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Sep 27, 2017 - Community at Temperate Salt Marsh. Doongar R. ..... Henry et al. [46] .... coastal wetlands, a salt marsh with halophytes has higher methane ...
Microb Ecol https://doi.org/10.1007/s00248-017-1083-y

PLANT MICROBE INTERACTIONS

Influences of Different Halophyte Vegetation on Soil Microbial Community at Temperate Salt Marsh Doongar R. Chaudhary 1,2 & Jinhyun Kim 3 & Hojeong Kang 3

Received: 3 July 2017 / Accepted: 27 September 2017 # Springer Science+Business Media, LLC 2017

Abstract Salt marshes are transitional zone between terrestrial and aquatic ecosystems, occupied mainly by halophytic vegetation which provides numerous ecological services to coastal ecosystem. Halophyte-associated microbial community plays an important role in the adaptation of plants to adverse condition and also affected habitat characteristics. To explore the relationship between halophytes and soil microbial community, we studied the soil enzyme activities, soil microbial community structure, and functional gene abundance in halophytes- (Carex scabrifolia, Phragmites australis, and Suaeda japonica) covered and un-vegetated (mud flat) soils at Suncheon Bay, South Korea. Higher concentrations of total, Gram-positive, Gram-negative, total bacterial, and actinomycetes PLFAs (phospholipid fatty acids) were observed in the soil underneath the halophytes compared with mud flat soil and were highest in Carex soil. Halophyte-covered soils had different microbial community composition due to higher abundance of Gram-negative bacteria than mud flat soil. Similar to PLFA concentrations, the increased activities of

β-glucosidase, cellulase, phosphatase, and sulfatase enzymes were observed under halophyte soil compared to mud flat soil and Carex exhibited highest activities. The abundance of archaeal 16S rRNA, fungal ITS, and denitrifying genes (nirK, nirS, and nosZ) were not influenced by the halophytes. Abundance bacterial 16S rRNA and dissimilatory (bi)sulfite (dsrA) genes were highest in Carex-covered soil. The abundance of functional genes involved in methane cycle (mcrA and pmoA) was not affected by the halophytes. However, the ratios of mcrA/pmoA and mcrA/dsrA increased in halophytecovered soils which indicate higher methanogenesis activities. The finding of the study also suggests that halophytes had increased the microbial and enzyme activities, and played a pivotal role in shaping microbial community structure. Keywords Enzymes . Functional gene abundance . Microbial community . PLFA . Salt marsh

Introduction Electronic supplementary material The online version of this article (https://doi.org/10.1007/s00248-017-1083-y) contains supplementary material, which is available to authorized users. * Doongar R. Chaudhary [email protected] * Hojeong Kang [email protected] 1

Marine Biotechnology and Ecology Division, Central Salt and Marine Chemicals Research Institute (CSIR), Bhavnagar, Gujarat 364 002, India

2

Academy of Scientific and Innovative Research (AcSIR), CSIR, New Delhi, India

3

School of Civil and Environmental Engineering, Yonsei University, Seoul 03722, South Korea

Salt marshes are highly productive intertidal coastal ecosystems that provide habitat to salt-tolerant halophytes in an ecotone between land and sea. The global area of salt marshes is estimated from 2.2 to 40 Mha [1]. Interest in salt marshes has increased as the area declines due to sea level rise and the loss occurs of beneficial ecosystem services that salt marshes provide [2]. Coastal salt marshes withstand dynamic and complex environmental factors and therefore provide many valuable ecological functions to coastal human communities, including coastal defense, shoreline stabilization, protection against storms, wildlife conservation, and carbon and nutrient cycling [2, 3]. Halophytes are representative vegetation of salt marshes that possess special morphological, anatomical, and physiological features for adaptation to saline environments

Chaudhary D. R. et al.

