Soil microbial community structure and function relationships: A heat stress experiment

Applied Soil Ecology 86 (2014) 121–130

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Soil microbial community structure and function relationships: A heat stress experiment Wassila Riah-Anglet a, *, Isabelle Trinsoutrot-Gattin a , Fabrice Martin-Laurent b , Emilie Laroche-Ajzenberg a , Marie-Paule Norini a , Xavier Latour c, Karine Laval a a b c

Unité Agri'Terr, Esitpa, 3 rue du Tronquet 76134, Mont Saint Aignan, France INRA, UMR Agroécologie, Écologie des Communautés et Durabilité Systèmes Agricoles, Centre de Dijon, 17 rue Sully BP 86510, 21065 Dijon cedex, France Laboratoire de Microbiologie Signaux et Microenvironnement (LMSM EA 4312) Normandie Université - Université de Rouen - IUT Evreux, Evreux, France

A R T I C L E I N F O

A B S T R A C T

Article history: Received 5 May 2014 Received in revised form 25 September 2014 Accepted 1 October 2014 Available online xxx

Links between the microbial community structure and soil functions are unclear. The study of these relationships requires the development of highly specific experimental approaches. In this work, the soil microbial community structure and function relationship was evaluated in relation to heat stress in a soil microcosm incubated at 17
C and 50
C. We selected a luvisol that included two land uses. Samples were taken from the soil of a long-term (>10 year) arable cropping plot (CC) and a permanent grassland (PG) (>25 years). The soil functions were evaluated by measuring the enzyme activities, including cellulase, N-acetyl-glucosaminidase, b-glucosidase, xylanase and dehydrogenase. The total microbial biomass was assayed by the quantification of the total DNA extracted from the microcosm soils. The abundance of total bacterial and fungal communities and different bacterial taxa were measured by qPCR rRNA genes. For both soil types, heat stress induced changes in the microbial community structure and soil functions. In most cases, the results yielded effects following heat treatment. All of the enzymes were inhibited except xylanase. Heat stress significantly reduced the total microbial biomass and fungal abundance in the soils. The abundance of the total bacterial community was not affected by heat stress. In the two soils, the dominant taxa were Actinobacteria (13–40%) and Bacteroidetes (14–32%), while Planctomycetes and Gammaproteobacteria exhibited lower abundance (0–3%). Changes in the microbial community structure and changes in the functions were correlated; the correlation was positive in the PG soil and negative in the CC soil. The changes in the CC soil structural and functional state were greater than of those observed in PG soil. Our initial hypothesis was confirmed, indeed, grassland soil is more resistant to drastic stress due to its highly abundant and highly diversified microbial community. These results represent a contribution to the understanding of soil microbial community structure and functions relationships. ã 2014 Elsevier B.V. All rights reserved.

Keywords: Microbial diversity Bacterial taxa Enzymatic activities Heat stress Soil land use

1. Introduction Soils are considered the primary reservoirs of biodiversity (Jangid et al., 2010). Among this biodiversity, microflora represents a considerable fraction, which is highly diversified (Torsvik et al., 1990; Allison and Martiny, 2008). A large number of studies have documented how the microbial community structure, or the different types of microorganisms and their abundances (Fuhrman, 2009), constitute an essential element

* Corresponding author. Tel.: +33 232829198. E-mail address: [email protected] (W. Riah-Anglet). http://dx.doi.org/10.1016/j.apsoil.2014.10.001 0929-1393/ ã 2014 Elsevier B.V. All rights reserved.

for understanding the impacts of environmental and anthropogenic perturbations on soil functioning (Nannipieri et al., 2003). However, the close relationship between the structure of microbial communities and soil functions remain poorly understood because of functional redundancy of microbial communities. Yin et al. (2000) defined the functional redundancy as the potential for multiple species to be able to perform the same function and thus, changes in the microbial community structure do not necessarily lead to a change in soil functioning (Chapin et al., 1997). Microbial species that appear after an environmental stress or disruption but harbor the same ability to perform a function might (i) not have the same growth rate or competitive ability compared with the original community

