Soil microbial and plant responses to the absence of plant cover and monoculturing in low arctic meadows

Applied Soil Ecology 48 (2011) 142–151

Contents lists available at ScienceDirect

Applied Soil Ecology journal homepage: www.elsevier.com/locate/apsoil

Soil microbial and plant responses to the absence of plant cover and monoculturing in low arctic meadows Minna-Maarit Kytöviita a,∗ , Anne Pietikäinen a , Hannu Fritze b a b

Department of Biological and Environmental Science, Jyväskylä University, P.O. Box 35, FIN-40014, Finland Finnish Forest Research Institute, Vantaa Research Centre, P.O. Box 18, FIN-01301 Vantaa, Finland

a r t i c l e

i n f o

Article history: Received 2 March 2010 Received in revised form 23 February 2011 Accepted 27 March 2011 Keywords: Microbial community Erosion Soil nutrient levels Arbuscular mycorrhizal colonization Dark septate fungi Facilitation

a b s t r a c t Arctic ecosystems are sensitive to disturbance yet there is little information on the fate and recovery of soil microbial communities after disturbance and persistence in the absence of plants. Neighbouring plants may facilitate seedling establishment through amelioration of the physical environment and maintenance of arbuscular fungal community in soil. The inoculum of arbuscular mycorrhizal symbionts is critical for the establishment of low-latitude arctic herbs that are obligately mycorrhizal. In the present work, we investigated plant–soil and plant–plant interactions in low arctic meadow habitat. Plant cover was experimentally removed and field plots were left without plant cover, or a monoculture of the common local herb, Solidago virgaurea, was planted on the plots or the plant cover was left intact. After two years, three herb species were sown on the plots and their growth, mycorrhizal colonization and soil microbial communities were measured. Relative fungal abundance decreased in the soil community in the treatments where the soil had been disturbed. Soil microbial community structure by functional group was maintained in the soil two years after no plant cover and microbial biomass per organic matter was not reduced. Mycorrhizal colonization potential was not impaired after two years of absence of host plants and thus it is concluded that symbiotic propagules are able to persist for two years in the absence of host plants. The Solidago monoculture did not facilitate seedling establishment nor change soil microbial community markedly in short-term. Overall, the low arctic soil microbial community was markedly resistant to disturbance. © 2011 Elsevier B.V. All rights reserved.

1. Introduction Arctic areas are especially vulnerable to disturbances because of their harsh climate and low ecosystem productivity. The arctic is increasingly exploited by humans for extraction of industrial raw materials and for recreation. Increasing numbers of roads are constructed, and off-road vehicle use is increasing (Ives, 1974). Furthermore, trampling and grazing by large stocks of semidomesticated reindeer destroy the thin layer of vegetation in the most sensitive areas leaving the soil exposed to erosion (Kashulina et al., 1997; Loeffler, 2002). Mining activities result in large areas without vegetation and organic matter and the revegetation process in these areas resembles that of primary succession (Cargill and Chapin, 1987). Industrial activities in the extreme north result in square kilometres of land with stripped vegetation that leaves the soil vulnerable to physical erosion by wind and water (Kashulina et al., 1997). Precipitation is predicted to increase in the Arctic (Kattsov et al., 2005) and thus increase soil erosion rates. Despite

∗ Corresponding author. Tel.: +358 14 260 2293; fax: +358 14 260 2321. E-mail address: minna-maarit.kytoviita@jyu.fi (M.-M. Kytöviita). 0929-1393/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.apsoil.2011.03.011

intensive research in arctic and alpine ecosystems, integrated information on plant and soil responses to erosion and disturbance in cold climate habitats is limited (Jorgenson and Joyce, 1994). Recovery of vegetation depends on the severity of disturbance, the distance to the nearest source of new potential colonizers and local environmental conditions. Small patches experimentally denuded of vegetation are rapidly revegetated by clonal growth of neighbouring vegetation in subarctic systems (Olofsson et al., 2005). In tussock tundra not susceptible to erosion, vegetation recovered naturally within 10 years after experimentally removing plants from a 20 m × 50 m area, but leaving the underlying 10–20 cm organic soil layer (Chapin and Chapin, 1980). A degraded arctic system does not always return to the original state even under restoration efforts. Due to the complex feedbacks between biotic and abiotic factors, eroded systems may form a stable alternative state (Suding et al., 2004). Little information is available on the fate and recovery of the soil microbial community after disturbance in the arctic. Loss of plant cover and loss of organic matter inevitable in the process of erosion reduce the available C for saprophytic microorganisms. On the other hand, carbon stocks in arctic soils are formidable (McKane et al., 1997) and carbon availability does not seem to limit the

