhang et al. 2010; Kivlin and Hawkes 2011; Lekberg et al. 2013). First, invasive exotic plant species usually form large monospecific plant populations, thus reducing the diversity of host-plants available to the AMF community. Second, metabolites produced by invaders negatively affect native plants growth by disrupting their mutualistic associations with the AMF community from native soils (Zubek et al. 2016). Finally, the introduction of invasive exotic plant species might cause changes in soil chemical properties that may indirectly affect AMF community composition and, thus, contribute to a successful establishment and spread of invasive exotic plant species (Shah et al. 2009).

We examined the AMF community from P. juliflora root zone (EXO soils) and compared it with the AMF community from Mimosa tenuiflora root zone (NAT soils). Our study addressed the following questions. (1) Do invasive exotic plant species alter the composition of the AMF community in the studied areas? Based on the enhanced mutualisms hypothesis (Richardson et al. 2000; Reinhart and Callaway 2006), we expected to find evidence for changes in the original AMF community. (2) Is there evidence for differences in the relationship between invasive and native plants and the AMF community from NAT and EXO soils in field conditions? We hypothesized that invasive plants would experience a stronger interaction with a specific AMF community according to the invasion opportunity windows and resource enemy release hypotheses (Johnstone 1986; Agrawal et al. 2005; Blumenthal 2005; Kulmatiski and Kardol 2008). To accomplish this, we perform field sampling of two root zone types, i.e., invasive and native root zones, characterized by both the soil chemical properties and AMF communities.

Methods

Plant species and study sites

Field sampling was carried out in five different locations in the Brazilian tropical seasonal dry forest, Paraiba, Brazil (Algodão de Jandaíra, 06° 46′ 17.3″ S, 36° 01′55.3″ W; Esperança, 06° 56′ 45.7″ S, 35° 54′ 06.8″ W; Juazeirinho, 07° 06′ 33.3″ S, 36° 34′ 34.2″ W; Monteiro, 07° 48′ 19.8″ S, 37° 10′ 32.4″ W; and Natuba, 07° 37′ 34.9″ S, 35° 32′ 24.5″ W). These study areas are classified as Bsh following Köppen-Geiger climate classification, i.e., hot semi-arid with hot summers and mild to warm winters, annual precipitation and temperature of 600 mm and 30 °C, respectively. In these sites, rainfall is highly reduced, unpredictable, and irregular (Alves et al. 2009).

We selected the invasive exotic plant species P. juliflora and the native plant species M. tenuiflora which co-occur in a mixed plant community (well-mixed stand) in all studied areas. We also have selected these two plant species to perform our study because P. juliflora was introduced in 1942 and nowadays is the most common troublesome invasive exotic plant species in all studied areas and M. tenuiflora is highly abundant in Brazilian tropical seasonal dry forests (Pegado et al. 2006; Andrade et al. 2009; Souza et al. 2016b). The soil type of the studied areas was classified as a sandy loam Dystric Fluvisols (WRB 2006).

Field sampling and soil characterization

In each of the five study areas, we established 40 plots of 100 m2 according to Fortin and Dale (2005). Within each plot, we selected one plant of each target species in a well-mixed stand (i.e., 2 plants per plot, thus 80 plants per study location, or 400 total plants across all locations) according to the following criteria: (1) the plant had a diameter near the soil surface of >3 cm and (2) no individuals from a different plant species were growing in a 3-m radius to the sampling point in all directions (Daubenmire 1968; Fortin and Dale 2005; Caifa and Martins 2007; Costa and Araújo 2007; Durigan 2009). Soil samples (including soil and root fragments) were collected near the drip line and beyond (0–20 cm deep), during the dry period, i.e., at the beginning of September 2012. By sampling during the dry season, we guaranteed that we sampled the largest number of AMF species because fungal sporulation is expected to be higher at this time of the year in semi-arid environments (Silva et al. 2014). Samples from each plant species in each plot were bulked, mixed, and stored at 4 °C until host-plant bioassays. During sampling and handling of each soil sample, precautions (e.g., sterilization with ethanol and gloves) were undertaken to avoid cross contamination. Later, each sample collected from the field was divided into portions intended for chemical soil characterization and AMF community assessment.

To chemically characterize the soil from each plot, we analyzed soil pH, total organic carbon, total nitrogen, and available phosphorus (N = 40 by plant species). Soil pH was measured in a suspension of soil and distilled water (1:2.5 v:v, soil:water suspension) (Black 1965). Total organic carbon was estimated according to the methodology described by Okalebo et al. (1993). To quantify total nitrogen, soil samples were first digested with sulfuric acid plus potassium sulfate and we then followed the protocol described by Kjeldahl (Black 1965). Available phosphorus (Olsen’s P) was determined colorimetrically using a spectrophotometer at 882 nm by extraction with sodium bicarbonate for 30 min (Olsen et al. 1954).

Arbuscular mycorrhizal fungi community characterization

AMF communities extracted from native and exotic soils were classified as “NAT” if they occurred in the root zone of M. tenuiflora and as “EXO” if they occurred in the root zone of the invasive exotic plant species P. juliflora. Spores from field were extracted by the wet sieving technique (Gerdemann and Nicolson 1963) followed by sucrose centrifugation (Jenkins 1964). For this, we used 100 g of field soil. Initially, the extracted spores were examined in water under a dissecting microscope and they were separated based on morphology. Subsequently, they were mounted in polyvinyl alcohol lactoglycerol (PVLG) with or without addition of Melzer’s reagent (Walker et al. 2007). Species identification was based on the descriptions provided by Schenck and Perez (1987), publications with descriptions of new families and genera (i.e., Oehl et al. 2008), and by consulting the international culture collection of arbuscular mycorrhizal fungi database—INVAM (http://invam.caf.wvu.edu). In this work, we followed the classification proposed by Oehl et al. (2011), including recently new described taxa (i.e., Goto et al. 2012; Redecker et al. 2013; Sieverding et al. 2014). In addition to species identification, we also assessed spore abundance by counting the total number of spores, spore abundance of each AMF species by recording the number of spores of each AMF species recorded in the samples, and the species occurrence frequency (FOi) of each AMF species. FOi was calculated using the following equation:
$$ \mathrm{F}\mathrm{O}\mathrm{i} = \mathrm{n}\mathrm{i}/\mathrm{N} $$
where ni is the number of times an AMF species was observed and N is the total of AMF spores observed from each studied area.