[4, 5]. Salt marshes are characterized by high primary productivity and support a wide variety of vegetation [6], whereas decomposition is impeded because of waterlogged conditions and high salinity [7]. As a result, these coastal ecosystems play a crucial role in carbon (C) sequestration, contributing to climate change mitigation [8]. Although many studies report on productivity of salt marshes and the controlling variables [2, 4–6], relatively less effort is directed to understand microbially mediated decomposition processes in the rhizosphere [9–11]. The plant rhizosphere is a habitat in which microbes are directly influenced by roots, with effects due to root exudates, sloughed-off cells, lysates, litter decomposition, and exogenous enzymes that subsequently alter soil characteristics to promote the establishment of a specific rhizosphere microbial community [11–15]. The root exudates of halophytes provide C and energy to the microorganisms, and the quantity and quality of exudates released in soil depend on the plant species, growth stage and metabolism, and microbial response to changes in the nutritional condition of soil [16–18]. Extracellular enzyme activities are a useful indicator of soil quality and are involved in the hydrolysis of complex substrates during decomposition [12, 15]. In coastal ecosystems, halophyte-associated microbial communities and enzyme activities are influenced by variable salinity and waterlogging, providing a unique opportunity for investigation. The production of organic matter by halophytes is high, and the decay of litter or compounds released through roots in conjunction with waterlogging results in oxygen depletion in the sediments. Nitrate and sulfate are two primary ions that act as terminal electron acceptors to alleviate anaerobic respiration in sediments [19, 20]. The important enzyme for sulfate reduction is dissimilatory sulfite reductase (dsr). This enzyme catalyzes the reduction of sulfite to sulfide, which is an essential step in the anaerobic sulfate respiration pathway encoded by the dsrAB gene with α- and β-subunits (referred to as dsrA and dsrB, respectively) [21–23]. The denitrification processes (nitrite and nitrous oxide reduction) are catalyzed by nitrite and nitrous oxide reductase, respectively. Copper and a cytochrome cd1-containing nitrite reductase encoded by the genes nirK and nirS, respectively, catalyze nitrite reduction [24]. Nitrous oxide reductase (nosZ) is another gene also associated with the denitrification process [25]. The production of CH4 in salt marshes is an anaerobic microbial process conducted by methanogens. The mcrA gene is found in all methanogens and encodes the α-subunit of methyl-coenzyme M reductase involved in methanogenesis [26]. Aerobic methane-oxidizing bacteria (methanotrophs) conduct CH4 oxidation. The pmoA gene encoding the α-subunit of the particulate methane monooxygenase enzyme can be used as a functional marker for methanotrophic bacteria [27]. Information on these genes is of particular importance, because the final products CO2, CH 4 , and N 2 O are strong greenhouse gases. Targeted

functional gene approaches that examine the responses of specific genes related to nutrient cycling will help to understand fine-scale community responses, despite the overall similarity in composition of soil microbial communities [25]. Globally, many coastal marshes are experiencing rapid changes in vegetation composition. For examples, Phragmites invasion receives much attention in the USA [28], whereas Spartina expansion is of concern in China [29, 30]. Suncheon Bay was registered as a wetland of international importance by the Ramsar Convention in 2006 and consists of the largest high-density colonies of Phragmites in Korea, with the acreage continuing to increase. Such rapid changes in vegetation composition are expected to result in changes in microbial community composition and activities; however, detailed information remains unavailable. We hypothesized that different halophytes would change the microbial community composition and functional gene(s) abundance compared with non-vegetated mudflats. In this investigation, we evaluated the effects of different halophytes that commonly occur (Carex scabrifolia, Phragmites australis, and Suaeda japonica) in salt marshes at Suncheon Bay, South Korea, on the soil microbial community (phospholipid fatty acids, PLFAs), extracellular enzyme activities (β-glucosidase, cellulase, phosphatase, sulfatase, N-acetyl-β-glucosaminidase, and aminopeptidase), and abundance of bacteria, archaea, fungi, and functional genes (mcrA, pmoA, dsrA, nirK, nirS, and nosZ) related to greenhouse gas emissions. These effects were compared with those in an un-vegetated mudflat soil.