M.-M. Kytöviita et al. / Applied Soil Ecology 48 (2011) 142–151

microbial activity (Mack et al., 2004) and growth (Schimel and Weintraub, 2003) in undisturbed soils. However, it may be that it is not the low availability of carbon per se, but the carbon quality that limits microbial activity in tundra soils (Jonasson et al., 1999). Defoliation and thus reduction in plant C assimilation capacity reduces microbial respiration in arctic meadow indicating that at least a significant proportion of microorganisms are dependent on carbon in the form of root exudates (Stark and Kytöviita, 2006). Furthermore, although reduction in plant cover reduced competition for nutrients, nitrogen did not accumulate in microbial biomass in eroded arctic salt marsh soils (Buckeridge and Jefferies, 2007), suggesting that nutrient availability did not limit the microbial biomass. This indicates that other factors such as C availability or activity of organisms at other trophic levels in soil controlled microbial abundance. It is of great interest, especially in reference to restoration efforts to know how long the C stocks in soil maintain appreciable microbial populations in the absence of plants. Arbuscular mycorrhizal (AM) fungi are an important component of the soil microbial community. AM symbiosis is common in low arctic herbs, although the high arctic climate seems to discourage AM fungi (Kytöviita, 2005). In this symbiosis, the fungus efficiently explores the soil for water and nutrients which are exchanged for plant photosynthetates (Allen, 1991; Smith and Read, 1997). It has been suggested that AM may be important in restoration and rehabilitation of nutrient poor ecosystems (Reeves et al., 1979; White et al., 2008). AM fungi may improve plant performance in eroding soils through directly assisting plant in nutrient and water uptake, and indirectly through stabilizing soil and therefore improving soil nutrient status and plant nutrient acquisition (Miller and Jastrow, 1992). AM fungi may be particularly important in eroded soils as these fungi produce soil stabilizing proteins, glomalins, in addition to binding soil particles directly with hyphae (Rillig and Mummey, 2006). While the AM fungi are important in establishment of plant cover after disturbance, disturbance itself reduces the abundance of the AM fungi (Allen et al., 1987; Moorman and Reeves, 1979; Reeves et al., 1979). The dispersal of AM propagules to disturbed sites is low as AM propagules have poor dispersal ability (Maffia et al., 1993). Therefore, the presence and persistence of AM in disturbed arctic soils is a prerequisite to plant establishment in these sites. Arctic plant species disperse outside their immediate vicinity by producing seeds, although clonal reproduction is important in local survival in some species. Seedling establishment is a critical stage in the plant life cycle and it is affected by both abiotic and biotic environment. Interactions such as those with AM fungi and neighbouring plants are crucial for seedling establishment. Seedling–neighbourhood interactions may be positive when established plants facilitate the establishment of recruits and plant growth as has been shown in highly disturbed and harsh environments (Callaway et al., 2002; Baumeister and Callaway, 2006; Choler et al., 2001). Facilitation is considered to take place through physical amelioration of the environment (Brooker and Callaghan, 1998; Padilla and Pugnaire, 2006). However, facilitation could also take place by established plants providing mycorrhizal inoculum (Nara, 2005) or by maintenance of functional saprophytic rhizosphere microorganisms that improve nutrient cycling and the whole plant community nutrient uptake. Plant–plant interactions may also be negative as is classically seen in plant competition experiments (e.g. Pietikäinen and Kytöviita, 2007; Dormann et al., 2004). Therefore, in restoration attempts where neighbouring plants are planted, both negative and positive interactions are probably involved between neighbouring plants and establishing seedlings, but the net outcome of these interactions is difficult to predict. In the present work, we investigated plant–plant and plant–soil interactions in disturbed, eroding soil in low arctic meadow habitat.

143

Plant cover was experimentally removed (no-plant treatment) and a common local herb, Solidago virgaurea, was planted on the plots (Solidago monoculture treatment) or the meadow plots were left intact (control treatment). After two years, three local herb species were sown on the plots and their growth, mycorrhizal colonization and soil microbial communities and nutrient levels were measured. Specifically, we hypothesised: (i) Microbial community will be different in the vegetation removal plots compared to plots with plant cover due to decline of microorganisms associated with plants such as the AM fungi that are obligatorily dependent on the living plant and those that prefer plant-derived carbon such as the Gram negative bacteria (Kramer and Gleixner, 2006, 2008). (ii) The mycorrhizal inoculum potential of the soil will be significantly reduced after two years of no plant cover due to lack of host plants. (iii) Compared with the absence of plants, the Solidago monoculture will facilitate seedling establishment due to maintenance of functional soil microbial population and AM networks that provide the seedlings with AM inoculum. 2. Materials and methods The experiment was conducted at Kilpisjärvi, NW Finland. Two low-latitude arctic meadows were chosen at 2.5 km apart to serve as landscape level replicates. Both meadows are about 600 m a.s.l. with southern exposure and referred to as Saana (69◦ 03 N, 20◦ 50 E) and Jehkas (69◦ 05 N, 20◦ 47 E) hereafter. Both sites are located in intensive summer grazing area of reindeer and the history of reindeer herding in this area can be dated back at least a few centuries. The vegetation in these sites is similar and dominated by the grass Deschampsia flexuosa, but sedges and herbs, such as Carex bigelowii, C. vaginata, S. virgaurea, Trollius europaeus, Potentilla crantzii, and Bistorta vivipara, are also common. Only few species of dwarf shrubs, such as Betula nana, Juniperus communis and Vaccinium myrtillus, occur. The length of the growing season is about 90 days in the area and the mean annual temperature is −2.56 ◦ C (1951–1985) and precipitation 422 mm (1961–1985) measured at Kilpisjärvi meteorological station situated at 483 m a.s.l. (Järvinen, 1987). The soil has an organic layer about 5 cm in depth and the soil temperature at 3–5 cm depth during the snow-free months, July and August, is 10.8 ◦ C (2000–2006, Kytöviita, unpublished). In the end of June 1999, nine experimental plots, 3.5 m in diameter were established on both meadows. First the plots were randomly allocated to three treatments: control, monoculture and no-plant. Thus, we had three replicates for each treatment per site. Then all the vegetation was removed from the monoculture and the no-plant plots. Most of the roots were extracted, but small fragments of fine roots could not be completely removed. When extracting the roots, the very top soil profile high in organic matter (5 cm) got mixed with the soil underneath in the monoculture and no-plant plots resulting in initially lower soil organic matter concentration in these two treatments in the beginning of the experiment (Kytöviita et al., MS). At this stage, these manipulations left the no-plant and the monoculture plots without any aboveground vegetation. The monoculture plots were revegetated in August 1999 by planting 100 mature S. virgaurea plants collected from the surrounding undisturbed meadow to each plot. These plots are referred to as monoculture plots hereafter. In the year 2002, the coverage of the Solidago in the monoculture plots was 8% of total area. The control plots were left untouched presenting the diversity and composition of natural vegetation with 100% vegetation coverage of total plot area. All the plots (including control plots) were covered with fine transparent mesh every year from