Mycorrhizal root colonization assessment

Roots of P. juliflora and M. tenuiflora were examined for quantification of AMF colonization. The collected roots were stored in 50% ethanol until staining. Roots were cleared in 2% KOH for 1 h at 90 °C. Subsequently, they were left to acidify overnight in 1% HCl. Staining was done with blue ink (Parker Quink) for 30 min at 60 °C, followed by destaining in lactoglycerol. The amount of colonization was estimated using a grid-intersect method with examination of 100 intersects under a compound microscope at ×200 magnification (Phillips and Hayman 1970; McGonigle et al. 1990). All microscopic examinations were carried out by the same person. Root intersects that contained vesicles, arbuscules, and hyphae were scored as mycorrhizal. The decision to score hyphae as mycorrhizal was based on the associated presence of vesicles, arbuscules, spores, and the morphology of the mycelium. Roots that did not have cortex were excluded from the analysis. In total, 40 samples (20 samples from EXO soils plus 20 samples from NAT soils) per studied area were examined to score 100 intersections.

Glomalin analysis

To quantify total glomalin, soil samples (1.0 g) were used with 8.0 mL of 50 mmol L−1 sodium citrate at pH 8.0 over three cycles of autoclaving at 121 °C, for 1 h per cycle. The extractor was separated from the soil via protein centrifugation at 1720g for 15 min. The supernatant was quantitated in a mass spectrophotometer by the protocol described by Wright and Upadhyaya (1998).

Ecological indices and statistical analyses

After AMF species identification, we calculated the following ecological indices: diversity index (H) proposed by Shanon and Weaver (1949), and dominance index (C) proposed by Simpson (1949) for each studied AMF community.

Differences in soil properties and AMF community structure between EXO and NAT soil groups were determined by non-parametric t test followed by Monte Carlo test (100 replicates). Univariate analyses (t test, ANOVA, and Tukey’s test) were performed using SAS 9.1.3 Portable, whereas multivariate analysis (principal component analysis) was performed using MVSP (MultiVariate Statistical Package) 3.1 (Kovach 2007). All data were checked for normality and homogeneity of variances before analyses. Count data and environmental variable values were transformed (function square root) before multivariate analysis. The relationships between the AMF community structure and soil properties were examined using the correlation analyses by the Pearson correlation coefficient.

Results

Soil properties

Significant differences (p ≤ 0.05) between the EXO and NAT soils were found for soil chemical properties. On average, EXO soils were higher in soil pH, total organic carbon (TOC), total nitrogen (total N), and available phosphorus (P) (Table 1). Within the EXO soils, there were no significant differences among the EXO sites for soil pH, total organic carbon, and total nitrogen, but we found significant differences among EXO sites for available P. The Monteiro samples showed the highest soil pH and amounts of P, while the Natuba samples showed the highest amounts of TOC and total N, whereas in the NAT soils, we found significant difference from each studied site in soil pH, total organic carbon, total nitrogen, and available P. In the NAT soils, the Esperança and Juazeirinho soil samples showed the highest soil pH, while the Natuba samples showed the highest amounts of TOC, total N, and P (Table 1).
Table 1

Chemical soil attributes of exotic (EXO) and native (NAT) soils from the Brazilian tropical seasonal dry forest (mean ± SD, N = 40)

Soil type/site

Soil pH (H2O)

Total organic carbon (g kg−1)

Total nitrogen (g kg−1)

Available P (ppm)

EXO

 Algodão de Jandaíra

6.64 ± 0.26 a

8.56 ± 0.55 a

0.86 ± 0.15 a

8.49 ± 0.07 c

 Esperança

6.71 ± 0.20 a

8.44 ± 0.30 a

0.82 ± 0.10 a

8.42 ± 0.30 c

 Juazeirinho

6.35 ± 0.46 a

8.99 ± 0.57 a

0.88 ± 0.09 a

6.97 ± 0.47 d

 Monteiro

6.87 ± 0.26 a

8.81 ± 0.71 a

0.87 ± 0.07 a

10.70 ± 0.52 a

 Natuba

6.50 ± 0.35 a

9.14 ± 1.78 a

0.92 ± 0.17 a

9.22 ± 0.22 b

NAT

 Algodão de Jandaíra

5.04 ± 0.03 b

2.78 ± 0.16 d

0.15 ± 0.04 d

1.93 ± 0.02 f

 Esperança

5.20 ± 0.10 b

2.00 ± 0.10 e

0.21 ± 0.01 c

2.00 ± 0.10 f

 Juazeirinho

5.20 ± 0.29

2.52 ± 0.59 d

0.22 ± 0.07 c

1.74 ± 0.26 f

 Monteiro

4.87 ± 0.26 b

3.71 ± 0.15 c

0.28 ± 0.09 bc

2.78 ± 0.45 e

 Natuba

4.27 ± 0.17 c

4.35 ± 0.17 b

0.45 ± 0.12 b

2.50 ± 0.09 e

EXO versus NATa

9.73**

10.54**

25.01**

26.10**

Same letters represent no significant differences by Tukey’s test (p ≤ 0.05)

**p ≤ 0.01

aIndependent sample t test comparing EXO × NAT soil groups

AMF abundance and community structure

For all study areas, the species richness, spore abundance, root colonization, total glomalin, diversity index, and dominance index were significantly different between the EXO and NAT soils (Table 2). The root colonization (p ≤ 0.01), total glomalin (p ≤ 0.01), and dominance (p ≤ 0.01) were significantly higher in the EXO soils than in the NAT soils. Within the EXO soil, we found significant differences from each studied site in AMF abundance and community structure, except for diversity index that we did not find any difference between the EXO soils. The highest root colonization (43.74%) was found in the soil samples from Esperança, while the Monteiro samples showed the highest total glomalin (7.67 mg g−1 soil) and dominance index (0.97), whereas in the NAT soils, the species richness (p ≤ 0.01), spore abundance (p ≤ 0.01), and diversity index (p ≤ 0.01) were significantly higher in the NAT soils than in the EXO soils. Within the NAT soil, we found significant differences among NAT sites for AMF abundance and community structure. The highest species richness (18 species) was found in the soil samples from Algodão de Jandaíra, while the Esperança and Monteiro samples showed the highest spore abundance (18.80 spores g−1 soil) and diversity index (2.98), respectively (Table 2).
Table 2

AMF community properties and ecological indexes for the AMF communities of exotic (EXO) and native (NAT) soils from the Brazilian tropical seasonal dry forest (mean ± SD, N = 40)