Materials and Methods Research Site and Soil Sampling The research site included the salt marshes at Suncheon Bay, South Korea (Fig. 1). Suncheon Bay is located on the southern coast of the Korean Peninsula (34°84′ N, 127°45′ E). The total area of mudflats with and without vegetation is 5.4 and 21.6 km2, respectively. Mudflats are flooded with sea water by the tide twice a day. Soil texture of mudflats is mostly loam in this area. The Suncheon Bay area has a temperate climate. Mean annual air temperature is 14.3 °C, with the lowest winter temperature of − 0.8 °C and the highest summer temperature of 28.9 °C. Mean annual precipitation is 1439 mm. Carex scabrifolia (hereafter Carex), Phragmites australis (hereafter Phragmites), and Suaeda japonica (hereafter Suaeda) dominate the study site. Soil samples were collected in July 2016 from four quadrats (1 × 1 m) (four replicates) from uniform, monospecific populations of individual halophyte species (n = 12). Ten meters separated replicate quadrats. From each quadrat, soil samples (0–20 cm) were collected using a trowel under the canopy (near the root zone area) of

Influences of Different Halophyte Vegetation on Soil Microbial Community at Temperate Salt Marsh Fig. 1 Map of location of Suncheon Bay (34°84′ N, 127°45′ E)

four to five individual plants (subsamples) for one composite sample per quadrat. Similarly, four mudflat soil samples (without any halophyte vegetation) were also collected (n = 4). All instruments were sterilized using alcohol and distilled water before and between soil samplings. The soil samples were kept in an ice box and returned to the laboratory. Soil samples were divided into two subsamples: one set of subsamples was maintained at 4 °C for enzyme and PLFA analyses, and the other set of subsamples was maintained at − 80 °C for molecular analysis. Soil Analysis Soil moisture content was determined after drying at 105 °C for 24 h in a hot air oven. The pH and electrical conductivity (EC) were measured in a 1:2 (soil:water) suspension. Organic matter was determined by loss on ignition at 600 °C in a muffle furnace. Dissolved organic carbon was extracted from soil (2 g) with water (20 mL), filtered, and then measured with a TOC analyzer (TOCVCHP; Shimadzu, Kyoto, Japan). To measure soil microbial community composition, fatty acids from soil were extracted following the methods of Bardgett et al. [31] and Frostegård et al. [32] using chloroform, methanol, and citrate buffer (1:2:0.8 v/v) followed by fractionation (neutral-, glyco-, and phospho-lipid) using a silicic column. Phospholipids were esterified in alkaline methanol and then analyzed using a gas chromatography (Agilent 6850; Agilent Technologies, Inc.) equipped with an ultra 2 capillary column, a flame ionization detector, and MI System version 6.0 (MIDI Inc., Newark, DE, USA) using the TSBA6 method. An internal standard of methyl nonadecanoate (19:0) was used for calculation of fatty acid concentrations. The concentrations of signature biomarker fatty acids were used for the following microbial taxonomic groups [33, 34]: Gram-positive bacteria (i15:0, a15:0, i16:0, a16:0, i17:0,