144

M.-M. Kytöviita et al. / Applied Soil Ecology 48 (2011) 142–151

mid August to early June to prevent the natural seed rain and the subsequent seedling establishment. All the plots were trenched and disconnected from surrounding soil and vegetation. All the plots were fenced to exclude the reindeer and remained without any further manipulations until June 2001. In June 15–16, 2001, seeds of S. virgaurea, Alchemilla glomerulans and P. crantzii were sown to each plot. These plants are common in the study site (Pietikäinen et al., 2007). The seed material was collected from the experimental meadows in September 1999 and stratified for 7 weeks in wet sterilized sand at 5 ◦ C prior to sowing to the experimental plots. On each plot, the seeds were sown in a line crossing the plot, lightly covered with soil and watered. We counted the seedlings repeatedly in 2001 and 2002 and the seedlings were thinned when necessary to avoid crowding. Only very few Potentilla and no Alchemilla germinated in 2001. However, Solidago germinated well during summer 2001 and very few seeds germinated 2002. Ten Solidago seedlings from each plot were sampled in August 13–14, 2001 and these represent young, maximally about two-month old seedlings. In August 9–11, 2002, 40 Solidago and Potentilla seedlings were harvested from each plot, except in one control plot at Jehkas there were only 9 Potentilla seedlings. There were fewer seedlings of Alchemilla and 18–30 seedlings were harvested in each plot, except in one control plot at Saana there were only 5 seedlings. The seedlings collected in 2002 represent young, maximally two and half month old seedlings in case of Alchemilla and Potentilla and one year and two months old in case of Solidago. The roots were gently washed clean of soil and stored in 50% ethanol until analysed for fungal colonization intensity. The dry weight of the shoots (80 ◦ C overnight) was recorded and the shoot N concentration analysed (CE Instruments EA 1110 Elemental Analyzers). For fungal colonization intensity in Solidago seedlings 2001, 10 plants were collected per plot 14–16 in August. Per plot, 10–20 root fragments of 1–3 cm length were included in the analyses. For fungal colonization intensity in Solidago, Alchemilla and Potentilla seedlings in the year 2002, 17 individual plants were harvested per plot (except in two plots where the material was less as specified above). Per plot, 17–30 root fragments were analysed. Fungal colonization intensity in the roots was assessed under a light transmission microscope using the gridline intersection method (McGonigle et al., 1990) after staining with trypan blue (Phillips and Hayman, 1970). For each root fragment, 20 intersections were scored for the presence of arbuscular mycorrhizal structures (hyphae, arbuscules, vesicles and fine endophyte), of dark septate (DS) fungi and yeasts. Fine endophytic structures were identified due to their very thin and intensively staining hyphae that form a characteristic morphology (Gianinazzi-Pearson et al., 1981). Fine endophytic colonization is assigned to the AM fungus Glomus tenuis, although there is some uncertainty whether several fungi are associated or not (Brundrett et al., 1996). Dark septate endophytes are a taxonomically diverse range of fungi belonging to Ascomycota (Addy et al., 2005). They are characterised by formation of non-staining, dark, pigmented and septate hyphae and sclerotia in plant roots. Root colonization by other fungi was negligible. Soil samples were collected each year in the beginning of the growing season in June and in the end part of the growing season in August (15–16 June 2001, 9 August 2001, 9–10 June 2002 and 12–13 August 2002). A bulk sample consisting of 12 soil cores (diameter 3 cm, 6 cm length) was taken from each plot. The soil samples were transferred to the laboratory and immediately sieved through 2 mm sieve, frozen and kept frozen until the PLFA and nutrient analyses. The phospholipid fatty acids (PLFAs) were extracted from 1 g fresh weight (Frostegård et al., 1993, modified by Pennanen et al., 1999). Briefly, the humus was extracted with 1.9 ml chloroform, 3.75 ml methanol and 2 ml Blight and Dyer mix-

ture (Blight and Dyer contains chloroform:methanol:citrate buffer [0.15 M; pH 4] at volume ratios 1:2:0.8), and the lipids were separated into neutral lipids, glycolipids, and phospholipids in a silicic acid column. The phospholipids were subjected to mild alkaline methanolysis, and the fatty acid methyl esters were separated by gas chromatography (Hewlett Packard 5890) equipped with a flame ionisation detector and a HP-5 (phenylmethyl silicone) capillary column, 50 m in length, using He (30 ml min−1 ) as a carrier gas. The peak areas were quantified by adding methyl nonadecanoate fatty acid (19:0) as an internal standard, and the peaks were identified by using HP ChemStation software. 40 PLFAs were identified. Of these, the following were considered to indicate bacteria: i15, a15, 15:0, i16, 16:1␻9, 16:1␻7t, i17, a17, 17:0, cy17, 18:1␻7, cy19. Gram positive bacterial signal PLFAs were a15, i16, i17, and Gram negative 16:1␻7t/c, 16:1␻5, cy17, 18:1␻7, 19:1a. The methylated PLFA 10Me16, 10Me17 and 10Me18 were considered as actinomycete indicators. The 18:2␻6,9 PLFA was used as the indicator of fungal abundance (Ascomycota and Basidiomycota) and the 16:1␻5 as the indicator of AM fungal abundance (Glomeromycota). The following physico-chemical variables were analysed on the soil collected in August 2001 and 2002: pHH2 O , both extractable ions (P, Ca, Mg, Mn, S, Fe, Al, and Na) and dissolved organic carbon (DOC) and nitrogen (DON), which could be eluted into 100 ml water from 2 g dry soil by shaking for 2 h at 200 rpm was measured after filtrating the solution through a <2 ␮m paper filter. Extractable ions were analysed by inductively coupled plasma atomic emission spectrometer (TJA Iris-Advantage) and DOC/DON with an Analytic Jenan MULTI-nitrogen-carbon Analyser. C, N, and C/N ratio, were analysed according to descriptions given in Tamminen and Starr (1990) except that total C and N were determined with a LECO carbon-hydrogen-nitrogen-1000 Analyser. Organic matter content was analysed by loss-on-ignition at 550 ◦ C for 4 h. 2.1. Statistical analyses Mean values per plot were used as the statistical variates when several measurements had been made per plot at the same time. The significance of treatment and site effects on plant biomass, N concentration and N content was analysed with two factor ANOVA with treatments (control, monoculture, and no plant) and site (Saana and Jehkas) as fixed factors followed by Tukey’s multiple comparison test. The effect of treatments (control, monoculture, and no-plant) and site (Saana and Jehkas) on fungal colonization parameters (root length colonized by AM hyphae, arbuscules, vesicles, fine endophyte, DS fungi and yeasts) was tested with two factor ANOVA for the three plant species separately. Solidago colonization data was also tested with repeated measures ANOVA with time (2001 and 2002) as the within subject factor. Since arbuscular frequency was significantly affected by time, the two years were tested separately with two factor ANOVA. The significance of treatment and site effects and sampling time on the total amount of PLFA extracted, and the mole% of single or pooled signature PLFAs and bacterial to fungal ratios were analysed with repeated measures ANOVA with time as within subject factor (June 2001, August 2001, June 2002, and August 2002) and treatment (control, monoculture, and no-plant) and site (Saana and Jehkas) as the between subject factors. The mole% of the signature PLFAs included in the repeated measures ANOVA were the AM fungal indicator 16:1␻1, sum of the bacterial Gram positive signature PLFAs (a15, i16, and i17), sum of the Gram negative signature PLFAs (16:1␻7t/c, cy17, 18:1␻7, and 19:1a), and sum of the actinomycete indicator PLFAs (10Me16, 10Me17, and 10Me18). The relative abundances of the 40 identified PLFA were reduced to two principal components, PLFA-PC-1 and PLFA-PC-2, which explained 44% and 10% of the variation, respectively. These two