Soil type/site

Species richness

Spore abundance (spore g−1 soil)

Col. (%)

Total glomalin (mg g−1 soil)

H

D

EXO

 Algodão de Jandaíra

11.0 ± 0.40 cd

5.91 ± 0.19 d

40.24 ± 3.10 ab

5.87 ± 0.81 b

1.73 ± 0.20 d

0.93 ± 0.02 a

 Esperança

6.0 ± 0.60 f

6.12 ± 0.23 d

43.74 ± 1.11 a

6.35 ± 0.67 b

1.78 ± 0.12 d

0.95 ± 0.02 a

 Juazeirinho

7.0 ± 0.30 e

5.14 ± 0.18 e

35.33 ± 2.32 b

5.90 ± 0.98 b

1.81 ± 0.11 d

0.90 ± 0.02 b

 Monteiro

10.0 ± 0.50 c

5.12 ± 0.46 e

38.02 ± 1.56 b

7.67 ± 0.75 a

1.69 ± 0.12 d

0.97 ± 0.03 a

 Natuba

9.0 ± 0.60 d

11.70 ± 1.42 b

40.20 ± 3.14 ab

6.35 ± 0.81 ab

1.80 ± 0.10 d

0.91 ± 0.01 b

NAT

 Algodão de Jandaíra

18.0 ± 0.80 a

7.80 ± 0.12 c

6.03 ± 0.12 f

0.55 ± 0.03 d

2.20 ± 0.02 c

0.86 ± 0.01c

 Esperança

16.5 ± 0.30 b

18.80 ± 0.21 a

16.57 ± 1.12 d

0.81 ± 0.11 d

2.17 ± 0.01 c

0.85 ± 0.01 c

 Juazeirinho

17.5 ± 0.20 a

12.27 ± 0.28 b

19.10 ± 0.98 c

0.67 ± 0.09 d

2.64 ± 0.02 b

0.68 ± 0.01 f

 Monteiro

15.8 ± 0.80 b

11.38 ± 0.18 b

7.94 ± 0.67 e

0.69 ± 0.08 d

2.98 ± 0.04 a

0.76 ± 0.01 e

 Natuba

17.2 ± 0.60 a

12.02 ± 0.19 b

14.83 ± 2.12 d

0.98 ± 0.11 c

2.58 ± 0.06 b

0.79 ± 0.02 d

EXO versus NATa

12.34**

11.63**

16.32**

23.73**

12.65**

11.98**

Same letters represent no significant differences by Tukey’s test (p ≤ 0.05)

Col. (%) root colonization, H’ diversity index, D dominance index

**p ≤ 0.01

aIndependent sample t test comparing EXO × NAT soil groups

The principal component analysis (PCA) showed two well-defined clusters segregating the soil attributes and fungal communities of EXO and NAT soils (Fig. 1). PCA also indicated that the EXO soils were correlated with higher soil pH, total organic carbon (TOC), total nitrogen (TN), available phosphorus (P), total glomalin (TG), root colonization (Col), and dominance index (D), whereas NAT soils were correlated with species richness (S), spore abundance (Spore), and diversity index (H’) (Fig. 1). Available P, diversity index, spore abundance, and species richness were the main factors differing between the EXO and NAT soils. The two axes explained 88.78% of the variation present in the samples.
Fig. 1

PCA score plot of soil properties and AMF community structure for the five studied sites. Alg Algodão de Jandaíra, Esp Esperança, Jua Juazeirinho, Mon Monteiro, Nab Natuba. The points represent samples from each plot by exotic (EXO) native (NAT) soils. S species richness, H’ diversity index, Spores spore abundance, D dominance index, pH soil pH, TOC total organic carbon, P available phosphorus, TG total glomalin, TN total nitrogen, and Col root colonization

The order Glomerales, specifically the family Glomeraceae, was the most abundant in all the soil samples (Table 3). Funneliformis was statistically significantly (p ≤ 0.01) more abundant in all the studied soils, with exception of Natuba that was dominated by Claroideoglomus (60.5%) and Glomus (22.0%) in the EXO and NAT soils, respectively. The orders Diversisporales and Gigasporales were less abundant in the EXO soils at statistical significant level (p ≤ 0.01) than the NAT soils. The EXO soils showed significant lower abundance of Acaulospora (p ≤ 0.01), Quatunica (p ≤ 0.05), Gigaspora (p ≤ 0.01), Racocetra (p ≤ 0.01), and Scutellospora (p ≤ 0.05) than the NAT soils (Table 3).
Table 3

Relative abundance (%) of the arbuscular mycorrhizal fungi (based on morphological classification) of exotic (EXO) and native (NAT) soils from the Brazilian tropical seasonal dry forest (mean ± SD, N = 40)