a17:0), Gram-negative bacteria (16:1ω5, 16:1ω7, 17:1ω9, 18:1ω7, cy17:0, cy19:0), fungi (18:2ω6,9), and actinomycetes (10Me16:0, 10Me17:0, 10Me18:0). The absolute concentration (nmol g−1 soil) and relative abundance (molar %) were calculated for these signature PLFA biomarkers. Extracellular enzyme activities in soil samples were measured using methylumbelliferyl (MUF)-linked substrates [35]. MUF-labeled β-glucoside, β-D-cellobioside, phosphate, sulfate, N-acetyl β-glucoside, and L-leucine-7amino-4-methylcoumarin hydrochloride were used to measure the activities of β-glucosidase, cellulase, phosphatase, sulfatase, N-acetyl glucosaminidase, and aminopeptidase, respectively. Soil enzymes were extracted with acetate butter (50 mM, pH 5.0), and the extract was mixed with MUF-linked substrate solution for hydrolases. The amount of MUF substrate consumed was measured using a fluorescent assay (excitation: 355 nm; emission: 460 nm; FLUO-star OPTIMA, BMG LABTECH, Germany) after incubation at 25 °C. Soil microbial DNA was extracted from 400 mg of soil sample using a PowerSoil® DNA Isolation Kit (MOBIO, CA, USA). To estimate the abundances of specific groups of microbes and functional genes, real-time PCR was performed using the SYBR Green Real-time PCR Master MIX (Toyobo Co., Japan) as a detector. Target genes were bacterial 16S rRNA, archaeal 16S rRNA, fungal ITS, mcrA, pmoA, dsrA, nirS, nirK, and nosZ. The primers for each gene are provided in the references in Table 1. Fifty cycles of three-step PCR were used for amplification: denaturation at 95 °C for 25 s, annealing at 50, 52, 50, 64.5, 55, 53, 65, 58, and 56 °C (for bacterial 16S rRNA, archaeal 16S rRNA, fungal ITS, mcrA, pmoA, dsrA, nirS, nirK, and nosZ, respectively) for 25 s, and extension at 72 °C for 25 s. The CFX96TM Real-time PCR Detection System (BIO-RAD, USA) performed the reaction. The total volume of each reaction mixture was

Chaudhary D. R. et al. Table 1

The references for primers and PCR protocols

Target gene Bacterial 16S rRNA

Primer

Reference

accounting for Type III error in which the fixed effects sum to zero.

Forward

Lane [36]

Reverse

Lane [36]

Results

Archaeal 16S rRNA

Forward Reverse

Takai and Horikochi [37] Takai and Horikochi [37]

Soil Characteristics

Fungal ITS

Forward Reverse

Gardes and Bruns [38] White et al. [39]

Forward

Steinberg and Regan [40]

mcrA

Reverse

Hales et al. [41]

pmoA

Forward

Holmes et al. [42]

Reverse

Costello and Lidstrom [43]

dsrA

Forward

Kondo et al. [21]

Reverse

Kondo et al. [21]

Forward

Liu et al. [44]

nirS nirK nosZ

Reverse

Braker et al. [45]

Forward

Henry et al. [46]

Reverse

Braker et al. [45]

Forward

Rich et al. [47]

Reverse

Rich et al. [47]

20 μL, with 6.4 μL of distilled water, 10 μL of SYBR Green Master Mix, 0.8 μL of each forward and reverse primer (10 μM), and 2 μL of extracted soil DNA solution. Three independent real-time PCR assays were performed on each DNA sample for replications. The gene copy number was calculated by the standard curve method. The standard curve was deduced using 10-fold diluted series of plasmids containing each target gene.

Statistical Analyses All quantitative data were subjected to statistical analyses using SigmaPlot 12.0 (Systat Software, Inc., San Jose, CA, USA) and PC-ORD [48] software. The influence of halophyte on soil PLFA concentrations, soil enzyme activities, and abundance of functional genes were analyzed with one-way ANOVA followed by Tukey’s post-hoc tests (Tukey’s HSD test; p ≤ 0.05) using SigmaPlot 12.0 (Tables S1 and S2). All data sets were tested for a normal distribution using the Shapiro-Wilk test (SigmaPlot 12.0) before analysis of variance. The PLFAs (molar %) were used for non-metric multidimensional scaling (NMS) analysis using PC-ORD software with a Bray-Curtis distance matrix. A second matrix was prepared with summed values of taxonomic groups to construct a joint plot overlaid on NMS. The variability in soil microbial communities was assessed using the PerMANOVA procedure

