Substrate limitation to soil microbial communities in a subalpine volcanic desert on Mount Fuji, Japan

European Journal of Soil Biology 73 (2016) 34e45

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European Journal of Soil Biology journal homepage: http://www.elsevier.com/locate/ejsobi

Original article

Substrate limitation to soil microbial communities in a subalpine volcanic desert on Mount Fuji, Japan S. Yoshitake a, *, M. Fujiyoshi b, T. Masuzawa c, H. Koizumi d a

Takayama Field Station, River Basin Research Center, Gifu University, 919-47 Iwaimachi, Takayama, Gifu 506-0815, Japan School of Humanities and Culture, Tokai University, 4-1-1 Kitakaname, Hiratsuka, Kanagawa 259-1292, Japan c Graduate School of Science and Technology, Shizuoka University, 836 Ohya, Suruga, Shizuoka, Shizuoka 422-8017, Japan d Faculty of Education and Integrated Arts and Sciences, Waseda University, 2-2 Wakamatsucho, Shinjuku-ku, Tokyo 162-8480, Japan b

a r t i c l e i n f o

a b s t r a c t

Article history: Received 27 September 2015 Received in revised form 11 December 2015 Accepted 6 January 2016 Available online xxx

We examined two hypotheses based on laboratory amendment experiments: (1) that development of the soil microbial community on volcanic deserts was regulated by substrate limitation; and (2) that the type and the extent of substrate limitation would change along the succession gradient. Soils were collected from the early (Stage B) and the late (Stage F) stages of primary succession of a subalpine volcanic desert on Mt. Fuji and they were amended with three carbon (C) sources (glucose, cellulose, or lignin), inorganic nitrogen (N), or phosphorus (P) sources alone or in a mixture. Respiration rates were monitored for 25 days and changes in microbial biomass and community structure were examined using the content and composition of phospholipid fatty acids. For both soils, the magnitude of the microbial response differed depending on the type of C source and it decreased in the following order reflecting the availability to microorganisms: glucose > cellulose > lignin. For Stage B soil, although any single amendment did not affect the microbial properties, combined amendment of C (glucose) and N increased microbial respiration and biomass and shifted the microbial community structure. In contrast, microbial properties in Stage F soils responded positively to single amendments of C source. Our results suggest that the microbial community in the early stage of succession is primarily limited by simultaneous shortage of C and N sources but the quality of the C source becomes more important in the late successional stages, which have large, but recalcitrant, organic matter pools in the soil. © 2016 Elsevier Masson SAS. All rights reserved.

Keywords: Microbial biomass Microbial community structure Microbial respiration Primary succession Subalpine volcanic deserts Substrate limitation

1. Introduction In volcanic deserts, we can observe the development of ecosystems on virgin land surfaces covered by lava flows and ash deposits from eruptions. Volcanic deserts are thus a good model for primary succession and many previous studies have investigated the various ecological processes involved in primary succession, including changes in the microbial community [1e4]. In particular, alpine and subalpine volcanic deserts are thought to provide a distinct advantage for studying primary succession because of the presence of isolated “island-like vegetation communities” within small areas [5,6]. The structure of these communities reflects the time since the first colonization of pioneer plants, and important

* Corresponding author. E-mail addresses: [email protected] (S. Yoshitake), [email protected]. ac.jp (M. Fujiyoshi), [email protected] (T. Masuzawa), hkoizumi@waseda. jp (H. Koizumi). http://dx.doi.org/10.1016/j.ejsobi.2016.01.002 1164-5563/© 2016 Elsevier Masson SAS. All rights reserved.

factors such as climatic conditions and substrate type are the same, which enables us to determine actual successional series within a small area [6]. Volcanic deserts are extremely nutrient-poor ecosystems, and therefore the various soil microbial functions such as organic matter decomposition and accompanying nutrient release, N2 fixation, and mycorrhizal symbiosis with plant roots are very important for the maintenance and development of whole ecosystem functions. It is well known that some important soil conditions affecting soil microorganisms such as pH, soil organic matter (SOM) contents and inorganic nutrient concentrations can change drastically within a small area of volcanic desert along a primary succession [5,3,6]. These facts suggest that clarifying the changes in soil microbial characteristics during succession and its controlling factors would be important for understanding the shifts in ecosystem function and the rate and direction of ecosystem development. In general, biomass and diversity of the soil microbial