Order

EXO vs NATa

Family

EXO vs NAT

Genera

EXO

NAT

EXO vs NAT

Alg

Esp

Jua

Mon

Nab

Alg

Esp

Jua

Mon

Nab

Archaeosporales

2.13ns

Ambisporaceae

2.13ns

Ambispora

0.0 ± 0.0 b

0.0 ± 0.0 b

0.0 ± 0.0 b

0.0 ± 0.0 b

0.0 ± 0.0 b

0.0 ± 0.0 b

0.0 ± 0.0 b

2.1 ± 0.1 a

3.0 ± 0.2 a

0.0 ± 0.0 b

2.13ns

Diversisporales

9.16**

Acaulosporaceae

9.16**

Acaulospora

0.0 ± 0.0 e

0.0 ± 0.0 e

0.0 ± 0.0 e

0.0 ± 0.0 e

0.0 ± 0.0 e

5.3 ± 0.4 b

7.5 ± 0.3 a

3.2 ± 0.2 c

0.9 ± 0.4 d

1.8 ± 0.5 d

9.16**

Gigasporales

15.33**

Dentiscutataceae

11.96**

Dentiscutata

9.0 ± 0.8 c

8.4 ± 0.6 c

7.5 ± 0.2 d

5.3 ± 0.1 e

28.0 ± 1.2 a

7.2 ± 0.4 d

9.0 ± 0.6 c

13.7 ± 1.2 b

1.4 ± 0.3 f

2.7 ± 0.2 f

10.13**

Quatunica

2.5 ± 0.1 e

2.4 ± 0.3 e

5.4 ± 0.2 b

3.1 ± 0.1 d

0.0 ± 0.0 g

0.7 ± 0.1 f

0.6 ± 0.2 f

3.9 ± 0.3 c

5.4 ± 0.3 b

11.0 ± 0.9 a

8.03*

Gigasporaceae

11.15**

Gigaspora

0.0 ± 0.0 f

0.0 ± 0.0 f

7.5 ± 0.2 b

0.0 ± 0.0 f

2.3 ± 0.1 d

0.6 ± 0.2 e

20.0 ± 3.2 a

8.3 ± 0.8 b

3.7 ± 0.5 c

7.3 ± 0.2 b

11.15**

Racocetraceae

−9.37**

Racocetra

4.9 ± 0.2 c

4.4 ± 0.2 c

9.6 ± 0.8 b

4.2 ± 0.3 c

2.3 ± 0.1 d

2.3 ± 0.1 d

2.1 ± 0.2 d

18.2 ± 1.5 a

1.8 ± 0.3 d

10.1 ± 0.5 b

9.37**

Scutellosporaceae

−8.99**

Scutellospora

0.5 ± 0.1 e

0.7 ± 0.3 e

1.7 ± 0.2 c

0.4 ± 0.1 e

0.0 ± 0.0 f

1.2 ± 0.2 d

1.0 ± 0.2 d

5.3 ± 0.4 b

6.3 ± 0.2 a

1.8 ± 0.3 c

8.99*

Glomerales

18.54**

Entrophosporaceae

14.75**

Entrophospora

0.0 ± 0.0 e

0.0 ± 0.0 e

0.0 ± 0.0 e

0.0 ± 0.0 e

0.0 ± 0.0 e

0.6 ± 0.0 d

0.6 ± 0.0 d

1.8 ± 0.1 c

0.0 ± 0.0 d

2.9 ± 0.3 a

4.15ns

Claroideoglomus

0.9 ± 0.1 f

3.0 ± 0.5 e

0.0 ± 0.0 g

7.5 ± 0.2d

60.5 ± 4.5 a

18.9 ± 3.1 b

10.5 ± 0.9 d

0.0 ± 0.0 g

22.5 ± 2.5 b

14.6 ± 0.9 c

15.11**

Glomeraceae

16.76**

Funneliformis

60.7 ± 2.1 b

57.0 ± 1.7 b

67.9 ± 3.9 ab

75.6 ± 3.5 a

4.6 ± 0.9 g

39.7 ± 1.2 c

28.3 ± 2.7 d

17.6 ± 0.9 e

39.2 ± 1.8 c

11.0 ± 1.1 f

13.72**

Glomus

5.0 ± 0.2 c

4.8 ± 0.1 c

0.4 ± 0.1 f

1.2 ± 0.3 e

0.0 ± 0.0 g

5.1 ± 0.3 c

4.9 ± 0.4 c

7.4 ± 0.2 b

3.6 ± 0.4 d

22.0 ± 1.9 a

8.76*

Rhizoglomus

18.4 ± 0.9 a

19.3 ± 1.2 a

0.0 ± 0.0 e

2.7 ± 0.2 d

2.3 ± 0.1 d

17.3 ± 2.9 a

10.5 ± 0.5 c

16.6 ± 0.8 ab

12.2 ± 1.7 b

13.8 ± 1.4 b

9.76**

Same letters represent no significant differences by Tukey’s test (p ≤ 0.05

ns not significant

**p ≤ 0.01

aIndependent sample t test comparing EXO × NAT soil groups

The AMF community present in all soil samples was composed of 29 species, most of them classified as orders Glomerales (44.8%) and Gigasporales (41.4%) (Additional file 1). The EXO soil considering all sites was composed of 18 AMF species; of those, only one from the order Diversisporales, seven from the order Gigasporales, and ten from the order Glomerales (Additional file 1), whereas the NAT soil considering all sites was composed of 29 AMF species: 18 similar to EXO soil group, plus one from the order Archaeosporales, two from the order Diversisporales, five from the order Gigasporales, and three from the order Glomerales (Additional file 1).

Discussion

Up to date, the arbuscular mycorrhizal fungal (AMF) community in the EXO soils has been poorly described when compared to other sites, such as mining areas, desertic areas, and undisturbed habitats (Silva et al. 2005; Souza et al. 2003; Panwar and Tarafdar 2006; Mergulhão et al. 2007; Silva et al. 2014; Souza et al. 2016a). To our knowledge, this is the first study assessing the soil AMF composition and diversity of EXO soils in the Brazilian tropical seasonal dry forest compared to their respective NAT soils by using morphological characterization. As revealed by our study, soil properties (e.g., soil pH, total organic carbon, total nitrogen, and available P) were higher in the EXO soils compared with the NAT soils. These results are in agreement with previous work (Soumare et al. 2015; Majewska et al. 2015) that reported higher values of soil pH and available P in the root zone of invasive exotic plants, such as Acacia senegal, Acacia seyal, Acacia albida, Eragrostis albensis, and Olpidium spp. By altering the chemical properties of the soil below their canopy, invasive plant species may alter the nutrient cycle (Follstad Shah et al. 2010) and thus may be responsible for the modification AMF community composition in the EXO soils (Zubek et al. 2016; Zubek et al. 2013).

Significant differences in the species richness, spore abundance, root colonization, total glomalin, diversity index, and dominance index were observed between the EXO and NAT soils. These findings support our hypothesis that EXO soils have lower AMF diversity than NAT soils. Several studies (e.g., Richardson et al. 2000; Hawkes et al. 2006; Rodríguez-Echeverría et al. 2009; Oehl et al. 2010; Carneiro et al. 2015; Souza et al. 2016b) also showed a lower AMF diversity in disturbed soils by biological invasion of invasive exotic species (e.g., A. senegal, A. seyal, A. albida, E. albensis, Olpidium spp., Cryptostegia madagascariensis, Sesbania virgata, P. juliflora, and Parkinsonia aculeata) in comparison with soil in natural conditions.

These results are in agreement with previous studies (Soumare et al. 2015; Ayanu et al. 2015; Zubek et al. 2016; Callaway et al. 2008; Tanner and Gange 2013) and support our hypothesis that invasive plants are associated with specific AMF species. As a consequence, P. juliflora seemed to be in advantage comparing with M. tenuiflora by profiting from beneficial AMF species (e.g., AMF from the order Glomerales) (Shah et al. 2009). The differences in AMF community structure between EXO and NAT soils were revealed by the decreased AMF species richness in all studied sites from the EXO soils and a lesser root colonization of the NAT soils. According to studies from elsewhere (Shah et al. 2009; Callaway et al. 2008; Tanner and Gange 2013), we hypothesize that three different mechanisms may be involved in the detected changes in the AMF community. First, P. juliflora usually form large monospecific plant populations, as was the case in our study, thus reducing the diversity of host-plants available to the AMF community. Consequently, this changes soil organic carbon inputs (Andrade et al. 2009; Dandan and Zhiwei 2007; Oehl et al. 2010; Jansa et al. 2014; Silva et al. 2014; Carneiro et al. 2015; Sousa et al. 2011; Souza et al. 2016a) and decreases AMF’s growth and proliferation in the absence of a diverse mycorrhizal plant community (Zubek et al. 2013).