Soil characteristics showed considerable variability, particularly organic matter, dissolved organic carbon, and moisture content (Table 2). The pH and EC ranged from 7.56 to 7.60 and from 4.15 to 5.59 (dS m−1), respectively. Organic matter content was higher under halophyte-covered soil than under mudflat soil. Phragmites soil had the highest dissolved organic carbon content, followed by Carex and Suaeda, with the lowest content in mudflat soil. The moisture content was highest in the soil beneath Carex and lowest in the mudflat soil. Soil Microbial Community Composition and Structure There were significant influence of halophytes on the concentrations of total PLFAs (F3,12 = 5.38, p = 0.01), Gram-positive (F3,12 = 6.028, p = 0.01), Gram-negative (F3,12 = 6.68, p = 0.007), total bacterial (F3,12 = 9.23, p = 0.002), and actinomycetes (F3,12 = 4.31, p = 0.02) PLFAs. The highest concentrations of PLFAs for the total, Gram-positive, Gram-negative, total bacteria, and actinomycetes were observed in the soil underneath Carex, which were comparable with those in other halophyte-covered soils but significantly higher than those in the mudflat soil (Table 3). However, halophytes did not significantly affect concentrations of the fungal biomarker PLFA. The NMS ordination of molar percent PLFAs resulted in a two-dimensional solution, and both ordination axes explained 89.3% of the total variability in soil microbial community structure (Fig. 2). Gram-positive (r = − 0.54, p < 0.01), Gram-negative (r = − 0.68, p < 0.01), and total bacteria (r = − 0.77, p < 0.01) were significantly and negatively correlated with axis 2, whereas actinomycetes were positively (r = 0.64, p < 0.01) correlated with axis 2 and negatively (r = − 0.59, p < 0.01) correlated with axis 1. Halophytes had a significant influence on soil microbial communities (PerMANOVA, pseudo-F = 1.95, p = 0.017). Based on scores of axis 1, the soil microbial community of Phragmites was significantly differentiated from the communities under Suaeda and in mudflat soil. The scores of axis 2 differentiated the soil microbial community of mudflat soil from those in Carex- and Suaeda-covered soils. The abundance of PLFA biomarkers for the microbial taxonomic groups is depicted in Fig. 3 (all biomarker PLFAs kept at 100%). Among different microbial taxonomic groups, halophytes significantly affected the abundance of Gram-

Influences of Different Halophyte Vegetation on Soil Microbial Community at Temperate Salt Marsh Table 2 Characteristics of soils of Suncheon Bay. [Different letters denote significant differences (Tukey’s HSD test) at p ≤ 0.05]

Soil

EC(1:2) (dS m−1)

pH (1:2)

Organic matter (%)

DOC(mg g−1)

Moisture (%)

Mudflat

4.56

7.60

5.99b

0.08c

66.28b

Carex Phragmites

4.15 4.78

7.57 7.56

6.38ab 6.60a

0.21b 0.31a

86.77a 76.45ab

Suaeda

5.59

7.56

6.58a

0.16b

74.44ab

negative bacteria (F3,12 = 5.34, p = 0.01), which increased in halophyte-covered soils compared with that in mudflat soil.

Soil Extracellular Enzyme Activities Halophytes significantly influenced soil extracellular enzyme activities compared with those in mudflat soil (Table 4). The activities of β-glucosidase (F3,12 = 4.40, p = 0.02), cellulase (F3,12 = 4.06, p = 0.03), phosphatase (F3,12 = 3.62, p = 0.045), and sulfatase (F3,12 = 4.57, p = 0.02) increased in halophytecovered soil compared with those in mudflat soil; however, halophytes did not significantly affect N-hydrolyzing enzymes (N-acetyl-β-glucosaminidase and aminopeptidase). Similar to PLFA concentrations, the highest activities of βglucosidase, phosphatase, sulfatase, and cellulase occurred in soils covered with Carex, followed by Suaeda, Phragmites, and mudflat soils.