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community are very low during the initial phase of primary succession [7,8]. Similar trends have been reported in some volcanic deserts; microbial biomass and some indices for microbial activity were very low in bare ground in the earlier stages of succession [1,2]. We examined the successional changes in soil microbial biomass and community structure along the development of island communities in a subalpine volcanic desert of Mount Fuji, Japan, and found that biomass and richness of the soil microbial community was extremely low in the bare ground between island communities [6]. However, microbial biomass increased with primary succession (development of island communities) and was positively correlated with the increase in SOM and soil total carbon (C) and nitrogen (N) contents. Halvorson & Smith [3] examined soil and microbial properties in the volcanic desert of Mount St. Helens and reported clear correlations between soil C and N contents and microbial biomass. In addition to the biomass, microbial community structure shifted significantly along the succession on Mount Fuji and it was thought to be related to changes in the quality of SOM along the plant succession [6]. These results suggest that quantity and/or quality of SOM are important factors regulating the development of the soil microbial community. Heterotrophic microorganisms, important drivers of organic matter decomposition and mineralization in soil, need not only C as their energy source but also inorganic nutrients such as N and phosphorus (P) for growth [9]. Cleveland and Liptzin [10] reported a relatively consistent microbial biomass element ratio (C:N:P ratio) at the global scale and suggested that the soil microbial community was homeostatic based on the concept of ecological stoichiometry. Griffiths et al. [9] also confirmed that the C:N:P ratios of microbial biomass were homeostatic under near optimum soil conditions. This fact indicates that disparities between soil element ratios (substrate element ratios) and the soil microbial biomass ratio would suppress the activity of the soil microbial community. In fact, many examples demonstrated that substrate limitation to the microbial community, either a shortage of C source and/or inorganic nutrients, limited microbial activity in various ecosystems [11,12]. In particular, activity and growth of the soil microbial community was regulated by substrate limitation in ecosystems characterized by very low SOM contents such as an Antarctic dry valley [13], high arctic tundra [14], highly acidic solfatara [15] and dry desert [16]. Volcanic deserts are also nutrient-poor ecosystems, and it is possible that shortage of C and/or inorganic nutrients such as N and P could be one of the major factors regulating the soil microbial community. Availability of substrate in the soil (labile or recalcitrant) as well as the absolute amount should be another important aspect explaining substrate limitation to the soil microbial community [11]. However, information about substrate limitation to the soil microbial community in subalpine volcanic deserts has not yet been obtained and therefore the mechanisms of microbial succession, especially in the early stages of primary succession of volcanic deserts, are not well understood. Here, we hypothesized that development of the soil microbial community on volcanic deserts was regulated by substrate limitation, and the type and the extent of substrate limitation would differ between the early and late stages of succession. Our objective was to examine these hypotheses by conducting laboratory amendment experiments using soils collected from different successional stages. In this study, three types of organic C sources with different availability for soil microorganisms and inorganic N and P sources were added to the soils alone or in combination; the changes in quantity (biomass) and quality (respiration and community structure) of the soil microbial communities were examined.

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2. Materials and methods 2.1. Study site This study was conducted at the same site as Yoshitake et al. [6] and was on the southeastern slope of Mount Fuji, between 1500 and 1550 m above sea level (35
200 N, 138
470 E). The annual mean air temperature in 2009 was 7.8
C, and the monthly mean air temperature ranged from 2.8
C in January to 17.9
C in August. Mean annual precipitation at Gotenba, about 15 km southeast of the study site, averaged 2801 mm between 2000 and 2014. An accumulation of basaltic ejecta composed mainly of scoria (typically 2e30 mm wide) from Mount Hoei (2702 m), which is a parasitic volcano that formed on the mid-slope of Mount Fuji during the 1707 eruption, covers much of the southeastern slope [5]. In this area, previous studies have investigated the pattern and mechanisms of plant succession, which begins with isolated islandlike communities of the perennial herb Polygonum cuspidatum Sieb. et Zucc. (¼ Reynoutria japonica Houttuyn) [17,18]. As P. cuspidatum spreads outward, its shoot density decreases in the center, thus providing a colonization site for later successional species (the “central dieback” phenomenon [17]). As a result of differences in the timing of the initial colonization by the pioneer P. cuspidatum and its slow growth rate, a number of island-like communities with different successional stages are scattered within the study site. Detailed descriptions of the successional pattern of these islandlike communities are provided in Fujiyoshi et al. [5] and Yoshitake et al. [6]. In this study, the bare ground between the islands of vegetation dominated by P. cuspidatum (cf. stage I [6]) was chosen as the first pioneer stage (Stage B), whereas the Larix kaempferi forest adjacent to areas that contained the island-like communities was chosen as a late successional stage (Stage F). The late stage is thought to be formed by integrating the island communities containing the L. kaempferi in the central part. Stages B and F in this study were identical to those in a previous study [6], and some soil properties in Stage B and F soils are summarized in Table 1. 2.2. Soil sampling Three study plots (each 1 m  1 m) were selected for Stages B and F (i.e., n ¼ 3 plots for each successional stage). At these study plots, mineral soil samples (0e5 cm below the mineral soil surface) were collected in late October 2009 using a sterilized stainless-steel soil corer (diameter (Ø) ¼ 5 cm, height (H) ¼ 5 cm, volume ¼ 100 cm3). The thin organic soil layer (almost zero at Stage B and less than 3 cm at Stage F) was removed carefully before sampling. Three core samples per plot were collected and composited. All collected samples were brought back to the laboratory with a cooler and plant roots and gravel were removed using a 4-mm mesh sieve and tweezers. These soil samples were stored at 20
C (the temperature at which respiration measurements were conducted) with their original water contents for up to 5 days after sampling until the amendment experiments were carried out. 2.3. Changes in microbial respiration after substrate amendment The respiration rates after the addition of the C, N and P sources individually, or in combination, were measured for 25 days (respiration was measured on days 0, 1, 2, 5, 10, 15, 20 and 25) to determine whether the availability of these elements limited microbial respiration. We used three types of C source independently in the experiments: C source 1 (C1), glucose, representing a very labile monosaccharide; C2, cellulose, known as the main

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Table 1 Soil characteristics of the study sites (0e5 cm of mineral soil layer). Values are means (±SD) for three replicates (n ¼ 3). Significance was tested by T-test and expressed as*, P < 0.05; **, P < 0.01; ***, P < 0.001.