Secondary metabolites produced by P. juliflora negatively affect native plants growth by disrupting their mutualistic associations with the unaltered AMF community (Andrade et al. 2009; Zubek et al. 2013). Studies by Stinson et al. (2006), Callaway et al. (2008) and Yuan et al. (2014) have provided evidence that invasive plant species produce metabolites that are novel for native AMF community in their introduced areas, and these secondary compounds directly limit AMF growth, spore germination, and root colonization (Callaway et al. 2008). Consequently, the most beneficial AMF (e.g., AMF species from the order Glomerales) from the native AMF community composition are favored, while the growth of the less favorable ones (e.g., AMF species from the order Diversisporales) is inhibited. Finally, the introduction of invasive exotic plant species might cause changes in soil chemical properties that may indirectly affect AMF community composition and thus contribute to a successful establishment and spread of invasive exotic plant species (Shah et al. 2009).

Previous studies revealed that Brazil is the diversification and dispersion center of species from the order Gigasporales (Goto et al. 2012; Marinho et al. 2014). In this study, we observed the same pattern for AMF communities from the NAT soils. But, we did not observe the same for the EXO soils. The biological invasion altered the composition of AMF communities in all studied EXO soils, promoting species from the order Glomerales, thus suggesting an effect of exotic plant species on the AMF community selection (Hausmann and Hawkes 2009; Jansa et al. 2014). Actually, the EXO soils showed disturbances and changes in soil properties and also showed the highest soil pH and the highest amount of total organic carbon, total nitrogen, and available P from the other NAT soils. Accordingly, the study done by Souza et al. (2016b) invasive plant species can directly affect the soil properties, which in turn can affect indirectly AMF community composition. We observed an increase in the relative abundance of AMF species from Claroideoglomus, Glomus, and Funneliformis in the EXO soils. So we can confirm that this effect was a result of the soil properties changes in response to biological invasion processes (Wetzel et al. 2014).

The low soil pH and low amounts of available phosphorus observed in the NAT soils could explain the high predominance of AMF species from Acaulospora, Gigaspora, Quatunica, Racocetra, and Scutellospora. Ramos et al. (2008) suggest that the AMF community composition in acid soils with low phosphorus availability is a result of enzymes (H+-ATPase and H+-pyrophosphatase) that act in the spore germination, and mycelium growth, improving absorption, translocation, and nutrient exchange utilization by the AMF species from the orders Archaesporales, Diversisporales, and Gigasporales.

Treseder and Turner (2007) have reported that the total glomalin is positively correlated with the primary productivity and soil organic matter, and according to the studies done by Comis (2002) and Rillig (2004), this soil protein can improve soil quality due to its positive effects improving soil aggregate and decreasing soil erosion (Gillespie et al. 2011). In our survey, we observed a significantly higher amount of total glomalin in the EXO soils. We also observed a positive correlation with total glomalin and total organic carbon and total nitrogen in these sites. Based on these results, we hypothesize that the high amounts of total glomalin in the EXO soils are involved in the changes of soil properties (e.g., soil pH, soil total carbon, and soil organic matter) as described by King (2011) and Purin and Rillig (2007). However, the potential for higher total glomalin amounts by the biological invasion process influence cannot be discarded and should be investigated in the future. Controversial results are observed in the literature showing negative effect of land use on concentration of glomalin in soil (Rillig 2004; Rillig et al. 2003; Treseder and Turner 2007).

Conclusions

Our study revealed that AMF community composition in EXO and NAT soils were dissimilar. We find evidence for changes in the original AMF community and soil properties as we have hypothesized based on the enhanced mutualism hypothesis. Differences in AMF community composition were associated with (1) differences in the dominant plant species (P. juliflora vs M. tenuiflora) and (2) changes in soil chemical factors (soil, pH, total organic carbon, total nitrogen, and available P) in exotic soils. Soil properties, total glomalin, root colonization, and dominance index were correlated with EXO soils, while species richness, number of spores, and diversity index were correlated with NAT soils. The most abundant AMF order in all the soil samples was the order Glomerales. Funneliformis, Claroideoglomus, and Glomus species were more abundant in the EXO soils than the NAT soils, but the EXO soil showed lower abundance of Acaulospora, Quatunica, Gigaspora, Racocetra, and Scutellospora species than the NAT soils. Thus, future studies should include molecular studies of the functional diversity of arbuscular mycorrhizal fungi in the EXO soils and the relation with exotic plant species.

Declarations

Acknowledgements

Special thanks to Joana Costa and Susana Rodriguez-Echeverria for valuable discussions and checking of English grammar. The authors also thank the anonymous reviewers for helpful comments, which greatly improved a previous version of the manuscript.

Authors’ contributions

We declare that all the authors made substantial contributions to the conception, design, acquisition, analysis, and interpretation of the data. All the authors participate in drafting the article, revising it critically for important intellectual content; and finally, the authors gave final approval of the version to be submitted to Ecological Processes.

Authors’ information

TAF Souza is broadly interested in the arbuscular mycorrhizal fungi (AMF) symbiosis and the morphological and molecular characterization of AMF. Specific areas of research include the following: (1) effects of biologic invasion on the arbuscular mycorrhizal diversity from the Brazilian tropical seasonal dry forest and Mediterranean forest; (2) AMF taxonomy; and (3) conservation biology of AMF. He holds a degree in Agronomy (BSc), Soil and Water Management and Conservation (M.S.), and Soil sciences (Ph.D.) from the Federal University of Paraíba, Brazil.

H. Freitas conducts research on the ecology and taxonomy of mycorrhizal fungi, plant ecology, nature conservation, plant-soil microorganism interaction, and microbial ecology. She holds a degree in Ecology (Ph.D.) from the University of Coimbra, Coimbra, Portugal, and Ecology of Ecosystems (Post-doc) from Stanford University, Stanford, USA. She is the coordinator of the Centre for Functional Ecology and has an affiliation (full-professor) with the University of Coimbra.