M.-M. Kytöviita et al. / Applied Soil Ecology 48 (2011) 142–151

145

Table 1 Soil variables in the experimental sites Saana (S) and Jehkas (J). Mean and one standard error (SE) values of soil samples collected in August 2001 and August 2002 are shown for the control (C), monoculture (M) and no-plant (No) treatments, n = 12. Statistically non-significant (ns) and significant differences between the treatments (Trt) and the sites (Site) are indicated according to repeated measures ANOVA followed by Tukey’s multiple comparison test. Saana

Jehkas

Control Mean pH OM (%) DOCa DONa C (g g−1 OM) N (mg g−1 OM) Soil C (%) Soil N (%) C/N Pa Caa Mga Mna Sa Fea Ala Naa a

4.88 7.92 560 66 394 31 3.3 0.24 13.8 5.25 36.2 10.3 11.4 66 12.9 16.6 13.9

SE 0.03 0.75 70 10 8 1 0.3 0.02 0.3 0.65 4.35 1.3 2.8 4 1.0 1.5 1.5

Monoculture

No-plant

Mean

Mean

5.20 4.94 179 22 400 33 2.0 0.16 12.23 2.22 13.7 4.3 2.1 43 11.9 11.1 9.3

SE 0.07 0.56 17 2 8 1 0.3 0.02 0.2 0.07 2.6 0.5 0.3 4 1.9 1.2 1.0

5.06 4.72 185 29 394 32 1.9 0.15 12.4 2.14 21.67 6.1 2.5 49 11.8 12.1 12.13

Control SE 0.07 0.14 11 3 10 1 0.1 0.06 0.1 0.02 1.9 0.4 0.3 2 1.3 0.6 1.7

Mean 4.91 10.0 793 92 446 34 4.4 0.34 13.2 8.65 57.1 13.5 8.4 73 16.3 21.8 15.5

SE 0.03 1.51 81 7 16 1 0.5 0.04 0.3 0.83 5.4 1.2 1.3 6 1.6 1.4 1.3

Monoculture

No-plant

Mean

Mean

5.01 3.75 289 32 407 34 1.54 0.13 11.9 3.04 25.5 7.2 2.2 47 13.5 15.5 9.5

SE 0.02 0.19 23 3 12 1 0.1 0.01 0.4 0.24 3.6 0.7 0.4 4 0.5 0.4 1.2

4.86 4.17 270 45 376 32 1.6 0.14 11.5 2.70 34.8 9.7 2.8 55 14.3 17.1 15.13

SE 0.07 0.25 17 6 6 1 0.1 0.01 0.3 0.30 2.3 0.7 0.5 3 1.6 1.5 1.6

ns Trt (C > M, No) Trt (C > M, No), Site (J > S) Trt (C > M), Site (J > S) Trt (C > No) Site (J > S) Trt (C > M, No) Trt (C > M, No) Trt (C > M, No), Site (S > J) Trt (C > M, No) Trt (C > No > M) Trt (C > No > M), Site (J > S) Trt (C > M, No) Trt (C > No > M) ns Trt (C > M, No), Site (J > S) Trt (C, No > M)

mg kg−1 dry weight soil.

PCs were subjected to repeated measures ANOVA as above. The soil nutrient contents were reduced to one PC (nutrient-PC) that explained 68% of total variation. Since soil nutrient content PCs did not differ significantly between the years, we used the average values of the two measurement times (August 2001 and August 2002 to explore which soil parameters were responsible for overall differences between the sites and the treatments) and tested soil pH, organic matter fractions and nutrient concentrations with two-factor ANOVA. Site (Saana and Jehkas) and treatment (control, monoculture, and no-plant) were included as fixed factors and the ANOVA was followed by Tukey’s multiple range test to elucidate the differences between the three treatments. To explore the relationship between plant growth and root fungal colonization rates and the nutrient and microbial composition in the soil, regression analyses were conducted on 2002 August data where plant growth, N content and fungal colonization parameters were the dependent variables and PLFA-PC-1 and soil nutrient-PC-1 on August 2002 data and plant coverage in the treatments were the independent variables. Variables were log-transformed to gain normal distribution and variance homogeneity when necessary, but proportional variables (fungal frequency, mole%) were arc sin square root transformed. All analyses were conducted with SPSS version 16.0.

3. Results 3.1. Soil nutrient levels Overall, reducing plant cover reduced nutrient levels in the soil (Table 1). Also the amounts of DON and DOC per soil dry weight and soil C% and N% were reduced in these treatments (Table 1). The amount of carbon in the soil organic matter was 42% under control conditions and declined in the monoculture and no-plants plots (Table 1). The soil pH was close to 5, and did not differ significantly between sites or treatments (Table 1). Despite having no differences in soil pH, the Jehkas site was more nutrient rich than the Saana site and had higher DOC, DON, organic matter N content, Mg and Al concentrations (Table 1). The soil C% and N% was not significantly affected by the site, but C/N ratio was significantly higher in the Saana site. Overall, soil nutrient levels were not affected by the year of collection (data not shown).