Soil structure (% of total weight) >4 mm 2e4 mm <2 mm pH (H2O)a Water content (g g1 dry soil)b Watar holding capacity (WHC) (g g1 dry soil)b,c Organic matter content (mg g1 dry soil)d Total carbon (C) (mg g1 dry soil)e Total nitrogen (N) (mg g1 dry soil)e C/N ratio Dominant vegetation a b c d e

Stage B

Stage F

P value

Significance

45.7 (5.3) 16.2 (2.4) 38.1 (7.4) 5.5 (0.02) 0.09 (0.001) 0.37 (0.012) 4.16 (1.6) 0.8 (0.05) 0.05 (0.006) 15.0 (2.0) None

39.8 (18.3) 13.6 (2.6) 46.7 (15.8) 5.8 (0.07) 0.29 (0.005) 0.65 (0.032) 68.7 (41.8) 34.3 (16.1) 1.57 (0.96) 23.0 (2.9) Larix kaempferi

0.62 0.27 0.44 <0.01 <0.001 <0.001 0.056 <0.05 0.052 <0.05

ns ns ns ** *** *** ns * ns *

Soil (air-dried): water ¼ 1: 5. Determined on 30 October 2009. Determined by the methods of Paul et al. [19]. Determined by loss in weight on ignition (550
C, 4 h). Measured with a CN analyzer (Sumigraph NC-22; Sumika Chemical Analysis Service, Tokyo, Japan).

component of plant cell walls and representing a relatively labile polysaccharide; and C3, lignin, a recalcitrant polymer with a polyaromatic nature. Ammonium nitrate (NH4NO3) and potassium dihydrogen phosphate (KH2PO4) were used as N and P sources, respectively. A portion of each replicate soil sample from each successional stage (stages B and F) was allocated to each of the following treatments: C1þ, C2þ, C3þ, Nþ, Pþ, C1Nþ, C2Nþ, C3Nþ, C1Pþ, C2Pþ, C3Pþ, NPþ, C1NPþ, C2NPþ, C3NPþ, and a control (Fig. 1). A soil sample equivalent to 30 or 10 g dry weight for Stage B and F soils, respectively, was placed in a cylindrical plastic case (Ø ¼ 9 cm, H ¼ 2.5 cm). C, N and P sources were added with water to achieve the desired water content (60% of water-holding capacity) at the rate equivalent to 20 mg C, 2 mg N and 0.4 mg P, respectively, for each 1 g of dry soil. Soils supplied with water alone served as controls. The amount of C source was considered in advance based on the basal respiration rate and the length of the incubation period (25 days); sufficient C source was added to monitor the changes in microbial respiration rate throughout the experimental period. The C:N:P ratio (50:5:1) was within the range of those reported in previous studies [20e22] and was close to the value for soil microbial biomass (60:7:1) in Cleveland & Liptzin [10]. Reagents except for cellulose and lignin were added in solution, while

Fig. 1. Concept of the carbon (C), nitrogen (N), and phosphorus (P) amendment experiment using three types of C source. Positions overlapping with two or three elements indicate combined amendment.

cellulose and lignin were added directly as powder together with water or N and P solution because solubility of these reagents was very low. After reagent addition, the soils were mixed adequately and immediately using a spatula. All soil samples were incubated in the dark at 20
C. Microbial respiration (as the CO2 emission rate) was measured at 20 ± 0.3
C in an open-flow system with an infrared gas analyzer (LI-6262; Li-Cor Inc., Lincoln, USA). The plastic case containing the sample was placed in a cylindrical chamber (Ø ¼ 9.2 cm, H ¼ 3 cm) connected to the system. Ambient air containing 400e410 ppm CO2 was pumped into the system at a suitable flow rate (100, 200, or 500 mL min1), depending on the amount of CO2 released from the sample. During measurements, the chamber was placed in a water bath to maintain a constant temperature of 20 ± 0.3
C. Respiration rate was calculated based on the differences in CO2 concentration between the chambers with and without sample, taking into account the flow rate and temperature. 2.4. Changes in microbial biomass and community structure after substrate amendment The phospholipid fatty acid (PLFA) contents in soil before and after substrate addition (C, N and/or P sources) were determined to elucidate the changes in microbial biomass and community structure. The experimental design was almost identical to that for the respiration measurement (described above) but samples for PLFA analyses were obtained on days 0, 10 and 15 after substrate addition for Stage B soil and days 0, 5 and 15 for Stage F soil, taking into consideration the timing of the major increase in microbial respiration rate in each stage soil. On sampling days, a portion of each soil sample was collected, freeze-dried, sieved (<2 mm) and stored at 80
C until analysis. Lipids were extracted using the extraction method of Bligh & Dyer [23], as modified by White et al. [24] and Frostegård et al. [25]. Briefly, 1.5 or 3.0 g (dry weight) of soil for Stage B and Stage F soil, respectively, were extracted in a chloroformemethanolecitrate buffer (0.15 M, pH 4.0) (1:2:0.8 v/v/v). The lipids were separated into neutral lipids, glycolipids, and phospholipids on a silicic acid column (Sep-Pak Plus Silica; Waters Corp., Milford, USA). The phospholipids were esterified with 5% HClemethanol (Tokyo Kasei Kogyo Co., Ltd., Tokyo, Japan [26]). The resultant fatty acid methyl esters were separated using a gas chromatograph (GC-2014; Shimadzu Corp., Kyoto, Japan) equipped with a capillary column (PhenyleMethyl/Silicone (30 m DB-5); J&W Scientific Inc., Folsom, CA, USA). Helium was used as the

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carrier gas. Column oven temperature was ramped from 80
C to 200
C at 15
C min1 and then to 240
C at 2
C min1 and held for 3.5 min. Peak areas were quantified against methyl nonadecanoate fatty acid (19:0) as an internal standard. We used the fatty acid nomenclature of Frostegård et al. [27] and the total PLFA content (TotPLFA; nmol g1) was used to indicate the total microbial biomass [27]. We used the PLFA composition as an index of microbial community structure because phospholipids are good indicators of the composition of living soil microbes [24].

normal and a dissimilarity matrix of PLFA composition using the BrayeCurtis dissimilarity coefficients was calculated. Generated dissimilarity matrices were subjected to NMDS using R software (version 3.0.2 [30]) and we obtained two-dimensional ordination graphs.