Competing interests

The authors declare that they have no competing interests.

Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made.

Authors’ Affiliations

(1)
Centre for Functional Ecology, Department of Life Sciences, University of Coimbra

References

  1. Agrawal AA, Kotanen PM, Mitchell CE, Power AG, Godsoe W, Klironomos J (2005) Enemy release? An experiment with congeneric plant pairs and diverse above- and belowground enemies. Ecology 86:2979–2989View ArticleGoogle Scholar
  2. Alves JJA, Araújo MA, Nascimento SS (2009) Degradação da Caatinga: uma investigação ecofisiográfica. Caatinga (Mossoró, Brasil) 22(3):126–135Google Scholar
  3. Andrade LA, Fabricante JR, Alves AS (2008) Algaroba (Prosopis juliflora (Sw.) DC.): Impactos sobre a Fitodiversidade e Estratégias de Colonização em Área Invadida na Paraíba, Brasil. Nat Conserv 6:61–67Google Scholar
  4. Andrade LA, Fabricante JR, Oliveira FX (2009) Invasão biológica por Prosopis juliflora (Sw.) DC.: impactos sobre a diversidade e a estrutura do componente arbustivo-arbóreo da caatinga no estado do Rio Grande do Norte, Brasil. Acta Bot Bras (Impresso) 23:935–943View ArticleGoogle Scholar
  5. Ayanu Y, Jintsch A, Müller-Mahn D, Rettberg S, RomanKiewicz C, Koellner T (2015) Ecosystem engineer unleashed: Prosopis juliflora threatening ecosystem services? Reg Environ Chang 15:155–167View ArticleGoogle Scholar
  6. Black CA (1965) Methods of soil analysis, part 2. In: Black CA (ed) Agronomy monograph no 9. American Society of Agronomy, MadisonGoogle Scholar
  7. Blumenthal D (2005) Interrelated causes of plant invasion. Science 310:243–244View ArticleGoogle Scholar
  8. Caifa AN, Martins FR (2007) Taxonomic identification, sampling methods, and minimum size of the tree sampled: implications and perspectives for studies in the Brazilian Atlantic Rainforest. Funct Ecosyst Commun 1:95–104Google Scholar
  9. Callaway RM, Cipolini D, Barto K, Thelen GC, Hallett SG, Prati D, Stinson K, Klironomos J (2008) Novel weapons: invasive plant suppresses fungal mutualists in American but not in its native Europe. Ecology 89:1043–1055View ArticleGoogle Scholar
  10. Carneiro MAC, Ferreira DA, Souza ED, Paulino HB, Saggin Junior OJ, Siqueira JO (2015) Arbuscular mycorrhizal fungi in soil aggregates from fields of “murundus” converted to agriculture. Pesq Agrop Brasileira Brasília 50:313–321View ArticleGoogle Scholar
  11. Comis D (2002) Glomalin: hiding place for a third of the world’s stored soil carbon. In: Agricultural Research (United States Department of Agriculture Agricultural Research Service)., pp 4–7Google Scholar
  12. Costa IR, Araújo FS (2007) Organização comunitária de um encrave de cerrado sensu stricto no bioma Caatinga, chapada do Araripe, Barbalha, Ceará. Acta Bot Bras 18:759–770View ArticleGoogle Scholar
  13. Dandan Z, Zhiwei Z (2007) Biodiversity of arbuscular mycorrhizal fungi in the hot-dry valley of the Jinsha river, southwest China. Appl Soil Ecol 37:118–128View ArticleGoogle Scholar
  14. Daubenmire RE (1968) Plant communities: a textbook of plant synecology. Haper and Row, New YorkGoogle Scholar
  15. Durigan G (2009) Estrutura e diversidade de comunidades florestais. In: Martins SV (ed). Ecologia de florestas tropicais do Brasil. Vicoça, MG: Editora UFV, 185-215.Google Scholar
  16. Follstad Shah JJ, Harner MJ, Tibbets TM (2010) Elaeagnus angustifolia elevates soil inorganic nitrogen pools in riparian ecosystems. Ecosystems 13:46–61View ArticleGoogle Scholar
  17. Fortin M, Dale MR (2005) Spatial analysis: a guide for ecologists. Cambridge University Press, CambridgeGoogle Scholar
  18. Gerdemann JW, Nicolson TH (1963) Spores of mycorrhizal Endogone species extracted from soil by wet-sieving and decanting. T Brit Mycol Soc 46:235–244View ArticleGoogle Scholar
  19. Gillespie AW, Farrill RE, Walley FL, Ross ARS, Leinweber P, Eckhardt K, Regier TZ, Blyth RIR (2011) Glomalin-related soil protein contains non-mycorrhizal-related heat-stable proteins, lipids and humic materials. Soil Biol Biochem 43(4):766–777. doi:10.1016/j.soilbio.2010.12.010 View ArticleGoogle Scholar
  20. Goto BT, Silva GA, Assis DMA, Silva DKA, Souza RG, Ferreira ACA, Jobim K, Mello CMA, Vieira HEE, Maia LC, Oehl F (2012) Intraornatosporaceae (Gigasporales), a new family with two new genera and two new species. Mycotaxon 119:117–132View ArticleGoogle Scholar
  21. Hausmann NT, Hawkes CV (2009) Plant neighborhood control of arbuscular mycorrhizal community composition. New Phytol 183:1188–1200View ArticleGoogle Scholar
  22. Hawkes CV, Belnap J, D’Antonio C, Firestone MK (2006) Arbuscular mycorrhizal assemblages in native plant roots change in the presence of invasive exotic grasses. Plant Soil 281:369–380View ArticleGoogle Scholar
  23. Hodge A, Storer K (2014) Arbuscular mycorrhizal and nitrogen: implications for individual plants through to ecosystems. Plant Soil 386:1–19View ArticleGoogle Scholar
  24. IUSS WORKING GROUP WRB (2006) World reference base for soil. World soil resources reports. n. 103. FAO, Rome, p 128Google Scholar
  25. Jansa J, Erb A, Oberholzer HR, Smilauer P, Egli S (2014) Soil and geography are more important determinants of indigenous arbuscular mycorrhizal communities than management practices in Swiss agricultural soils. Mol Ecol 23:2118–2135View ArticleGoogle Scholar
  26. Jenkins WR (1964) A rapid centrifugal flotation technique for separating nematodes from soil. Plant Dis Rep 48:692Google Scholar