3.2. PLFA profiles The amount of total PLFA extracted in control samples declined with time, but was nevertheless significantly higher than in the no-plant and monoculture soils (Fig. 1a). The total amount of PLFA as nmol g−1 organic matter did not show clear treatment or time trends (Fig. 1b). The within treatment microbial composition remained stable through the two years as indicated by the results of repeated measures ANOVA on PLFA-PC-1 and PLFA-PC-2 on the relative abundance of the 40 individual PLFAs (statistics not shown). The fungi were more abundant in control soils than in the monoculture or no-plant soils (Figs. 1c and 2). The AM fungal signature PLFA 16:1␻5 was significantly (p < 0.001) more abundant in the control soil than in the other two treatments, but the monoculture plots did not differ from the no-plant plots (Fig. 1d). The pooled Gram positive bacteria indicator PLFA was significantly lower in the control plots in comparison to the other two disturbed treatments (p < 0.001, Fig. 2), and significantly higher in the Jehkas site than in the Saana site (p < 0.002). The pooled Gram negative bacteria indicator had the opposite response, and it was significantly higher in control than in the disturbed treatments (p < 0.05) while there was no difference between the sites (Fig. 2, p = 0.379). The relative abundance of actinomycetes was marginally affected by treatments (Fig. 2, p = 0.072), and significantly higher in the Jehkas site (p = 0.016). Altogether, the control plot microbial community was significantly different from the monoculture and the no-plant plots, but monoculture did not result in significant changes compared to no-plant treatment (Figs. 2 and 3, statistics not shown). 3.3. Mycorrhizal colonization Only Solidago seedlings germinated in large numbers 2001 and therefore two year sampling was possible. The seedlings became highly colonized already in their first year in all treatments (Fig. 4a, c, and d). In 2002, the Solidago roots were significantly more intensively colonized by vesicles and DS fungi in control plots than in the two other treatments (Fig. 4b). Also in Alchemilla, the roots contained significantly more AM fungal vesicles in control plots (Fig. 4c). However, in Potentilla roots, the frequency of fine endophyte and arbuscules was the lowest in the control plots (Fig. 4d). The site did not affect significantly any of the fungal colonization

146

M.-M. Kytöviita et al. / Applied Soil Ecology 48 (2011) 142–151

A) total nmol PLFA / g dwt soil

Trt p < 0.003 Time p = 0.001 Time x Trt p < 0.001

450

control

400

monoculture

no-plant

350 300 250 200 150 100 50 0

5000 4500 4000 3500 3000 2500 2000 1500 1000 500 0

control

June 2001 Aug 2001 June 2002 Aug 2002 C) bacterial to fungal PLFA ratio 35

Trt =p 0.006 Time p = 0.015 Trt x Time p = 0.002

5 4,5 4 3,5 3 2,5 2 1,5 1 0,5 0

20 control monoculture no-plant

5

no-plant

D) mole % AM signature PLFA

25

10

monoculture

June 2001 Aug 2001 June 2002 Aug 2002

30

15

Trt p = 0.100 Time p = 0.343 Time x Trt p = 0.019

B) total nmol PLFA / g OM

0

June 2001 Aug 2001 June 2002 Aug 2002

control

Trt p< 0.001 Time p = 0.073 Trt x Time p = 0.248

monoculture

no-plant

June 2001 Aug 2001 June 2002 Aug 2002

Fig. 1. Phospholipid fatty acids (PLFAs) measured in the treatments (Trt) control, monoculture and no-plant plots at the four sampling occasions, the error bars indicate one standard error. (A) Total amount of PLFA nmol g−1 dry weight soil, (B) total amount of PLFA nmol g−1 organic matter in soil, (C) bacterial to fungal PLFA ratio, (D) amount of the arbuscular mycorrhiza signature PLFA 16:1␻5 in the control, monoculture and no-plant soils at the four sampling occasions. Repeated measures ANOVA results are shown in the graph.

parameters in any of the species (data not shown). In regression analyses, the PLFA PC-1 did not significantly explain the fungal colonization in any of the plant species. In Potentilla, the frequency of arbuscules and hyphae were significantly explained by the pooled nutrient profile index PC-1 (p < 0.001, r2 = 0.41 and r2 = 0.69, respectively). DS fungal frequency responded to nutrients, but also relative coverage by plants explained significantly their frequency in Solidago and Alchemilla roots (p < 0.01, r2 = 0.45 and r2 = 0.72, respectively).

Potentilla seedlings was markedly lower at the Saana site when compared to the more nutrient rich Jehkas site (Fig. 5a and c). The N% in plants was significantly higher in the no-plant plots than in the control plots (data not shown). The total N content of the seedlings reflected the N% and growth differences and was significantly higher in the no-plant plots when compared to control plots (Fig. 6). Only Solidago seedlings growing in the monoculture plots at the Saana site accumulated significantly less N when compared to the same treatment at Jehkas, although the mean N accumulation in all seedling species in the Saana monoculture plots was low (Fig. 6a). The coverage of competing plant foliage explained seedling biomass accumulation significantly in all cases (p < 0.01, r2 = 0.28 for Potentilla, r2 = 0.35 for Alchemilla, r2 = 0.89 for Solidago).

3.4. Seedling biomass and [N] In all plant species, the biomass accumulation was lowest when growing in the control plots (Fig. 5). The growth of Solidago and

Relative abundance (mole %)

12

control monoculture no-plant

10

8

6

4

2

Gram +

Gram Bacteria

Actinomycetes

18:2ω6

16:1ω5

10Me18

10Me17

10Me16

cy19

a17

17:0

15:0

i15

cy17

19:1a

18:1ω7

16:1ω7t

16:1ω9

i17

i16:0

a15

0

Fungi

AM fungi

Fig. 2. Selected individual phospholipid fatty acids in the control, monoculture and no-plant plots. Data from the four sampling occasions (June 2001, August 2001, June 2002, and August 2002) and the two sites (Saana and Jehkas) are pooled. The error bars indicate one standard error.

M.-M. Kytöviita et al. / Applied Soil Ecology 48 (2011) 142–151

2 1,5

PLFA PC-1

1 0,5 0 -2

-1

0

1

2

3

-0,5 -1 -1,5 -2 Nutrient PC-1

Fig. 3. The principal components on the relative abundance of individual PLFA (PLFA PC-1) and soil nutrient levels (nutrient PC-1) in the three treatments at the two sites (open symbols = Saana, filled symbols = Jehkas). Squares = control, triangles = monoculture, circles = no-plant plots. Average component values and their standard errors given for the two sampling dates are shown for the two sites separately, n = 6. PLFA PC-1 explained 44% and nutrient PC-1 explained 68% of the total variation.