observed in Carex-covered soil followed by mudflat, Phragmites, and Suaeda soils (Table 5). We selected two functional genes (mcrA and pmoA) involved in the methane cycle. The abundance of methanogens (methyl-coenzyme M reductase enzyme, mcrA) was similar in all soils, with relatively higher abundance under Carex and Phragmites and the lowest abundance in mudflat soil. Halophytes significantly affected the abundance of methaneoxidizing bacteria (particulate methane monooxygenase enzyme, pmoA) (F3,12 = 4.75, p = 0.02), and the highest abundance was observed under Carex, followed by mudflat, Phragmites, and Suaeda soils (Table 5). Halophytes also significantly influenced the relative proportions of methanerelated functional genes, mcrA/pmoA (F3,12 = 3.97, p = 0.04) and mcrA/dsrA (F3,12 = 4.91, p = 0.02), with the lowest proportions observed in mudflat soil (Fig. 4).

Discussion Gene Abundances Soil Microbial Community Structure The amount of bacterial 16S rRNA ranged from 9.6 × 108 to 15.2 × 108 copies per gram of soil, and the highest abundance was observed in Carex-covered soil, whereas the lowest was observed under Suaeda, which was significantly different from other soils (F3,12 = 3.73, p = 0.04; Table 5). Halophytes did not significantly affect copy number for archaeal 16S rRNA (4.0 × 108 to 5.19 × 108) or fungal ITS (1.7 × 107 to 4.3 × 107) genes. Similarly, the halophytes did not strongly influence the abundance of denitrifying genes nirK (2.8 × 108 to 4.3 × 108), nirS (5.1 × 109 to 8.1 × 109), and nosZ (6.1 × 108 to 11.6 × 108). Halophytes significantly affected the abundance of the dissimilatory (bi)sulfite (dsrA) gene (F3,12 = 7.82, p = 0.004), and the highest copy number was

Table 3 Concentrations of PLFAs (nmol g−1) in soils. [Different letters denote significant differences (Tukey’s HSD test) at p ≤ 0.05]

Soil microbial biomass and activity reveal the size of the microbial population involved in biological activities and nutrient cycling processes. Soil microbial activity can be measured using both culture-dependent and culture-independent methods. We used PLFA analysis to determine microbial biomass and microbial community structure, which is a cultureindependent technique and widely used for plant-soil-microbe interaction studies [33, 34]. In the present study, the hypothesis was that halophytes would directly influence soil microbial communities [49, 50]. Based on the PLFA analysis, in the soils beneath the halophytes, total, Gram-positive, Gram-negative, total bacteria, and actinomycete biomarker PLFA

Soil

Total PLFAs

Gram-positive

Gram-negative

Total bacteria

Fungi

Actinomycetes

Mudflat Carex Phragmites Suaeda

67.11b 86.72a 80.74ab 82.71a

11.04b 14.72a 13.91a 13.69ab

21.20b 28.95a 26.05ab 28.39a

32.25b 43.67a 39.95a 42.08a

1.31 1.85 1.75 1.88

4.87b 6.22a 5.81ab 5.00ab

Chaudhary D. R. et al. Fig. 2 NMS representation of soil sample distance based on molar % of PLFAs extracted from halophyte-covered soils and mudflat soil. Error bars indicate standard error of the mean. (GM+ : Gram-positive; GM-: Gramnegative; F/B: ratio of fungal to bacterial PLFAs)

concentrations increased compared with those in mudflat (unvegetated) soil (Table 3). The PLFA concentration is an indicator of microbial biomass [33]. The total PLFA concentration in halophyte-covered soil was significantly higher than that in mudflat soil, which was attributed to increased organic matter and dissolved organic carbon contents, and therefore increased microbial populations and activity [10, 11, 51]. The higher amounts of bacteria and actinomycetes in halophyte-