  27. Johnstone IM (1986) Plant invasion windows: a time-based classification of invasion potential. Brol Rev 61:369–394Google Scholar
  28. King GM (2011) Enhancing soil carbon storage for carbon remediation: potential contributions and constraints by microbes. Trend Microbiol 19(2):75–84. doi:10.1016/j.tim.2010.11.006 View ArticleGoogle Scholar
  29. Kivlin SN, Hawkes CV (2011) Differentiating between effects of invasion and diversity: impacts of aboveground plant communities on belowground fungal communities. New Phytol 189:526–535View ArticleGoogle Scholar
  30. Kovach WL (2007) MVSP—A MultiVariate Statistical Package for windows, ver. 3.1. Kovach Computing Services, Pentraeth, WalesGoogle Scholar
  31. Kulmatiski A, Kardol P (2008) Getting plant-soil feedbacks out of the greenhouse: experimental and conceptual approaches. In: Esser K, Lüttige UE, Beyschlag W, Murata J (eds) Progress in Botany 69. Springer, Berlin, Germany, pp 449–472Google Scholar
  32. Lekberg Y, Gibbons SM, Rosendahl S, Rawsey PW (2013) Severe plant invasions can increase mycorrhizal fungal abundance and diversity. ISME J 7:1423–1433View ArticleGoogle Scholar
  33. Majewska ML, Błaszkowski J, Nobis M, Role K, Nobis A, Lakomiec D, Czachura P, Zubek S (2015) Root-inhabiting fungi in alien plant species in relation to invasion status and soil chemical properties. Symbiosis 65:101–115View ArticleGoogle Scholar
  34. Marinho F, Silva GA, Ferreira ACA, Veras JSN, Goto BT, Maia LC (2014) Bulbospora minima, a new species in the Gigasporales from semi-arid Northeast Brazil. Sydowia (in press).Google Scholar
  35. McGonigle TP, Miller MH, Evans DG, Fairchild GL, Swan JA (1990) A new method which gives an objective measure of colonization of roots by vesicular-arbuscular mycorrhizal fungi. New Phytol 115:495–501View ArticleGoogle Scholar
  36. Mergulhão ACES, Oliveira JP, Burity HA, Maia LC (2007) Potencial de infectividade de fungos micorrízicos arbusculares em áreas nativas e impactadas por mineração gesseira no semiárido brasileiro. Hoehnea 34:341–348View ArticleGoogle Scholar
  37. Oehl F, Souza FA, Sieverding E (2008) Revision of Scutellospora and description of five new genera and three new families in the arbuscular mycorrhiza-forming Glomeromycetes. Mycotaxon 106:311–360Google Scholar
  38. Oehl F, Laczko E, Bogenrieder A et al (2010) Soil type and land use intensity determine the composition of arbuscular mycorrhizal fungal communities. Soil Biol Biochem 42:724–738View ArticleGoogle Scholar
  39. Oehl F, Sieverding E, Palenzuela J, Ineichen K, Silva GA (2011) Advances in Glomeromycota taxonomy and classification. IMA Fungus 2:191–199View ArticleGoogle Scholar
  40. Okalebo JR, Gathua KW, Woomer PL (1993) Laboratory methods of plant and soil analysis: a working manual. Technical bulletin no. 1 Soil Science Society East AfricaGoogle Scholar
  41. Olsen SR, Cole CV, Watanable FS, Dean LA (1954) Estimation of available phosphorous in soils by extraction with sodium bicarbonate. US Department of Agriculture, Washigton, Circular 939Google Scholar
  42. Panwar J, Tarafdar JC (2006) Arbuscular mycorrhizal fungal dynamics under Mitragyna parvifolia (Roxb.) Korth. in Thar Desert. Appl Soil Ecol 34:200–208View ArticleGoogle Scholar
  43. Parkash V, Aggarwal A (2009) Diversity of endomycorrhizal fungi and their synergistic effect on the growth of Acacia catechu Willd. J For Sci 55:461–468Google Scholar
  44. Pegado CMA, Andrade LA, Félix LP, Pereira IM (2006) Efeitos da Invasão Biológica de algaroba - Prosopis Juliflora (Sw.) DC. sobre a composição e a estrutura do estrato arbustivo-arbóreo da caatinga no município de Monteiro-PB, Brasil. Acta Bot Bras (Impresso) 20:887–898Google Scholar
  45. Phillips JM, Hayman DS (1970) Improved procedure for clearing root and staining parasitic and vesicular-arbuscular mycorrizical fungi for rapid assessment of infection. Trans Br Mycol Soc 55:158–161View ArticleGoogle Scholar
  46. Purin S, Rillig MC (2007) The arbuscular mycorrhizal fungal protein glomalin: limitations, progress, and a new hypothesis for its function. Pedobiologia 51(2):123–130. doi:10.1016/j.pedobi.2007.03.002 View ArticleGoogle Scholar
  47. Ramos AC, Façanha AR, Feijó JA (2008) Proton (H+) flux signature for the presymbiotic development of the arbuscular mycorrhizal fungi. New Phytol 178:177–188View ArticleGoogle Scholar
  48. Redecker D, Schüßler A, Stockinger H, Stürmer SL, Morton JB, Walker C (2013) An evidence-based consensus for the classification or arbuscular mycorrhizal fungi (Glomeromycota). Mycorrhiza. doi:10.1007/s00572-013-0486-y Google Scholar
  49. Reinhart KO, Callaway RM (2006) Soil biota and invasive plants. New Phytol 170:445–457View ArticleGoogle Scholar
  50. Richardson DM, Allsopp N, D’Antonio CM, Milton SJ, Rejmánek M (2000) Plant invasions—the role of mutualisms. Biol Rev 75:65–93View ArticleGoogle Scholar
  51. Rillig MC (2004) Arbuscular mycorrhizae, glomalin, and soil aggregation. Can J Soil Sci 84(4):355–363. doi:10.4141/S04-003 View ArticleGoogle Scholar
  52. Rillig MC, Ramsey PW, Morris S, Paul EA (2003) Glomalin an arbuscular mycorrhizal fungal soil protein, responds to land-use change. Plant Soil 253(2):243–299. doi:10.1023/A:1024807820579 View ArticleGoogle Scholar