4. Discussion Microbial community composition responds to changes in plant community composition such as presence of invasive species in temperate systems (Callaway et al., 2004) and monocropping (Garbeva et al., 2008). In our low arctic habitat, the microbial community composition responded to monoculture and absence of plant cover as indicated by changes in PLFA profile. The relative abundance of fungi decreased in the two treatments, monoculture and no-plant, where soil was disturbed. This finding is in line with many reports of reduced fungal abundance in mechan-

147

ically disturbed soils in temperate regions (e.g. Allison et al., 2005). In addition to mechanical disturbance, also lower levels of plant litter may have contributed to the decline in fungal abundance in monoculture and no-plant plots. Gram negative bacteria respond strongly to easily assimilable C inputs such as rhizodeposition (Griffiths et al., 1999) while Gram positive bacteria seem to use more soil organic matter derived carbon sources (Kramer and Gleixner, 2008). Therefore the relative increase in the Gram negative indicator 18:1␻7 in plots that had remained two years unvegetated is unexpected. However, differential responses to carbon source within Gram negative bacteria have been noted previously (Griffiths et al., 1999). Compared to a report of abundance of microorganisms in arctic tussock tundra, birch-willow tundra and sedge tundra (Zak and Kling, 2006), the present low arctic meadow soil had higher relative abundance of actinomycetes. When comparing different arctic habitats, actinomycete abundance was highest in glacier forefront (Zabavski and Zuravska, 1977) indicating adaptation to soil conditions prevalent in eroded soils. Also Peacock et al. (2001) report high relative abundances of actinomycetes in disturbed temperate soils. Similarly, in our case, the actinomycete marker 10Me16 was higher in the disturbed soils, although the group index of actinomycetes did not respond to treatments. Fungi dominate the microbial biomass in organic horizons of arctic and subarctic soils (Schmidt and Bölter, 2002), which must be the case in the present meadow habitat as well. Microbial population has strong seasonal shifts in cold climate soils (Björk et al., 2008; Schadt et al., 2003). In the present study, the microbial community did not change markedly between early (June) and late (August) growing season as indicated by the stability of the total amount and composition of the PLFA extracted. The PLFA analyses do not give indication of the absolute amounts of fungi and bacteria, but only comparative information concerning the effects of the applied treatments. The microbial biomass was significantly reduced in the no-plant plots and in the monoculture plots with relatively sparse (10 Solidago plants per m2 ) vegetation cover. This

Fig. 4. Average fungal colonization intensity in the seedlings grown in the control, monoculture and no-plant plots. Mean ± SE frequency of hyphae of arbuscular mycorrhizal fungi (Hyphae), arbuscules (Arbuscules), vesicles (Vesicles), dark-septate fungi (DSF), yeasts (Yeasts) and fine endophyte (FE) in roots of Solidago (A and B), Alchemilla (C) and Potentilla (D) are shown. Letters indicate significant differences between means (ANOVA followed by Tukey’s multiple test, p < 0.05). The error bars indicate one standard error.

148

M.-M. Kytöviita et al. / Applied Soil Ecology 48 (2011) 142–151 Trt F(2,12)=48.261, p= 0.000 Site F(1,12)=6.713, p= 0.024 TxS F(2,12)= 4.489, p= 0.028

A) Solidago

average seedling biomass (mg)

b

Saana Jehkas

300

average seedling biomass (mg)

average seedling biomass (mg)

350

b

250

b

200 150 100

a

a a

50 0

control

monoculture

no-plant

Trt F(2,12)=6.856, p= 0.010 Site F(1,12)=12.920, p= 0.004 TxS F(2,12)= 3.314, p= 0.071

C) Potentilla 35 30 25

b

Saana Jehkas

b

20 15 10

a

5 0

control

monoculture

no-plant

Trt F(2,12)=23.55, p= 0.000 Site F(1,12)=0.816, p= 0.384 TxS F(2,12)= 3.014, p= 0.087

B) Alchemilla 12

Saana 10

b

b

Jehkas

8 6 4

a

2 0

control

monoculture

no-plant

Fig. 5. Average seedling biomass (mg dry weight per seedling) in the treatments (Trt) control, monoculture and the no-plant plots at the end of the experiment in August 2002 in the two sites Saana and Jehkas. Factorial ANOVA results are shown in the graph. When significant interactions occurred, Tukey’s multiple test was employed and the significantly different groups are indicated with different letters. The error bars indicate one standard error.

was expected as the absence of plants should result in shortage of easily assimilable carbon in form of rhizodeposition as well as in reduced amounts of more complex C source in form of root litter. However, microbial biomass per organic matter was not affected Trt F(2,11) = 58.016, P<0.001 Site F(1,11) = 11.434 P=0.005 TxS F(2,11) = 5.637, P=0.019

A) Solidago

10000

b

Saana Jehkas

b

8000

b

6000 4000 2000

a

a

a

control

monoculture

1000

Saana Jehkas

d

800 600

cd

400 200

bc a

a

ab

no-plant

control

monoculture

no-plant

Trt F(2,11) = 21.204, P<0.001

B) Alchemilla

Site F(1,11) = 1.798, P=0.207 TxS F(2,11) = 3.130, P=0.084

1200

total N per plant (μg)

1200

0

0

1000

Trt F(2,11) = 36.499, P<0.001 Site F(1,11) = 21.671 P=0.001 TxS F(2,11) = 4.784, P=0.032

C) Potentilla

total N per plant (μg)

total N per plant (μg)

12000

by the removal of plants, which suggests that the rhizodeposition was not as important as the root litter or older soil organic matter as a source of C for the microorganisms. Tundra soils are known to contain large proportions of labile carbon (Sjögersten et al., 2003)

Saana Jehkas

800 600 400 200

b

b

a

0

control

monoculture

no-plant

Fig. 6. Average seedling N content (␮g N per seedling) in the treatments (Trt) control, monoculture and the no-plant plots at the end of the experiment in August 2002 in the two sites Saana and Jehkas. Factorial ANOVA results are shown in the graph. When significant interactions occurred, Tukey’s multiple test was employed and the significantly different groups are indicated with different letters (p < 0.05). The error bars indicate one standard error.