2.5. Statistics

For both successional stages, microbial respiration rates of control soils did not change significantly throughout the experimental period (one-way ANOVA, P > 0.05) and the rates in Stage F (1.6e3.3 mg CO2eC g1 h1) were almost tenfold those in Stage B (0.1e0.3 mg CO2eC g1 h1). In addition, microbial respiration rates of soils with no added C source (Nþ, Pþ, NPþ) were similar to those in control soils throughout the experimental period (Figs. 2a and 3a).

We conducted a three-way analysis of variance (ANOVA) for each day with C, N and P source as factors to test the effects of the addition of C, N and/or P on microbial respiration rates and total PLFA contents. We used Dunnett's test for multiple comparisons for each day. Significance was inferred in cases where P < 0.05. Changes in microbial community structure among the amendment treatments were examined based on PLFA composition with the nonmetric multidimensional scaling (NMDS) ordination technique [28]. NMDS is an ordination method that iteratively searches for the best way to represent the data in a reduced number of dimensions so that the distances in the ordination diagram reflect the dissimilarities in community structure of the original samples. This method is considered better than many other ordination techniques because it does not assume a linear relationship among variables [28,29]. For the PLFA composition data (averaged mol% data matrix), the arcsine square-root transformation was used according to Hannam et al. [28] because the mol% data were non-

3. Results 3.1. Microbial respiration

3.1.1. Changes in microbial respiration rate in Stage B soils Respiration rates in soils with C source added changed remarkably but the pattern differed depending on the type of carbon source used (Fig. 2bed). When glucose was used as a C source (Fig. 2b), significant interaction among C, N, and P addition (C  N  P) was detected from day 2 onwards, except for day 15 (three-way ANOVA, P < 0.01; Table A.1). The microbial respiration rates in soils with C source only (C1þ) and soils with both C and P sources (C1Pþ) were slightly higher than those in control soils on days 2 and 5 only (Dunnett's test, P < 0.01; Fig. 2b). In contrast, the

Fig. 2. Responses of soil microbial respiration rate to substrate amendments for Stage B soil. a) control and N and/or P addition without any type of C source (Control, Nþ, Pþ and NP þ treatments); b) glucose addition with and without N and/or P (Cþ, CNþ, CPþ and CNP þ treatments); c) cellulose addition with and without N and/or P; d) lignin addition with and without N and/or P. Values are means; vertical bars represent ± SD (n ¼ 3). Bars marked with an asterisk are significantly different from the control (Dunnett's test, P < 0.05). Note that the y-axis scale in panel a) differs from the other panels.

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Fig. 3. Responses of soil microbial respiration rate to substrate amendments for Stage F soil. a) control and N and/or P addition without any type of C source; b) glucose addition with and without N and/or P; c) cellulose addition with and without N and/or P; d) lignin addition with and without N and/or P. Values are means; vertical bars represent ± SD (n ¼ 3). Bars marked with an asterisk are significantly different from the control (Dunnett's test, P < 0.05). Note that the y-axis scale in panel a) differs from the other panels.

respiration rates of soils with C and N source added (C1Nþ) increased greatly after day 2 and the highest respiration rate in this treatment was 28.0 mg CO2eC g1 h1 on day 20. The respiration rates in soils with C, N and P sources added (C1NPþ) increased more rapidly and to a greater extent than the C1N þ treatment and reached a maximum value (39.3 mg CO2eC g1 h1) on day 10. When cellulose was used as a C source (Fig. 2c), a clear and significant interaction among C, N and P addition (C  N  P) was detected after day 2 (three-way ANOVA, P < 0.001; Table A.1). The respiration rates in C2þ, C2Nþ and C2P þ treatments were generally the same or slightly higher than those in control soils. In contrast, the respiration rate in the C2NP þ treatment was significantly higher than in the control after day 2 and the maximum respiration rate in this treatment was 27.0 mg CO2eC g1 h1 on day 15 (Dunnett's test, P < 0.001; Fig. 2c). When lignin was used as a C source (Fig. 2d), significant interaction among C, N and P addition (C  N  P) was remarkable on day 10 and interaction between C and N addition (C  N) was significant on day 15 (three-way ANOVA, P < 0.05 for both days; Table A.1). The respiration rates in C3þ and C3P þ treatments were similar to those in the control soil. The respiration rates in C3Nþ were significantly higher than the control on days 10 and 15 and C3NPþ was higher than the control on day 10 (Dunnett's test, P < 0.01; Fig. 2d).

3.1.2. Changes in microbial respiration rate in Stage F soils When glucose was used as a C source (Fig. 3b), significant C  N  P interaction was observed for days 1e10 and on day 25 (three-way ANOVA, P < 0.01; Table A.2). On days 15 and 20, significant C  N and C  P interactions were observed (three-way ANOVA, P < 0.001; Table A.2). The respiration rates of soils with

glucose addition (C1þ, C1Nþ, C1Pþ, C1NPþ) were all significantly higher than those in control soils (Dunnett's test, P < 0.001; Fig. 3b). The maximum respiration rates (in mg CO2eC g1 h1) for each treatment were in the following order: C1NPþ (49.3) > C1Nþ (34.8) > C1Pþ (21.9) > C1þ (9.3). When cellulose was used as a C source (Fig. 3c), significant C  N  P interaction was observed for days 2e15 (three-way ANOVA, P < 0.05; Table A.2). The respiration rates in C2þ, C2Nþ and C2P þ treatments were generally similar. In contrast, the respiration rates in C2NPþ were significantly higher than those in the control throughout the experimental period (Dunnett's test, P < 0.01; Fig. 3c) and the maximum rate for this treatment was 17.6 mg CO2eC g1 h1 on day 5. When lignin was used as a C source (Fig. 3d), the largest changes in microbial respiration rate were observed between days 1 and 5. Significant C  N  P interaction (P < 0.001) and C  N interaction (P < 0.05) was detected on day 2 and days 1e5, respectively, and the respiration rates in C3þ, C3Nþ, C3Pþ and C3NP þ treatments were all significantly higher than that in the control on these days (Dunnett's test, P < 0.05; Fig. 3d).