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members (Petterson and Bååth, 2003), or (ii) not perform functions with the same efficiency and/or generate the same metabolic by-products, or (iii) directly or indirectly influence the activity of other populations in the community. The small numbers of studies addressing the link between microbial community structure and function have shown that the manipulation of species diversity is a key to establishing empirical relationships (Griffiths et al., 2000). Two types of experimental approaches could be identified from the literature for microbial community manipulations: (i) the destructive approach and (ii) the constructive approach. The destructive approach reduces the microbial diversity by creating realistic rates of diversity but has the disadvantage of selecting the most abundant microorganisms (Dimitriu et al., 2010). Other authors have used more destructive methods, such as repeated fumigations to progressively decrease microbial diversity (Dickens and Anderson, 1999; Griffiths et al., 2000) or applications of different stress (including biocides) to kill specific microorganisms. The second approach involves the reconstitution of microbial communities through the establishment of a combination of culturable microorganisms (Liebich et al., 2007; Griffiths et al., 2008). The two approaches cited here were biased in the selection of diversity and composition, but are helpful methods for understanding functions of microbial community structural changes or alteration (Griffiths and Philippot, 2013; Bell et al., 2005). These studies have demonstrated that a decrease in the diversity is not necessarily followed by loss of function. Indeed some authors argued that community structure is positively correlated with functions (Chesson, 2000), and other authors argued that redundancy is similar to an “insurance policy” against functional loss and that species loss is unlikely to impact the function (Yachi and Loreau, 1999). This research aimed to examine the relationship between the microbial community structure and soil functions. As defined by Fuhrman, (2009), here we consider microbial structure simply to include the number of different fungal and bacterial species, and their relative abundance in soil microflora. We opted to apply a destructive approach by subjecting soil to heat stress to simplify the microbial community structure and observe consequences on the resultant functions. We studied two soils with the same edaphic characteristics and great difference in microbial community structure: (i) a permanent grassland (PG) and (ii) a conventional crop (CC). Several authors have already demonstrated that the initial microbial community structure was different between PG soil and CC soil (Jackson et al., 2003; Bissett et al., 2011). Moreover, PG has been described as resilient soil because of the very abundant and highly diversified microbial communities (Jangid et al., 2008). Thus, we hypothesized that the soil function responses to the heat stress would be different in PG soil and in CC soil. Soil functions were studied by evaluating soil enzymatic activities that are considered indicative of specific biochemical reactions of the entire soil microbial community (Nannipieri et al., 2002). Soil microbial community structures were examined by the quantification (qPCR) of the total bacterial and fungal biomasses and ten bacterial taxa.

2. Materials and methods 2.1. Soil sampling The soil samples were collected in the experimental site of Yvetôt (northwestern France). The climate was temperate oceanic (+10
C mean annual temperature and 850 mm annual precipitation). The soils characteristics, determined by the Soil Analysis Laboratory (LAS, INRA, Arras, France), are presented in Table 1. The soil is defined as a silty soil and classified as a luvisol. The soils were sampled according to two land uses, including a long-term arable conventional cropping plot (>10 year, wheat, maize, flax or beet crop rotation) (CC) and a permanent grassland (PG) (>25 years, dominant ray-grass species). The soil samples were collected by using an auger at a depth of 0–20 cm by pooling five subsamples. Prior to use, the samples were sieved to 2 mm. The water content was determined after drying 10 g of soil at 105
C in an oven for 24 h. 2.2. Microcosm preparation and heat stress treatments Microcosms were prepared in 20 cm3 glass jars. Each microcosm contained 200 g of freshly sieved soil. Several temperatures were tested to choose the appropriate heat stress that most impacted the soil’s microorganisms (results not shown). Heat stress at 50
C and 60
C similarly impacted the microbial communities and the enzymatic activities. A temperature of 50
C was selected for this study. For controls without heat stress, the soils were incubated at 17
C, which corresponded to the soil temperature at the time of sampling. The heat stress treatments included PG and CC soil microcosms incubated at 17
C and 50
C in a ventilated oven for 28 days. The soils were used directly after sieving without stabilization period. A daily weight of the microcosms allowed for controlling the moisture of the soils. The water losses were replaced using sterile distilled water. Three replicates were performed for each soil at each sampling time (5, 14, 21 and 28 days) and at each temperature treatment, for a total of 48 samples (24 destructive microcosms for the PG plot and 24 destructive microcosms for the CC plot). T = 0 corresponded to 5 days after the start of the incubation to avoid including the disturbances of soil microbial community caused by the establishment of the experiment. 2.3. Total DNA extraction and real-time PCR quantification (qPCR) For the 48 replicates, the total DNA of the soil was extracted from 0.5 g fresh soil using the BIO101 Fast DNA Spin Kit for soil (MP-Biomedicals, France). The total DNA extracts were re-suspended in sterile deionised water in a final volume of 50 mL and were quantified using the Fluorescent DNA Quantification Kit (picoGreen, invitrogen) (Gangneux et al., 2011). The DNA extracts were stored at 20
C. The total bacterial communities and some bacterial taxa (Actinobacteria, Acidobacteria, Firmicutes, Alphaproteobacteria, Bacteroidetes, Gemmatimonadetes, Verrucomicrobiales,

Table 1 Physicochemical properties of soils. Site

Plot

Soil properties Clay (g kg

Yvetot Mean  (SD).