M.-M. Kytöviita et al. / Applied Soil Ecology 48 (2011) 142–151

that could be used to fuel microbial activities in the absence of plant litter and exudates. In previous studies in temperate systems, ‘old’ carbon in soil is known to fuel considerable microbial activities decades after the absence of plants (Kelly et al., 1996) and this seems likely in our low arctic case as well. In contrast to our original hypothesis, arbuscular mycorrhizal fungal colonization rates in the seedlings growing in plots that had been left for two years without plant cover was overall comparable to seedlings growing among intact vegetation. This clearly indicates that the AM fungal propagules were able to survive two years in the soil without host plant. Mycorrhizal fungi have been shown to persist considerable periods also in eroded prairie soils (White et al., 2008). The source of AM inoculum was most likely only spores in these low arctic vegetation free plots (Pietikäinen et al., 2007). The AM-signature PLFA was considerable and did not decline in the no-plant plots within the time span (June 2001–August 2002) examined in this study suggesting (i) non-specificity of the signal and consequently a large contribution of 16:1␻5 of bacterial origin (Hedlund, 2002), (ii) 16:1␻5 is extracted from the AM spores in the soil, or (iii) stable amount of living AM hyphae in the no-plant plots. There were certainly AM spores in the no-plant plots, but the method used to extract the PLFA does not extract the fatty acids in AM spores (Olsson and Johansen, 2000). Also, in the view of the obligatorily biotrophic nature of the AM fungi, considerable amounts of living hyphae after two years in the absence of host plants seem improbable. Therefore, the present results indicate that the PLFA 16:1␻5 in this low arctic site was to a notable extent of bacterial origin. The seedlings became highly colonized by AM already during their first year as shown previously for Solidago and Gnaphalium norvegicum (Pietikäinen et al., 2005). The colonization rates in Solidago changed only little during the second year suggesting that the AM colonization in this cold habitat is stable as observed previously for A. glomerulans and Trollius europaeus (Ruotsalainen et al., 2002). In Solidago and Alchemilla seedlings AM intensity in roots was not explained by nutrient availability. This conclusion agrees with an earlier study where fertilization did not affect the colonization levels in low arctic meadow seedlings (Pietikäinen et al., 2007). However, in Potentilla, AM colonization intensity seemed most responsive to treatments as well as it was explained by nutrient availability suggesting that AM colonization intensity response to environmental changes is host species specific. Fine endophytic colonization has been assigned to the AM fungus G. tenuis and it was the only AM fungus that could be identified by its morphological characteristics. Fine endophytic structures were observed frequently only in the roots of Potentilla. G. tenuis has been found in plants colonizing primary habitats (Daft and Nicholson, 1974). In line with G. tenuis occupying plant roots in primary habitats (Daft and Nicholson, 1974), fine endophytic colonization dramatically increased in the two disturbed treatments that represent earlier successional stages than the intact meadow vegetation. The possible mycorrhizal functions of dark septate (DS) fungi are unknown (Newsham, in press), but the DS fungi identified from plant roots have been phylogenetically closely related to known saprophytes (Vrålstad et al., 2002). The arctic and alpine plant roots are usually intensively colonized by this group of fungi (Pietikäinen et al., 2005; Read and Haselwandter, 1981; Treu et al., 1996). In the present work, DS fungal colonization in the seedling roots was explained by soil nutrient level and plant coverage in case of Alchemilla and Solidago, which may be explained by the saprophytic nature of DS fungi and their preference to soils with high organic matter content. We hypothesised that the monoculture plants might facilitate the establishment of seedlings geminating in their vicinity. Plant–plant interactions are reported to be more positive in harsh arctic environments and established plants facilitate establishing species (Klanderud and Tøtland, 2004; Carlsson and Callaghan,

149

1991). However, we could not find any positive effects in terms of biomass, N content or AM development of the monoculture on any of the seedling species studied compared with the no-plant treatment. Alchemilla and Potentilla seedlings grew equally well in the monoculture plots as in the no-plant plots. In addition, we expected the monoculture to have developed a specific AM fungal community in the soil that might assist the conspecific Solidago seedlings in particular. This expectation is based on the potential host specificity of some AM symbionts (Vandenkoornhuyse et al., 2003). However, the Solidago seedlings showed little response to the monoculture in terms of AM colonization intensity or in terms of AM community composition (Pietikäinen et al., 2007). Furthermore, the Solidago seedling growth and N acquisition was negatively affected by the established monoculture in the nutrient poor Saana site, which indicates strong intraspecific competition for resources. Also in prairie vegetation, competition between conspecifics has been shown to be stronger than between heterospecifics (Casper and Castelli, 2007). Established vegetation has been shown to impose strong competitive effects on seedlings in this low arctic area (Eskelinen, 2008), as well as in other cold climate experiments (Olofsson et al., 1999; Gough, 2006; Walker and Chapin, 1986). Although competition among seedlings is most frequently studied, competition between seedlings and established perennial vegetation is considered the major factor affecting seedling recruitment and vegetation composition (Walker and Chapin, 1986). Despite the higher nitrogen levels in control soils, seedling tissue N concentration and N acquisition were lower in these soils probably due to the intensive plant competition for soil resources. The vegetation cover was relatively low in the monoculture plots, the established monoculture plant canopy did not shade seedlings and the competition was confined to belowground. These field results are supported by a greenhouse experiment where low arctic adult plants competed strongly with seedlings for below-ground resources (Pietikäinen and Kytöviita, 2007). In conclusion, microbial biomass was related to amount of organic matter in the soil and not dependent on the presence or absence of plants in this short-term field experiment although microbial community composition changed when plant cover was either absent or low. Functional soil microbial community necessary for successful seedling establishment persisted in the soil for two years without vegetation cover. Contrary to the general concept of facilitation by neighbouring plants in harsh climates (Callaway et al., 2002), in the present low arctic meadows the net outcome of established plant-seedling interactions was either negligent or negative. It is concluded that plant–plant interactions are dominated by competition in the present low arctic meadow and that the microbial community is relatively resistant to perturbations in the short-term. Acknowledgements We wish to thank Kilpisjärvi Biological Station (University of Helsinki) for support during the field work, Petri Kärkkäinen for help during harvesting the field experiment, Arja Tervahauta (Finnish Forest Research Institute) for supervising the soil nutrient analyses and Tuulikki Pakonen (University of Oulu) for conducting the plant tissue [N] analyses. This research was funded by Kone Foundation and Academy of Finland (project numbers 1206981 and 7127657). References Addy, H.D., Piercey, M.M., Currah, R.S., 2005. Microfungal endophytes in roots. Can. J. Bot. 83, 1–13. Allen, M.F., 1991. The Ecology of Mycorrhizae. Cambridge University Press, Cambridge, 184 pp.