3.2. Microbial biomass For both successional stages, total PLFA content (TotPLFA; as an index of microbial biomass) in control soils did not change significantly throughout the experimental period (one-way ANOVA; P > 0.05). TotPLFAs were 8.2e10.9 and 147e194 nmol g1 for Stages B and F soils, respectively. TotPLFA contents of soils without any added C sources (Nþ, Pþ, NPþ) were constant throughout the experimental period and were not different from those in control soils (Dunnett's test, P > 0.05; Figs. 4a and 5a). Similar to the

Fig. 4. Changes in total PLFA content after substrate amendment for Stage B soil. a) control and N and/or P addition without any type of C source; b) glucose addition with and without N and/or P; c) cellulose addition with and without N and/or P; d) lignin addition with and without N and/or P. Values are means; vertical bars represent ± SD (n ¼ 3). Bars marked with an asterisk are significantly different from the control (Dunnett's test, P < 0.05). Note that the y-axis scale in panel a) differs from the other panels.

Fig. 5. Changes in total PLFA content after substrate amendment for Stage F soil. a) control and N and/or P addition without any type of C source; b) glucose addition with and without N and/or P; c) cellulose addition with and without N and/or P; d) lignin addition with and without N and/or P. Values are means; vertical bars represent ± SD (n ¼ 3). Bars marked with an asterisk are significantly different from the control (Dunnett's test, P < 0.05).

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microbial respiration rate, the response pattern of TotPLFA content to C source with and without N and P sources differed largely depending on the type of carbon source used (Figs. 4 and 5). 3.2.1. Changes in total PLFA content in Stage B soils When glucose was used as a C source, a clear and significant C  N  P interaction was observed on days 10 and 15 (three-way ANOVA, P < 0.001 for both days; Table A.3) for TotPLFA content. TotPLFA contents in C1þ and C1P þ soils did not differ significantly from those in control soils (Dunnett's test, P > 0.05; Fig. 4b). TotPLFA content in C1N þ soils increased significantly compared with the control and reached 131 nmol g1 on day 15. TotPLFA content in C1NP þ soils increased to a maximum value of about 680e700 nmol g1. Significant C  N  P interaction on PLFA content was observed for cellulose addition (three-way ANOVA, P < 0.001; Table A.3). PLFA content was significantly higher than the control only in C2NP þ soils (Dunnett's test, P > 0.05; Fig. 4c) and the maximum value in this treatment was 269 nmol g1 on day 10. The response of TotPLFA content to lignin addition (Fig. 4d) was generally small. Although the TotPLFA contents tended to increase in C3Nþ and C3NP þ on day 15, these increases were not statistically significant (Dunnett's test, P > 0.05; Fig. 4b). 3.2.2. Changes in total PLFA content in Stage F soils Significant C  N  P interaction on TotPLFA content occurred when glucose was added (three-way ANOVA, P < 0.01; Table A.3). TotPLFA contents in treatments with glucose added were all higher than that in the control soil on day 5 (Dunnett's test, P < 0.05;

Fig. 5b). The maximum TotPLFA values in C1Nþ and C1NP þ soils were about 460 and 740 nmol g1 on day 15, respectively. When cellulose was added as a C source, significant C  N  P interaction was observed only on day 5 (three-way ANOVA, P < 0.001; Table A.3) and C2Nþ and C2NP þ soils showed significantly higher TotPLFA content when compared with the control soil (Dunnett's test, P < 0.05; Fig. 5c). When lignin was used as a C source, TotPLFA contents in C3þ and C3NPþ were significantly higher than in the control soil on day 15 (Dunnett's test, P < 0.05; Fig. 5d). 3.3. Microbial community structure NMDS two-dimensional ordination graphs were obtained based on the averaged PLFA composition data. The dimensions (axes) in these graphs have no special significance and can be rotated or mirrored without influencing the relative distances between the points. In addition, the distances between any two data plots reflect the dissimilarities in PLFA composition, namely community structure. The “stress values” of NMDS, a measure of the goodness-of-fit of the reproduced distances to the observed distances [31], in our analyses were 0.100 and 0.073 for Stage B and F, respectively and they indicated the “fair” fitness of the NMDS plots produced according to the definition of stress in Kruskal [31]. 3.3.1. Changes in PLFA composition in Stage B soils The NMDS ordination of soils without any C source addition (control, Nþ, Pþ and NPþ) were relatively stable and located in the left part of the diagram (Fig. 6a). The positions of C1Nþ and

Fig. 6. Changes in soil microbial community structure after substrate amendment for Stage B soil according to nonmetric multidimensional scaling (NMDS) based on PLFA composition. a) control and N and/or P addition without any type of C source; b) glucose addition with and without N and/or P; c) cellulose addition with and without N and/or P; d) lignin addition with and without N and/or P. Data are mean values (n ¼ 3). Open symbols and a line within each treatment indicate the data for soils just after substrate addition (day 0) and representing temporal change (days 10 and 15), respectively.