PG CC

1

163  (5.38) 133  (3.95)

)

Silt (g kg1)

Sand (g kg1)

Total C (g kg1)

Total N (g kg1)

pH water

CEC (cmol  kg1)

P2O5 (g kg1)

633  (4.64) 671  (3.92)

204  (4.99) 196  (6.45)

26  (0.96) 11  (0.34)

2.50  (0.07) 1.10  (0.04)

5.46  (0.05) 6.43  (0.04)

8.08  (0.26) 7.04  (0.30)

0.13  (0.13) 0.20  (0.02)

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Betaproteobacteria, Planctomycetes and Gammaproteobacteria) were quantified using the taxon-specific 16S rRNA qPCR assays developed by Philippot et al. (2011) using an Applied Bioystem 7300. The qPCR assay was carried out in a total volume of 15 mL with 1 SYBR green PCR Master Mix (Absolute QPCR SYBR Green Rox ABgene), 1 mM of each primer, 16.66 ng mL1 of T4 bacteriophage gene 32 (QBiogene, France), and 2 ng of total DNA. Standard curves were obtained using serial dilutions of linearized plasmids containing the cloned 16S rRNA genes from the different taxa. The qPCR was performed as described by Philippot et al. (2011) in an Applied Bioystem 7300. The qPCR efficiency for the assays ranged between 86% and 99%. The no template controls showed null or negligible values. The presence of PCR inhibitors in the DNA extracted from the soil was estimated by (1) diluting the soil DNA extract and (2) mixing a known amount of standard DNA with the soil DNA extract prior to the qPCR analysis. No inhibition was detected in either case. Two independent qPCR assays were performed for each taxon. The results are expressed as the 16S rRNA gene copy number per gram of soil dry weight (g1 DW). The results of the bacterial taxa quantification are expressed as a percentage of the changes from control in the relative abundance of bacterial groups, and the soils incubated at 17
C were used as a reference point in the PG and CC soils. The changes in the relative abundance of the bacterial taxa were calculated for each bacterial group and for each incubation time. A negative percentage indicates a decrease in the relative abundance, and a positive percentage indicates an increase in the relative abundance of the bacterial taxa by heat treatments. For the total fungal communities, 18S rRNA qPCR amplifications were performed in a total volume of 25 mL with 1 qPCR Master Mix (SYBR Green I, Applied Bioystem), 0.5 mM of FU18S1 and Nu-SSU-1536 primers (Borneman and Hartin, 2000), 0.5 mg mL1 BSA (NEB) and 10 ng of total DNA. Standard curves were obtained using serial dilutions of linearized plasmids containing the cloned 18S rRNA gene of Fusarium graminearum. After an initial denaturation and enzyme activation step of 10 min at 95
C, 40 cycles of PCR were performed in an Applied Bioystem 7300 as follows: 20 s at 95
C, 30 s at 62
C and 30 s at 72
C. The qPCR efficiency ranged from 95% to 98%. The results are expressed as the 18S rRNA gene copy number per gram of soil DW. 2.4. Enzyme assays The activities of four enzymes were measured using the methods described in Table 2. Cellulase, N-acetyl-glucosaminidase, b-glucosidase, xylanase and dehydrogenase activities were examined in three replicates for each soil sample. A unit of enzyme activity was defined as the nanomoles of substrate hydrolyzed or oxidized per minute and per gram of soil DW. As in previous approaches (Sannino and Gianfreda, 2001; Floch et al., 2011), the enzyme activity values of the soils incubated at 17
C were used as a reference point, which allows for the conversion of the enzyme activity values of soils incubated at 50
C as a percentage of changes. The enzymatic activities were calculated