Fig. 3 Relative abundance of different microbial taxonomic groups in halophyte-covered soils and mudflat soil. Different letters denote significant differences (Tukey’s HSD test) at p ≤ 0.05

covered soil were well correlated with organic matter and dissolved organic carbon released by halophytes as root exudates [49, 50, 52], as revealed in Table 2. These observations are consistent with those of earlier studies that found that halophytes altered soil characteristics by affecting nutrient mineralization and litter decay [11, 15, 51, 52]. The biomass of Gram-negative bacteria was greater than that of Grampositive bacteria, and although halophytes did not affect their

Influences of Different Halophyte Vegetation on Soil Microbial Community at Temperate Salt Marsh Table 4 Enzyme activities (nmol g−1 soil min−1) in soils of Suncheon Bay. [Different letters denote significant differences (Tukey’s HSD test) at p ≤ 0.05]

Table 5 Gene abundance (× 107 copy number g−1 soil) in soils of Suncheon Bay. [Different letters denote significant differences (Tukey’s HSD test) at p ≤ 0.05]

Soil

βglucosidase

Cellulase

Phosphatase

Sulfatase

N-acetyl-βglucosaminidase

Aminopeptidase

Mudflat

1.18b

0.93b

4.53b

0.29b

0.88

9.08

Carex

1.48a

1.03a

5.17a

0.34a

1.01

11.14

Phragmites Suaeda

1.37ab 1.40ab

0.96ab 0.98ab

4.93ab 5.04ab

0.31ab 0.33ab

0.97 1.00

10.33 10.31

Soil

mcrA

pmoA

dsrA

nirK

nirS

nosZ

Bacterial

Archaeal

Fungal

Mudflat

0.64

7.89ab

24.99a

29.83

724.35

116.62

104.57ab

43.00

1.34

Carex

1.35

10.55a

28.24a

43.82

698.97

99.93

152.50a

51.99

4.26

Phragmites Suaeda

1.14 0.72

6.99ab 5.85b

19.77ab 15.03b

31.60 28.91

813.59 517.27

75.22 61.19

114.57ab 96.50b

44.68 40.02

2.15 1.70

ratio (Gram-negative to Gram-positive), this ratio was higher in halophyte-covered soil than in mudflat soil. Similar to our observations, others also report that Gram-negative bacteria contribute a large proportion to bacterial biomass [11, 51, 52], and studies also show that root exudates are preferentially used by Gram-negative bacteria [10, 53]. Our NMS analysis of PLFA profiles confirmed that soil microbial communities shifted under halophyte-covered soil compared with those in mudflat soil (Fig. 2). This shift in microbial community structure was attributed to increased abundance of Gram-negative bacteria in halophyte-covered soil (particularly under Suaeda) compared with that in mudflat soil (Fig. 3). The NMS clearly differentiated the halophytecovered soils from the mudflat soil. Furthermore, the soil microbial communities under Phragmites and Carex were similar, but the community under Suaeda was different from that of Phragmites. Phragmites and Carex have an adventitious root system, whereas Suaeda has a tap root system, and these differences affect rhizodeposition and the distribution in soil. Biochemical constituents of root exudates are plant speciesspecific, and although these compounds may occur in similar environmental conditions, they have different effects on the composition of the soil microbial community [16, 17].

concentrations supporting such a linkage (Table 3). These types of differences in enzyme activities have previously been reported in halophytes and in field crops [10, 11]. Therefore, with increases in root growth and branching, the synthesis of these enzymes may result from increased substrate availability in the form of root exudates, decomposition processes, and increases in microbial biomass.