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C1NP þ soils notably shifted to the upper right during the experimental period (Fig. 6b) and similarly, the positions of C2NPþ, C3Nþ and C3NP þ soils shifted to the upper right of the diagram (Fig. 6c and d). In these soils, the proportion of PLFA 18:2u6 was increased from about 2 mol% (day 0) to >40 mol% (day 15). In addition, the proportion of PLFA 18:1u9 was increased from about 10 mol% (day 0) to >20 mol% (day 15). 3.3.2. Changes in PLFA composition in Stage F soils The NMDS ordination of soils without any C source addition (control, Nþ, Pþ and NPþ) were relatively stable during the experimental period (Fig. 7a). In contrast, the positions of C1Nþ and C1NP þ soils shifted to the right of the diagram (Fig. 7b). The position of C2NP þ also shifted to the right (Fig. 7c). Similar to the case for Stage B soils, an increase in the proportion of 18:2u6 (from about 6 mol% to >24 mol%) and 18:1u9 (from about 9 mol% to >15 mol%) was observed in these soils. When lignin was used as a C source, the changes in NMDS ordination were generally small (Fig. 7d). 4. Discussion 4.1. Amendment experiment Amendment experiments are one of the major approaches to examine the soil response to external inputs such as organic matter and nutrients [11e13,32]. However, care must be taken in interpreting the results from laboratory amendment experiments

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because the addition of relatively high levels of substrate might bias the microbial community structure if they are toxic to oligotrophic microbes and/or promote the growth of copiotrophs [33]. Nevertheless, this approach has the great advantage that various types of substrates can be applied and detailed responses of soil microorganisms to each substrate, either alone or in combination, can then be determined. Therefore, this approach is a direct and powerful tool for understanding the relationship between soil C and N dynamics and the soil microbial community [34]. Several previous and recent studies have used this approach to examine substrate limitation of soil microorganisms and have succeeded in clarifying the potential but detailed responses of soil microbial communities to various substrate amendments [12,32,35]. In this study, an increase in soil microbial respiration after amendment (Figs. 2 and 3) was generally accompanied by an increase in the biomass (Figs. 4 and 5) and a shift in the community structure (Figs. 6 and 7). In addition, shifts in microbial community structure were generally accompanied by large increases in the proportions of 18:2u6 and 18:1u9, which are indices for fungal biomass [27,36]. Therefore, amendment itself might have had selective effects on the soil microbial community and increased the proportions of copiotrophic fast-growing fungi. However, the response patterns of soil microbial properties to the amendments differed greatly between successional stages and among treatments, providing valuable information about changes in potential substrate limitation during soil development along the primary succession.

Fig. 7. Changes in soil microbial community structure after substrate amendment for Stage F soil according to nonmetric multidimensional scaling (NMDS) based on PLFA composition. a) control and N and/or P addition without any type of C source; b) glucose addition with and without N and/or P; c) cellulose addition with and without N and/or P; d) lignin addition with and without N and/or P. Data are mean values (n ¼ 3). Open symbols and a line within each treatment indicate the data for soils just after substrate addition (day 0) and representing temporal change (days 5 and 15), respectively.

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An increase in microbial respiration with an increase in biomass occurred for all treatments except in the case of lignin amendment to Stage F soils (Fig. 3d). This exception suggests that the response of microbial respiration to lignin amendment was transient and that microbial growth was not inhibited by substrate (lignin) limitation. An increase in microbial respiration without an increase in biomass was also observed by Schimel and Weintraub [37]. They explained this phenomenon with a theoretical model that showed an increase in “waste respiration” through the overflow metabolisms in N-limited soil. Therefore, monitoring of both microbial physiological activity (i.e., respiration rate) and actual growth (i.e., changes in biomass and community structure) would be desirable to elucidate a more complete understanding of substrate limitation in the amendment experiments. 4.2. Substrate limitation in the early stage of succession (Stage B) Any single addition of C, N or P source did not have remarkable effects on the soil microbial properties (respiration rate, biomass and community structure) in Stage B soil. However, combined addition of C and N (CNþ) triggered an increase in the microbial respiration rate and biomass and shifts in the microbial community structure. In addition, there was significant interaction between C and N additions on microbial respiration rate and biomass during the experimental period. These facts suggest that the microbial community of Stage B soils was primarily limited by simultaneous shortage of C and N. Similar CN limitation was also found in other soils without vegetation cover in some desert ecosystems such as hot dry desert [16], high arctic glacier foreland [14], and highly acidic solfatara [15]. Absolute amounts of C and N were extremely low especially in the early stages of succession in these desert ecosystems, which would result in the shortage of both of C and N (CN limitation). Absolute amounts of available P were also very low [5] but combined amendment of C and P (CPþ) did not affect the microbial community (Figs. 2a, 4a and 6a). This emphasizes that shortage of N source was an important limiting factor for soil microorganisms. The microbial responses to the CNP þ treatment were faster and greater than those in the CN þ treatment (Figs. 2b and 4b). These results suggest that P shortage could be a secondary or additional limiting factor of the soil microbial community in Stage B soils. Cellulose is the main component of plant cell walls, and it constitutes 20e30% of the plant litter mass [38]. Microbial cellulolytic capabilities are restricted to certain functional groups: various fungi and certain groups of bacteria [39]. The fact that a remarkable increase in microbial respiration in response to cellulose addition only occurred in the CNP þ treatment (Fig. 2b) and that it was accompanied by an increase in fungal PLFAs (18:2u6 and 18:1u9) suggested that cellulose-decomposing fungi were strongly suppressed by both N and P limitation. Güsewell and Gessner [40] reported that cellulose decomposition was N or P limited depending on the N:P ratio and fungi were particularly important when decomposition was limited by P shortage. These facts suggest that P supply is an important factor for cellulose-decomposing fungi and plant litter decomposition. The response pattern of microbial respiration in Stage B soil after lignin amendment (Fig. 3d) was similar to the case with glucose (Fig. 3b), that is, microbial responses were mainly observed in CNþ and CNP þ treatments, suggesting the simultaneous shortage of C and N. However, the maximum values of the respiration rate and biomass in the CNP þ treatment were much smaller than those in the cases of glucose or cellulose addition and the increase in the microbial respiration rate was transient (during days 5e15; Fig. 2d). These weak responses reflect the fact that lignin is a recalcitrant compound because of the complex chemical structure of the aromatic polymer and can only be used by limited groups of microorganisms such as