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for each enzyme and for each incubation time. A negative percentage indicates enzymatic activities inhibition, and a positive percentage indicates stimulation by heat treatments. 2.5. Statistical analyses Statistical tests were performed to highlight the effects of heat stress on the soil total microbial biomass, the total bacterial and fungal communities, the abundance of different bacterial taxa and enzymatic activities. The data measured for soils incubated at 50
C and the control soils incubated at 17
C at each incubation time were compared by Student’s t-test with significant differences at 5% (P < 0.05). For each soil land use type, two Principal Component Analysis (PCA) were performed (see Supplementary data). The first PCA was performed with twelve structural variables, and the second PCA was performed with five functional variables. The projection of all the sample coordinates according to the first and second axis of the factorial plan of each PCA allowed us to extract these coordinates and calculate the Euclidean distances. Nine combinations of Euclidean distances were obtained because three replicates were performed for each soil land use type at each temperature and at each incubation time (see Section 2.2). The nine obtained Euclidean distances were used to calculate the means and standard deviations. The values of Euclidean distances provide information on the degree of the difference between the soil land use types for structural and functional profiles. For example, in the CC soil, the large distance measured at 5 days obtained with the structural variables indicates that heat stress had a large impact on the structure of the microbial communities in this soil. For each land use type, the correlation between the amplitude changes in structure and function (Euclidean distances) under heat stress was tested by Spearman’s correlations. All the statistical tests were computed with the R freeware (R Development Core Team, 2009). 3. Results 3.1. Abundance of microbial communities in PG and CC soil The total microbial biomass was measured by the total DNA quantification expressed as micrograms per gram of soil DW (Table 3). The heat stress significantly reduced the total microbial biomass (from 26.7  1.05 to 9.78  1.50 in PG and from 16.5  7.60 to 6.33  3.87 in CC). The microbial community was dominated by bacteria in both soil types. Fungi (expressed as the 18S rRNA gene copy number per gram of soil DW) were less abundant than bacteria in these soil types and were more sensitive to warming. Indeed, in PG soil the changes ranged from 2.28  108  0.74  108 to 0.28  108  0.15  108 and from 6.75  107  3.0  107 to 1.72  107  2.67  107 in CC soil. Bacteria (expressed as the 16S rRNA gene copy number per gram of soil DW) were not significantly affected by the heat stress. The bacterial biomass ranged (from 6.05  108  2  108 to 3.56  108  3.15  108 in PG and from 0.86  107  0.28  107 to 0.70  107  0.47  107 in CC soil).

Table 2 Protocols used for enzyme assays. Enzymes

E.C. number

Substrates (pH buffer and concentration)

References

Dehydrogenase Cellulase N-Acetyl-glucosaminidase b-Glucosidase Xylanase

1.1.1.1 3.2.1.4 3.2.1.30 3.2.1.21 3.2.1.8

2-(p-Iodophenyl)-3-(p-nitrophenyl)-5-phenyltetrazolium chloride p-NP-b-D-Cellobioside (pH 6; 10 mM) p-NP-N-Acetyl-glucosaminide (pH 6; 10 mM) p-NP-b-D-Glucopyranoside (pH 6; 10 mM) Xylan (pH 5.5, 12 g L1)

Schaefer, 1963 Trap et al., 2012 Trap et al., 2012 Eivazi and Tabatabai, 1988 Schinner and Von Mersi, 1990

NP: nitrophynyl.

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Table 3 The total nucleic acid and quantification of the ribosomal gene copies by qPCR from PG and CC soils at different incubation times. Land use

Days Treatment Total nucleic acid, mg g soil DW1

PG

5 14 21 28

– CC

5 14 21 28

Bacterial 16S rRNA genes copy number, g soil DW1 Fungal 18S rRNA genes copy number, g soil DW1 (1010) (108)

Con Warm Con Warm Con Warm Con Warm

25.6  (2.4)a 7.70  (2.6)b 28.0  (1.1)a 9.80  (1.5)b 27.1  (0.4)a 11.2  (5.6)b 26.2  (0.9)a 10.4  (4.2)b

4.30  (3.90) 0.82  (0.31) 6.20  (0.62) 4.50  (3.10) 8.80  (3.60)a 1.30  (0.40)b 4.90  (2.70) 7.60  (5.70)

1.20  (0.14)a 0.43  (0.03)b 2.80  (0.11)a 0.38  (0.18)b 2.70  (0.29)a 0.20  (0.09)b 2.40  (0.18)a 0.11  (0.07)b

Con Warm Con Warm Con Warm Con Warm

5.20  (2.2)a 12.1  (3.1)b 19.7  (1.8)a 4.00  (0.7)b 21.5  (0.9)a 4.90  (3.3)b 19.7  (1.3)a 4.30  (0.9)b

0.62  (0.15) 1.30  (0.13) 0.62  (0.20) 0.44  (0.39) 1.10  (0.21)a 0.23  (0.12)b 1.10  (1.00) 0.81  (0.51)

2.40  (2.00)a 5.70  (0.23)b 8.10  (5.20)a 0.76  (0.70)b 7.30  (6.30)a 0.17  (0.10)b 9.20  (6.40)a 0.24  (0.22)b

Mean  (SD). Letters (a and b) indicate significant difference between control (Con) and heated soil (Warm) in PG and CC soil at each incubation time (Student’s test P < 0.0 5, n = 3).