  53. Rodríguez-Echeverría S, Crisóstomo JA, Nabais C, Freitas H (2009) Belowground mutualists and the invasive ability of Acacia longifolia in coastal dunes of Portugal. Biol Invasions 11:651–661View ArticleGoogle Scholar
  54. Santos JC, Leal IR, Almeida-Cortez JS, Fernandes GW, Tabarelli M (2011) Caatinga: the scientific negligence experienced by a dry tropical forest. Trop Conservation Sci 4(3):276–286View ArticleGoogle Scholar
  55. Schenck NC, Perez Y (1987) Manual for the identification of VA mycorrhizal fungi. In: International Culture Collection of VA Mycorrhizal Fungi (INVAM), Secondth edn. University of Florida, GainesvilleGoogle Scholar
  56. Shah MA, Reshi ZA, Khasa D (2009) Arbuscular mycorrhizas: drivers or passengers of alien plant invasion. Bot Rev 75:397–417View ArticleGoogle Scholar
  57. Shanon CE, Weaver W (1949) The mathematical theory of communication. University of Illinois Press, UrbanaGoogle Scholar
  58. Sieverding E, Silva GA, Berndt R, Oehl F (2014) Rhizoglomus, a new genus of the Glomeraceae. Mycotaxon 129:373–386View ArticleGoogle Scholar
  59. Silva GA, Trufem SFB, Saggin-Júnior OJ, Maia LC (2005) Arbuscular mycorrhizal fungi in a semiarid copper mining area in Brazil. Mycorrhiza 15:47–53View ArticleGoogle Scholar
  60. Silva IRS, Mello CMA, Ferreira Neto RA, Silva DKA, Melo AL, Oehl F, Maia LO (2014) Diversity of arbuscular mycorrhizal fungi along an environmental gradient in the Brazilian semiarid. Appl Soil Ecol 84:166–175View ArticleGoogle Scholar
  61. Simpson EH (1949) Measurement of diversity. Nature 163:688View ArticleGoogle Scholar
  62. Smith SE, Read DJ (2008) Mycorrhizal symbiosis. Academic Press and Elsevier, LondonGoogle Scholar
  63. Soumare A, Manga A, Fall S, Hafidi M, Ndoye I (2015) Effects of Eucalyptus camaldulensis amendment on soil chemical properties, enzymatic activity, Acacia species growth and roots symbioses. Agrofor Syst 89:97–106View ArticleGoogle Scholar
  64. Sousa VC, Andrade LA, Bezerra FTC, Fabricante JR, Feitosa RC (2011) Avaliação populacional de Sesbania virgata (Cav.) Pers. (Fabaceae Lindl.) nas margens do rio Paraíba. Agrária (Recife Online) 6:314–320Google Scholar
  65. Souza RG, Maia LC, Sales MF, Trugem SFB (2003) Diversidade e potencial de infectividade de fungos micorrízicos arbusculares em área de Caatinga, na Região do Xingó, Estado de Alagoas, Brasil. Rev Bras Bot 26:49–60View ArticleGoogle Scholar
  66. Souza TAF, Rodríguez-Echeverría S, Andrade LA, Freitas H (2016a) Could biological invasion by Cryptostegia madagascariensis alter the composition of the arbuscular mycorrhizal fungal community in semi-arid Brazil? Acata Bot Bras 30(1):00–00. doi:10.1590/0102-33062015abb0190 Google Scholar
  67. Souza TAF, Rodriguez-Echeverría S, Andrade LA, Freitas H (2016) Arbuscular mycorrhizal fungi in Mimosa tenuiflora (Willd.) Poir from Brazilian semi-arid. Brazilian J Microbiol. http://dx.doi.org/10.1016/j.bjm.2016.01.023.
  68. Stinson KA, Campbell SA, Powell JR, Wolfe BE, Callaway RM, Thelen GC, Hallett SG, Prati D, Klironomos JN (2006) Invasive plant suppresses the growth of native tree seedlings by disrupting belowground mutualisms. PLoS Biol 4:e140View ArticleGoogle Scholar
  69. Tanner RA, Gange AC (2013) The impact of two non-native plant species on native flora performance: potential implications for habitat restoration. Plant Ecol 214:423–432View ArticleGoogle Scholar
  70. Treseder KK, Turner KM (2007) Glomalin in ecosystems. Soil Sci Soc Am J 71(4):1257–1266. doi:10.2136/sssaj.2006.0377 View ArticleGoogle Scholar
  71. Walker C, Vestberg M, Demircik F, Stockinger H, Saito M, Sawari H, Nishmura I, Schüßler A (2007) Molecular phylogeny and new taxa in the Archaeosporales (Glomeromycota): Ambispora fennica gen. sp. nov., Ambisporaceae fam. nov., and emendation of Archaeospora and Archaeosporaceae. Mycol Res 111:137–153. doi:10.1016/j.mycres.2006.11.008 View ArticleGoogle Scholar
  72. Wetzel K, Silva G, Matczinski U, Oehl F, Fester T (2014) Superior differentiation of arbuscular mycorrhizal fungal communities from till and no-till plots by morphological spore identification when compared to T-RFLP. Soil Biol Biochem 72:88–96View ArticleGoogle Scholar
  73. Wright SF, Upadhyaya A (1998) A survey of soils for aggregate stability and glomalin, a glycoprotein produced by hyphae of arbuscular mycorrhizal fungi. Plant Soil 198:97–107. doi:10.1023/A:1004347701584 View ArticleGoogle Scholar
  74. Yuan Y, Tang J, Leng D, Hu S, Yong JWH, Chen X (2014) An invasive plant promotes its arbuscular mycorrhizal symbioses and competitiveness through its secondary metabolites: indirect evidence from activated carbon. PLoS ONE 9:e97163View ArticleGoogle Scholar
  75. Zhang Q, Yang R, Tang J, Yang H, Hu S, Chen X (2010) Positive feedback between mycorrhizal fungi and plants influences plant invasion success and resistance to invasion. PLoS ONE 5:e12380. doi:10.1371/journal.pone.0012380 View ArticleGoogle Scholar
  76. Zubek S, Błaszkowski J, Seidler-Łożykowska K, Bąba W, Mleczko P (2013) Arbuscular mycorrhizal fungi abundance, species richness and composition under the monocultures of five medicinal plants. Acta Sci Pol-Hortoru 12:127–14Google Scholar
  77. Zubek S, Majewska ML, Błaszkowski J, Stefanowicz AM, Nobis M, Kapusta P (2016) Invasive plants affect arbuscular mycorrhizal fungi abundance and species richness as well as the performance of native plants grown in invaded soils. Biol Fertil Soils 52:879–893View ArticleGoogle Scholar

Copyright

© The Author(s). 2017