wood- and leaf-decaying basidiomycetes [41]. 4.3. Substrate limitation in the late stage of succession (Stage F) The patterns of microbial responses to substrate amendments for Stage F soils were generally similar to those of Stage B soils except as detailed below. The most important difference between these two stages was the microbial responses to the single addition of glucose (C1þ treatment); the microbial respiration rate in Stage F soils significantly increased and was maintained at a high level in the C1þ treatment (Fig. 3b) compared with the response in Stage B soil (Fig. 2b). In addition, the microbial responses to the amendments in Stage F soils were arranged in the following order: C1NPþ > C1Nþ > C1Pþ > C1þ > control. These results suggest that the soil microbial community in Stage F soils was primarily limited by the shortage of labile C source and the N and P were secondary and tertiary limiting factors, respectively. Previous studies have also reported C limitation to the microbial community and that the respiration rate of C-amended soil with N was higher than those without N [12,14,16]. This is probably because significant amounts of SOM and, by extension, C, N, and P sources, would have already accumulated at this stage, but most of them would have been in recalcitrant form with relatively high C/N ratio [7]. As a result, the soil microbial community in the late stage of succession was primarily limited by a shortage of labile C source. Similar to the case for Stage B soils, a significant microbial response to cellulose amendment was only observed in the C2NP þ treatment in Stage F soil and a strong interactive effect among C, N and P amendment existed (Fig. 3c), suggesting a simultaneous shortage of these three elements for cellulose-decomposing microorganisms. As mentioned above, the response of microbial respiration to lignin amendment (Fig. 3d) did not coincide with those of microbial biomass (Fig. 5d) and community structure (Fig. 7d). Large differences in the microbial response pattern between glucose and lignin amendments in Stage F soils emphasized that the quality of the C source rather than the absolute amount would be a very important limiting factor in the late stages of succession. 4.4. Succession and substrate limitation Our results indicated that the type of substrate limitation differed among microbial functional groups. This means that substrate quality (availability of C source and C:N:P ratio) and its change could regulate microbial community structure through selective effects on microbial functional groups. Many previous studies reported that litter quality was an important predictor for litter decomposition rate [42,43] and that changes in litter chemical properties during decomposition were related to shifts in the microbial community [44]. The primary succession gradient, which is a relatively long term ecological process compared with the litter decomposition process, would also involve changes in substrate quality, that is, changes in SOM quality. In our study site, vegetation cover developed remarkably from almost bare ground (Stage B) to the forest dominated by L. kaempferi (Stage F) that produced high-lignin-content, recalcitrant leaf litter [45]. This suggests that plant litter quality differed greatly between the stages. In addition, significantly higher soil C/N ratio was observed in the Stage F soils (Table 1) than in the Stage B soils. These facts imply a decrease in the quality of SOM with the progress of the primary succession, which would have a considerable impact on the soil microbial community via the substrate limitation mechanism. In the early stage of succession, the microbial community relies on newly supplied organic matter as a source of C, N and P, because both the SOM content and the concentrations of inorganic nutrients are quite low at this stage (c.f. Table 1) and therefore the microbial community suffers from quantitative

S. Yoshitake et al. / European Journal of Soil Biology 73 (2016) 34e45

substrate limitation. In contrast, microbial communities in the late stage of succession suffer from accumulated recalcitrant SOM but they would be able to use the relatively large soil N and P pools. As a result, the microbial community was regulated by the availability of labile C in this stage. Although vegetation development in the late stage of succession might cause competition for inorganic nutrients such as N and P between soil microbes and plant roots [46], microbial communities in Stage F did not seem to be primarily limited by N or P. This result would again emphasize the importance of an easily available C source (energy source) for soil microbial community growth and activity in the late stage of succession. Many previous studies reported that SOM (or soil CN contents) increased [14,47] but its quality (microbial availability) tended to decrease [7,48] with succession. In addition, shifts in microbial characteristics along successions showed a somewhat consistent trend: microbial biomass increased and the community structure shifted from being bacteria-dominated with relatively high respiration rate per biomass (i.e., low C use efficiency) to being fungaldominated with relatively low respiration rate per biomass (i.e., high C use efficiency) [48e50]. Change in the type of substrate limitation from CN limitation to labile C limitation along successional gradients was reported in various ecosystems such as high arctic glacier foreland [14], dry hot desert [16], highly acidic solfatara [25], and volcanic desert (this study). These common trends found in soil development, microbial succession and shifts in the

43

type of substrate limitation imply that there is a consistent trajectory in microbial community development regardless of the ecosystem type and that substrate limitation is a key and universal mechanism in understanding this successional trajectory. 5. Conclusions Soil microbial respiration rate and biomass in the early stage of succession increased after CNP amendment and reached the same level as that in the late stage of succession. These results support our first hypothesis that substrate limitation would be an important factor regulating the development of the soil microbial community. Our amendment experiment using three types of C sources indicated that the different functional groups of soil microorganisms were limited by different types of substrate limitation. Our second hypothesis that the type and extent of substrate limitation would change along the succession was also supported. The microbial community suffered from CN limitation in the early stage of succession, whereas it was regulated more by labile C limitation in the late successional stage. This common trend in soil and microbial development along the succession observed in various ecosystem types implies that there is a consistent trajectory in microbial community development regardless of the ecosystem type and that substrate limitation is a key and universal mechanism in understanding this successional trajectory.