3.2. Changes in bacterial population under heat stress To estimate the relative abundance of the different studied taxa within the total bacterial community in PG and CC soils, we calculated the ratio of the taxon-specific 16S rRNA gene copies per gram of soil DW to the total bacterial 16S rRNA gene copies per gram of soil DW. In the control PG and CC soils incubated at 17
C, the sum of the relative abundance of all the studied taxa reached a maximum of 68% in the PG soil and 92% in the CC soil. In both soils types, the dominant taxa were Actinobacteria (13–40%) and Bacteroidetes (14–32%). The Planctomycetes and Gammaproteobacteria taxa had lower abundances (0–3%). The abundance of other groups (Acidobacteria, Firmicutes, Alphaproteobacteria, Gemmatimonadetes, Verrucomicrobiales, and Betaproteobacteria) ranged between the abundance of dominant and less represented taxa. The results of the changes from the control in the relative abundance of bacterial taxa in the PG and CC soil types are presented in Fig. 1. In the PG soil, a significant decrease was observed from the start of the incubation for the Acidobacteria, Planctomycetes, Alphaproteobacteria, and Gammaproteobacteria taxa, this decrease on average reached 80% in comparison to PG control soils. For the Actinobacteria, Bacteroidetes and Gemmatinomonadetes taxa, the decrease appeared only at the end of the incubation. The Firmicutes, Verrucomicrobiales and Betaproteobacteria taxa seemed to be insensitive to warming even if transient effects were observed. In the CC soil, among the 10 analyzed bacterial taxa, Gammaproteobacteria, and Acidobacteria decreased about 95% and Actinobacteria and Planctomycetes decreased about 20%, respectively, under heat stress in comparison to CC control soils. While Bacteroidetes, Gemmatinomonadetes, Verrucomicrobiales, Betaproteobacteria, and Alphaproteobacteria were less sensitive, as shown by the transitory increased or decreased effects. The Firmicutes taxa increased at the end of incubation period. 3.3. Heat stress effects on PG and CC soil enzymatic activities Heat stress treatment did not have the same impact on all the enzymes tested in the PG and CC soils (Fig. 2). In most cases, heat stress inhibited the enzymatic activities for the two soils over time. The cellulase, b-glucosidase dehydrogenase and N-acetyl-

glucosaminidase activities decreased significantly following heat treatment during the incubation time (5–28 days). For the N-acetyl-glucosaminidase activity at 5 days of incubation, a transient stimulating effect was observed (153%) in the CC soil. A different effect of heat stress was observed for xylanase activity, depending on the soils. In the PG soil, this activity increased significantly at 21 days of incubation (104%). In the CC soil, after an initial decrease, this activity tended to return to its initial soil background level at 14 days of incubation. 3.4. Relationships between structure and functions of soil microbial communities To explore the relationships between the soil microbial community structure and functions, Euclidean distances were calculated, and the correlations were used as described in Section 2. The first step consisted of a comparison of PG and CC reference soils based on the calculated distances to evaluate the impact of the two land use types on microbial community structure and functions. The results highlighted a constant difference between the two soils over time (average distances of 2). This finding confirmed the impact of management on the soil microbial community structural and functional states (Fig. 3). The second step relied on a comparison of the microbial community structure and functions in each soil type (PG and CC) under heat stress based on the calculated distances. In the PG and CC soils, heat stress had an effect on the microbial community structure and functions (see PCAs in Supplementary data). Significant changes from the control in structure and functions were observed (Fig. 4). According to our measurement changes approach, the structure and function of microbial communities of the CC soil were more severely impacted by heat stress compared with PG soil. The CC soil showed impacts that were 3 times and 1.5 times greater on the structure and functions, respectively (Fig. 4). In the PG soil, there was a significant change from the control in the microbial community structure at the beginning of incubation (distance = 1.32). This progressive change achieved its maximum at 21 days of incubation (distance = 6.32). Changes in the functions occurred at the beginning of the incubation and then remained constant. Spearman’s correlation showed a relationship between the changes in microbial community structures and the changes in functions in the PG soil (r = 0.34, P < 0.05). In the CC soil, significant