Methanogens, Methanotrophs, and Sulfate Reducers Methane production is a primary process in the anaerobic carbon cycle in coastal wetlands, and the methanogens (mcrA), methanotrophs (pmoA), and sulfate reducers (dsrA) are the primary microbes involved in the process. Methanogens produce methane, methanotrophs oxidize the

Soil Extracellular Enzyme Activities Soil enzyme activities provide information about biochemical processes and are good indicators of soil quality and microbial functions. The activities of extracellular enzymes (βglucosidase, cellulase, phosphatase, and sulfatase) were higher in halophyte-covered soil than in mudflat soil, reflecting the positive effect of halophyte roots (Table 4). The increase in activities might be linked to increased substrate availability and microbial activity [10], with PLFA

Fig. 4 Ratios of functional gene copies of mcrA/pmoA and mcrA/dsrA in halophyte-covered soils and mudflat soil. Different letters denote significant differences (Tukey’s HSD test) at p ≤ 0.05

Chaudhary D. R. et al.

methane, and sulfate reducers compete with methanogens for acetate, a primary carbon source, resulting in competitive inhibition [54, 55]. The relative abundance of these three microorganisms, and the ratios mcrA/pmoA and mcrA/dsrA varied according to soils (Fig. 4), which might reflect the influence of halophytes. According to a previous study conducted in temperate coastal wetlands, a salt marsh with halophytes has higher methane emissions than a mudflat without halophytes [30]. We found the gene copy ratio of mcrA/dsrA was significantly higher in the halophyte-covered soil than in the mudflat soil (Fig. 4), and a higher mcrA/dsrA ratio indicates a relatively higher abundance of methanogens and lower abundance of sulfate reducers. Thus, the proportion of methanogens affected by competitive inhibition was reduced, and methanogenesis activity should increase. The higher mcrA/dsrA ratio in halophyte-covered soil than in mudflat soil might be explained by the increase in DOC under halophytes (Table 2). Zeleke et al. [29] suggested that excess carbon availability might reduce the competitive inhibition between methanogens and sulfate reducers. Therefore, sulfate reducers can outcompete methanogens when carbon availability is limited [56]; however, in the present study, methanogens likely escaped from sulfate reducer inhibition in halophyte-covered soil because of the higher DOC concentrations, which resulted in the higher mcrA/dsrA ratio. The gene copy ratio of mcrA/pmoAwas significantly higher in the halophyte-covered soils than in the mudflat soil, which is consistent with previous studies conducted in salt marshes [30]. Previous studies show that methane emissions and mcrA/ pmoA ratios are positively correlated [57]. Abundance of functional gene(s) provides information on in- situ activities of functional microbial groups. For mcrA/pmoA, a high ratio indicates relatively higher abundance of methanogens and lower abundance of methanotrophs. Therefore, in soils with a high mcrA/pmoA ratio, methanogenesis increases and methane oxidation decreases. Overall, the present study indicated that invasion of and shifts in vegetation induced distinctive changes in the microbial community, leading to much higher activities. Particularly, the relative abundance of methanogens and activities of enzymes involved in organic matter decomposition increased, which might have implications for the longterm sequestration of carbon in salt marshes.

strongly influenced by halophyte plant species. We also observed increased activities of enzymes in halophyte-covered soils. The differences in enzyme activities and soil microbial communities among halophyte species and in mudflat soil could be closely related with root exudates and plant litters that differ among plant species in biochemical composition. The ratio of relative abundance of methanogens (mcrA) to methanotrophs (pmoA) was higher in halophyte-covered soils than in mudflat soil, indicating that more methanogenesis and less methane oxidation might occur. This result was also supported by the relatively higher abundance of methanogens (mcrA) and lower abundance of sulfate reducers (dsrA) in halophyte-covered soils than in mudflat soil. Acknowledgements This work was supported by Brainpool fellowship of the Korean Federation of Science and Technology (KOFST). This paper was studied with the support of the Korean Ministry of Science, ICT and Future Planning (MSIP) and National Research Foundation of Korea (20110030040 and 2015K2A2A2002194), and Korean Ministry of Education (SGER; 2016R1D1A1A02937049).

References 1.

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Conclusions In the present study, soil microbial community structure was analyzed in halophyte-covered soils and un-vegetated mudflat soil in a salt marsh of Suncheon Bay, South Korea. The halophytes affected a shift in soil microbial community structure. Halophytes play an important role in shaping microbial community composition because belowground processes are

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