Table A.1 Results of ANOVA (P values) showing the effects of C, N, and P amendment on microbial respiration rate (Stage B soils).

C1 C2 C3 N P C1:N C2:N C3:N C1:P C2:P C3:P N:P C1:N:P C2:N:P C3:N:P

Day 0

Day 1

Day 2

Day 5

Day 10

Day 15

Day 20

Day 25

0.134 <0.001*** <0.001*** <0.001*** <0.001*** 0.018* 0.166 0.737 0.080 <0.001*** <0.001*** <0.001*** 0.308 0.086. 0.009**

0.002** <0.001*** 0.456 0.025* 0.019* 0.473 <0.001*** 0.021* 0.149 0.317 0.284 0.190 0.995 0.170 0.456

<0.001*** <0.001*** 0.264 <0.001*** <0.001*** <0.001*** <0.001*** 0.205 <0.001*** <0.001*** 0.846 <0.001*** <0.001*** <0.001*** 0.064

<0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** 0.083 <0.001*** <0.001*** <0.001*** 0.002**

<0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** 0.005** <0.001*** <0.001*** <0.001*** 0.010*

<0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** 0.016* 0.373 <0.001*** 0.629 <0.001*** 0.192 <0.001*** 0.563

<0.001*** <0.001*** 0.277 <0.001*** <0.001*** <0.001*** <0.001*** 0.999 0.002** <0.001*** 0.914 <0.001*** <0.001*** <0.001*** 0.862

<0.001*** <0.001*** 0.586 <0.001*** 0.016* <0.001*** <0.001*** 0.840 0.004** <0.001*** 0.895 0.010** 0.002** <0.001*** 0.941

C1 glucose, C2 cellulose, C3 lignin, N ammonium nitrate (NH4NO3), P potassium dihydrogen phosphate (KH2PO4). ***P < 0.001, **P < 0.01, *P < 0.05.

Table A.2 Results of ANOVA (P values) showing the effects of C, N, and P amendment on microbial respiration rate (Stage F soils).

C1 C2 C3 N P C1:N C2:N C3:N C1:P C2:P C3:P N:P C1:N:P C2:N:P C3:N:P

Day 0

Day 1

Day 2

Day 5

Day 10

Day 15

Day 20

Day 25

<0.001*** <0.001*** <0.001*** <0.001*** 0.087 0.766 0.556 0.662 0.655 0.450 0.009** 0.082 0.582 0.515 0.536

<0.001*** 0.003** <0.001*** 0.005** <0.001*** 0.210 0.206 0.027* <0.001*** 0.060 0.216 0.063 0.006** 0.444 0.717

<0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** 0.212 <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** 0.013* <0.001***

<0.001*** 0.069 <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** 0.726 <0.001*** <0.001*** <0.001*** 0.907

<0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** <0.001*** 0.460 <0.001*** <0.001*** 0.423 <0.001*** <0.001*** <0.001*** 0.709

<0.001*** 0.001** 0.010* <0.001*** <0.001*** <0.001*** <0.001*** 0.707 <0.001*** 0.002** 0.874 0.013* 0.121 0.022* 0.625

<0.001*** <0.001*** 0.148 <0.001*** <0.001*** <0.001*** 0.002** 0.491 <0.001*** 0.098 0.843 0.808 0.169 0.086 0.805

<0.001*** <0.001*** 0.371 <0.001*** <0.001*** <0.001*** 0.009** 0.751 <0.001*** 0.071 0.679 0.013* <0.001*** 0.499 0.990

C1 glucose, C2 cellulose, C3 lignin, N ammonium nitrate (NH4NO3), P potassium dihydrogen phosphate (KH2PO4). ***P < 0.001, **P < 0.01, *P < 0.05.

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S. Yoshitake et al. / European Journal of Soil Biology 73 (2016) 34e45

Table A.3 Results of ANOVA showing the effects of C, N, and P amendment on microbial biomass (total PLFA contents). P values

C1 C2 C3 N P C1:N C2:N C3:N C1:P C2:P C3:P N:P C1:N:P C2:N:P C3:N:P

Stage B

Stage F

Day 0

Day 10

Day 15

Day 0

Day 5

Day 15

0.009** <0.001*** <0.001*** 0.028* 0.002** 0.748 0.504 0.990 0.281 0.212 0.864 0.878 0.706 0.591 0.156

<0.001*** <0.001*** 0.688 0.000*** <0.001*** <0.001*** <0.001*** 0.817 <0.001*** <0.001*** 0.957 <0.001*** <0.001*** <0.001*** 0.976

<0.001*** <0.001*** 0.090. <0.001*** <0.001*** <0.001*** <0.001*** 0.168 <0.001*** <0.001*** 0.752 <0.001*** <0.001*** <0.001*** 0.542

0.006** 0.329 0.010* 0.460 0.003** 0.537 0.191 0.933 0.233 0.087. 0.002** 0.780 0.173 0.951 0.167

<0.001*** <0.001*** 0.894 <0.001*** <0.001*** <0.001*** <0.001*** 0.025* <0.001*** 0.002** 0.730 <0.001*** <0.001*** <0.001*** 0.019*

<0.001*** 0.946 <0.001*** <0.001*** 0.088 <0.001*** 0.127 0.651 <0.001*** 0.570 0.501 0.994 0.002** 0.322 0.737

PLFA phospholipid fatty acid, C1 glucose, C2 cellulose, C3 lignin, N ammonium nitrate (NH4NO3), P potassium dihydrogen phosphate (KH2PO4). ***P < 0.001, **P < 0.01, *P < 0.05.

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