W. Riah-Anglet et al. / Applied Soil Ecology 86 (2014) 121–130

Actinobacteria

40 0

PG -40

CC

*

-80

* ‡ 14

5

28

‡

100

PG CC

0

14

21

28

‡ Acidobacteria

PG

200

CC

0

* 5

* ‡ 14

* ‡ 21

% of changes from control

400

* ‡ 28 days

*

‡

800

PG 400

CC

* 0

* -40 0

5

14

21

28

0

CC

-10 0

*

100

PG

0

CC

* ‡

* ‡

* ‡

-20 0

% of changes from control

200

14

21

28

days

21

28

days

α-Proteobacteria

200 100

PG

0

CC

-10 0

*

*

* 5

14

21

* 28

days

β-Proteobacteria

200

*

‡

150 100

PG

50

CC

0

‡

14

21

28

days

γ-Proteobacteria

600 400 PG 200

CC

0

-20 0

5

*

‡

300

800

300

14

5

5

‡

*

‡

-50

days

Planctomycetes

400

days

PG

250

% of changes from control

1200

28

100

-20 0

Verrucomicrobiales

1600

‡

200

400

600

21

300

days

800

14

‡ Gemmatinomonadetes

-20 0

1000

% of changes from control

CC

0

400

200

5

% of changes from control

PG 100

5

-10 0

% of changes from control

200

days

% of changes from control

% of changes from control

21

Firmicutes

300

-10 0

300

-10 0

-12 0

-20 0

Bacteriodetes

400

% of changes from control

% of changes from control

80

125

* ‡

* 5

14

* ‡

* ‡

21

28

days

Fig. 1. The effect of heat stress on the relative abundance of bacterial phyla from the PG and CC soils at each incubation time. The error bars represent the standard deviation of the mean of three replicates (n = 3). The signs (*) and (z) above the histogram indicate significant differences between the control and the heated soil at 50
C in the PG and CC soil, respectively (Student’s test P < 0.05).

changes from the control in the microbial community structure and enzymatic activities were observed at the beginning of the incubation. Changes in the microbial community structure then remained constant. Changes in the enzymatic activities declined

significantly and progressively after 21 days of incubation (distance decreased from 4.28 to 2.33). The changes in the microbial community structure were negatively correlated with the changes in the functions in the CC soil (r = 0.69, P < 0.05).

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‡N-acetylglucosaminidase

Cellulase % of changes in enzyme activity

% of changes in enzyme activity

300

PG

200

CC

100 0 -100

*

*

‡

‡

*

‡

-200

5

7

14

28

0

-40 -60

-100 -120

PP GC

-20 -40

* -80

‡

*

7

‡

14

*‡

28 % of changes in enzyme activity

5

‡

*

GC

* ‡

* ‡

5

7

*

14

28

days

Dehydrogense % of change in enzyme activity

% of changes in enzyme activity

β-glucosidase

PP

* ‡

-80

days

0

-60

‡

-20

0

PP

-20

GC

-40 -60

-80

*

-100 -120

days

*

‡

5

*

‡ 7

*

‡

14

‡

28

days

Xylanase

150

PP

* 100

*

GC

28

days

50 0

*

-50 -100

5

‡

7

14

Fig. 2. The effect of heat stress on N-acetylglucosaminidase, b-glucosidase, cellulase, xylanase and dehydrogenase activities of the PG soil and CC soil at each incubation time. The error bars represent the standard deviation of the mean of three replicates (n = 3). The signs (*) and (z) above the histogram indicate significant differences between the control and the heated soil at 50
C in the PG and CC soil, respectively (Student’s test P < 0.05).

4. Discussion 4.1. Changes of microbial community structure under heat stress

Joergensen and Brookes (1991), for example, observed a decreased in the soil microbial carbon biomass of 1.72% per day at 35
C during a 50 day incubation experiment. In the majority of heating studies, heat stress causes no effect or a drastic reduction in the soil

In this study, total microbial biomass was assessed according to the DNA analysis, however, our findings agreed with reductions reported in microbial biomass C due to heat stress in other studies.

Euclidean distance (Changes between PG and CC)

4

Euclidean distance (changes from control)

function

CC function

CC structure

PG function

PG structure

10

structure

3

8 6 4 2 0

2

5

14

5con-5warm x 14con-14warm Structure (S) * Fun ction (F) ns ANOVA

1

PG

0

CC

5

14

21

28

Days

Fig. 3. Changes in the microbial community structure and enzymatic activity functions at each incubation time expressed by Euclidean distances between the PG and CC control soils. The error bars represent the standard deviation of the mean of nine replicates. No significant differences were observed between the changes in the structure and function at 5 days and 14 days or between 21 days and 28 days (Student’s test P < 0.05).

Structure (S) Fun ction (F)

ns *

21

28

5con-5warm x 5con-5warm x 21con-21warm 28con-28warm * * ns ns ns *

ns *

Days

Correlation between S and F r = 0,34 *

r = - 0,69

*

Fig. 4. The changes in the microbial community structure and enzymatic activity functions in the PG soil and CC soil at each incubation time expressed by Euclidean distances. The error bars represent the standard deviation of the mean of nine replicates. (*) indicates a significant difference between the changes at 5 days and 14 days and between 21 days and 28 days (Student’s test P < 0.05). ns: no significant differences. r: Spearman coefficient correlation between structure and function.

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microbial biomass (Arnold et al., 1998; Rinnan et al., 2007; Allison and Treseder, 2008; Chaer et al., 2009; Schindlbacher et al., 2011) regardless of the analytical methods used (Prieto-Fernandez et al., 2004; Frey et al., 2008). The effects of high temperatures on the total microbial biomass lead to cell degradation of cellular storage material in response to the elevated energy requirements and death (Joergensen and Brookes, 1991; Petersen and Klug, 1994). Our results showed that the soil microbial community was dominated by bacteria, and they were less sensitive to heat stress during the 28 days of incubation, especially in the cropped soil (CC). This could be explained by the fact that bacteria are often dominant in agricultural soils (Fierer et al., 2005; Janssen, 2006) and are more tolerant to heat than fungi (Allison and Treseder, 2008; Frey et al., 2008; Rinnan et al., 2007; Keiblinger et al., 2012). In the present study, Actinobacteria and Bacteroidetes were the predominant bacterial phyla in the control and heated soils. These results are in agreement with other studies, such as the studies described by Acosta-Martinez et al. (2010),Youssef and Elshahed, (2009), Castro et al. (2010) which were based on massive sequencing. Actinobacteria are less sensitive to heat shock, which might be related to the spore-forming ability that provides an advantage over other phyla (Hayden et al., 2012). Due to their adaptation to heating, Actinobacteria are among the most important decomposers in soils (Kopecky et al., 2011) and are commonly found in extreme environments, such as dry volcanic soils or deserts (Connon et al., 2007; Costello et al., 2009). Bacteroidetes are poorly described in the literature compared with Actinobacteria. Bacteria belonging to the phylum Verrucomicrobia are nearly ubiquitous in soil (Zhang and Xu, 2008) but are usually considered to be one of the less frequent bacterial phyla (Wessén et al., 2010; Philippot et al., 2011), as observed in our study. The abundance of this bacterial group increased during heat stress, especially in the CC soil. This behavior could be attributed to the fact that this mesophilic phylum is more resistant to heat stress than other taxa (Sangwan et al., 2004). Regarding the Proteobacteria, Acidobacteria, and Planctomycetes, our results were inconsistent with previous findings because Alphaand Gammaproteobacteria, Acidobacteria and Planctomycetes were found to be severely decreased by heat stress, especially in PG soil, but these phyla are commonly described as able to survive in hot environments (Zhang et al., 2013; Fierer et al., 2007; Yergeau et al., 2012). In these three phyla, species that are important for biogeochemical cycling are heat-resistant and often produce thermo-stable enzymes (Torsvik et al., 2002). Our results confirm that heat stress induces the death of sensitive species (Bécaert et al., 2006), which in turn could promote the proliferation of surviving species, reduce competition and facilitate their access to resources. Heat stress might lead to changes in microbial community composition and diversity, creating a new microbial community structure that is particularly well adapted to stress and has significant functional stability (Girvan et al., 2005; Schimel et al., 2007). 4.2. Changes of soil enzymatic activities under heat stress The b-glucosidase, cellulase, N-acetyl-glucosaminidase, and dehydrogenase activities obtained in the PG and CC soils were inhibited by heat stress and did not recover during the incubation period. Similar results showed reductions in b-glucosidase, a-1, 4-glucosidase, cellobiohydrolase, N-acetyl-glucosaminidase, phosphatase, and sulfatase enzymatic activities for several weeks or months in soil temperature incubation experiments (Staddon et al., 1998; Waldrop and Firestone, 2004). The enzymatic activities measured in this study were extra-cellular with the exception of the dehydrogenase activity, which is intracellular (Burns et al., 2013). These enzyme activities are known to react quickly to severe

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stress (Caldwell, 2005; Bécaert et al., 2006; Chaer et al., 2009; Floch et al., 2011). The reduction of the soil enzymatic activities might be due to: (i) the decrease of enzyme synthesis and secretion in response to warming (Allison, 2005; Wallenstein et al., 2012); and (ii) the denaturation and the degradative activities of extracellular proteases (Wallenstein et al., 2011), though the literature also reported that soil enzymes are more tolerant to high temperatures (Nannipieri et al., 2003; Trasar-Cepeda et al., 2007); and (iii) the evolution of microbial metabolism due to changes in soil organic matter composition according to heat stress. Dehydrogenase that is involved in the oxidoreductase mechanism and gives information about active microbial biomass, was more severely affected than the other enzyme activities (inhibited on average about 87% in both soils), by heat stress in our study, as found by DNA analysis. These results are consistent with those of Arnold et al. (1998) showing an increase in the temperature induced a significant decrease in dehydrogenase activity linked to a microbial biomass reduction. However, soil samples incubated at temperatures ranging from 5 to 40
C showed a high positive correlation between the dehydrogenase activity and the soil temperature, and the activity increased in a linear manner over the entire temperature range (Trevor, 1984; Subhani et al., 2001). In our study, xylanase activity only was not affected by heat stress in the CC soil and was significantly stimulated in the PG soil. Stimulations of this activity were observed in several studies with an optimal xylanase activity of approximately 40–60
C (Collins et al., 2005; Isil and Nilufer, 2005). Other studies reported that the xylanase enzymes isolated from bacteria and fungi are extremely thermostable (Simpson et al., 1991; Gomes et al., 1993; Hayashi et al., 1999; Li et al., 2009). This property might explain the resistance of the xylanase enzyme to heat stress (Burns et al., 2013). The increases in xylanase activity would depend on the survival or proliferation of microbial populations able to produce this enzyme (Chaer et al., 2009). Furthermore, fungi are considered better producers of xylanase than bacteria (Collins et al., 2005). Thus, the higher stimulation of xylanase activity in the PG soil could be attributed to the difference in fungal community diversity between the CC and PG soils. The different behavior between the studied enzymes after heat stress could be explained by selective modification of microbial community functions (Dell et al., 2012). 4.3. Relationships between structure and functions of soil microbial communities We opted to use heat stress to induce changes in the microbial communities and soil enzymatic activities in order to examine the relationship between the microbial community structure and soil enzymatic activities. The relationship between the microbial community structure and soil function is not easy to predict and is poorly documented in experimental conditions similar as our (Jacobsen and Hjelmsø, 2014). This study highlighted a difference between the PG and CC control soils incubated at 17
C based on the calculated distances (Fig. 3) during 28 days of incubation. Changes in the microbial structure and functions are commonly observed in situ under different disturbances, such as land use (Six et al., 2004; Kaye et al., 2005; Brown et al., 2007; Roesch et al., 2007). The comparison of the impact of heat stress on microbial community structures and functions in the PG and CC soil showed that: (i) in PG, progressive changes were observed in the microbial community structure, and the changes in the functions occurred at the beginning remain constant during the incubation period (r = 0.34); (ii) in CC soil, changes in the microbial community structures were not significant, but changes in the functions were observed during the incubation period (r = 0.69). This finding suggested that the microbial community structure-function relationship was different in the PG and CC soils (van Eekeren

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et al., 2008). The soil microbial community structure and functions were positively correlated in the PG soil and negatively correlated in the CC soil. In our experiment, the PG soil was more resistant to drastic stress. This result confirmed our hypothesis, which stated that the very abundant and highly diversified microbial community in PG soil makes it more resistant to drastic stress as shown by DuPont et al. (2010). Based on our approach using the Euclidean distance calculation, our results showed that the changes in the microbial community structure and function induced by heat stress in PG soil is nearly the same as the differences existing between native CC and PG soils. We observed that the decreases in these enzyme activities were particularly associated with the decreases in the abundance of Actinobacteria, Acidobacteria and Planctomycetes known to have an extraordinary range of enzyme activities that include beneficial effects for plant growth (Gardner et al., 2011; Nunvar et al., 2014). The intrinsic metabolic capacity of species within these groups can explain their ability to colonize specific niches and to grow on pasture or cropped soil. Actinobacteria and Acidobacteria, have various correlations with several soil hydrolytic enzyme activities in agricultural soils (Zhang et al., 2013; Stone et al., 2014). The increase of Verrucomicrobiales and Firmicutes during heat stress in the cropped soil may indicate the role of these groups on the levels of the enzyme activities sustained during heat stress. Concerning fungal community, more research is needed to determine in more detail the fungal populations. Although this study did not evaluate the distribution of fungal members in detail, the decreases in fungal diversity were in agreement with the decrease in the activity of cellulase and N-acetyl-glucosaminidase, enzymes that have been mainly attributed to be produced by fungal populations (Alfonso et al., 1992; Valášková et al., 2007). 5. Conclusion We demonstrated that changes in the microbial community structure, led to losses of soil functions related to biogeochemical cycling via enzymatic activities. This study demonstrated changes in the soil fungal and bacterial diversity and within bacterial phyla distribution, due to laboratory induced heat stress, that could represent important modifications to soil function. Fungal diversity was more sensitive to the heat stress than bacterial diversity. Since heat stress is far from natural conditions, observed changes in microbial communities and functions can't be interpreted as potentially realistic changes. Our aim was only to induce a drastic change in soil microbial communities in order to increase our knowledge on structure and functions relationship. The soil management history played a key role in the resilience of these microbial groups to heat stress as the changes in the microbial community and functions were more pronounced in the conventional cropped soil. This supported our hypothesis that a more diverse microbial community is more resistant to drastic stress. Heat stress reduced the activities of several enzymes involved in biogeochemical cycling, except for xylanase. Changes in the microbial community structure and functions were positively correlated in the permanent grassland soil and negatively correlated in the conventional crop soil. These results represent a contribution to the understanding of soil microbial community structure and functions relationships. Advances in RNA extraction from soil might permit to determine active species in soil (Griffiths et al., 2000; Hurt et al., 2001) and contribute to a better understanding of these relationships. Acknowledgments These works were supported by the Research Networks VASI from the region Haute-Normandie, the French Ministry of Research

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