150

M.-M. Kytöviita et al. / Applied Soil Ecology 48 (2011) 142–151

Allen, E.B., Chambers, J.E., Connor, K.F., Millen, M.F., Brown, R.W., 1987. Natural reestablishment of mycorrhizae in disturbed alpine ecosystems. Arct. Alp. Res. 19, 11–20. Allison, V., Miller, J., Jastrow, R.M., Matamala, J.D., Zak, R.D., 2005. Changes in soil microbial community structure in tallgrass prairie chronosequence. Soil Sci. Soc. Am. J. 69, 1412–1421. Baumeister, D., Callaway, R.M., 2006. Facilitation by Pinus flexilis during succession: a hierarchy of mechanisms benefits other plant species. Ecology 87, 1816–1830. Björk, R.G., Björkman, M.P., Andersson, M.X., Klemedtsson, L., 2008. Temporal variation in soil microbial communities in Alpine tundra. Soil Biol. Biochem. 40, 266–268. Brooker, R.W., Callaghan, T.V., 1998. The balance between positive and negative plant interactions and its relationship to environmental gradients: a model. Oikos 81, 196–207. Brundrett, M., Bougher, N., Dell, B., Grove, T., Malajczuk, N., 1996. Working with Mycorrhizas in Forestry and Agriculture, Canberra, ACIAR Monograph 32, 374 pp. Buckeridge, K., Jefferies, R., 2007. Vegetation loss alters soil nitrogen dynamics in an arctic salt marsh. J. Ecol. 95, 283–292. Callaway, R.M., Brooker, R.W., Choler, P., Kikvidze, Z., Lortie, C.J., Michelet, R., Paolini, L., Pugnaire, F.I., Newingham, B., Aschehoug, E., Armas, C., Kikodze, D., Cook, B.J., 2002. Positive interactions among plants increase with stress. Nature 417, 844–848. Callaway, R.M., Thelen, G.C., Barth, S., Ramsey, P.W., Gannon, J.E., 2004. Soil fungi alter interactions between the invader Centaurea maculosa and North American natives. Ecology 85, 1062–1071. Cargill, S.M., Chapin, F.F., 1987. Application of successional theory to tundra restoration: a review. Arct. Alp. Res. 19, 366–372. Carlsson, B., Callaghan, T.V., 1991. Positive plant interactions in tundra vegetation and the importance of shelter. J. Ecol. 79, 973–983. Casper, B.B., Castelli, J.P., 2007. Evaluating plant–soil feedback together with competition in a serpentine grassland. Ecol. Lett. 10, 394–400. Chapin, F.S., Chapin, M.S., 1980. Revegetation of an arctic disturbed site by native tundra species. J. Appl. Ecol. 17, 449–456. Choler, P., Michalet, R., Callaway, R.M., 2001. Facilitation and competition on gradients in alpine plant communities. Ecology 82, 3295–3308. Daft, J., Nicholson, T.H., 1974. Arbuscular mycorrhizas in plants colonizing coal wastes in Scotland. New Phytol. 73, 1129–1138. Dormann, C.F., van der Wal, R., Woodin, S.J., 2004. Neighbour identity modifies effects of elevated temperature on plant performance in the high Arctic. Global Change Biol. 10, 1587–1598. Eskelinen, A., 2008. Herbivore and neighbour effects on tundra plants depend on species identity, nutrient availability and local environmental conditions. J. Ecol. 96, 155–165. Frostegård, Å., Bååth, E., Tunlid, A., 1993. Shifts in the structure of soil microbial communities in limed forest as revealed by phospholipid fatty acid analysis. Soil Biol. Biochem. 25, 723–730. Garbeva, P., van Elsas, J.D., van Veen, J.A., 2008. Rhizosphere microbial community and its response to plant species and soil history. Plant Soil 302, 19–32. Gianinazzi-Pearson, V.G., Morandi, D., Dexheimer, J., Gianinazzi, S., 1981. Ultrastructural and ultracytochemical features of a Glomus tenuis mycorrhiza. New Phytol. 88, 633–639. Gough, L., 2006. Neighbour effects on germination, survival, and growth in two arctic tundra plant communities. Ecography 29, 44–56. Griffiths, B.S., Ritz, K., Ebblewhite, N., Dobson, G., 1999. Soil microbial community structure: effects of substrate loading rates. Soil Biol. Biochem. 31, 145–153. Hedlund, K., 2002. Soil microbial community structure in relation to vegetation management on former agricultural land. Soil Biol. Biochem. 34, 1299–1307. Ives, I.D., 1974. The impact of motor vehicles on the tundra environments. In: Ives, I.D., Barry, R.G. (Eds.), Arctic and Alpine Environments. Methuen & Co Ltd, London, pp. 907–910. Jonasson, S., Michelsen, A., Schmidt, I., 1999. Coupling of nutrient cycling and carbon dynamics in the Arctic, integration of soil microbial and plant processes. Appl. Soil Ecol. 11, 135–146. Jorgenson, M.T., Joyce, M.R., 1994. Six strategies for rehabilitating land disturbed by oil development in arctic Alaska. Arctic 47, 374–390. Järvinen, A., 1987. Basic climatological data on the Kilpisjärvi area NW Finnish Lapland. Kilpisjärvi Notes 10, 1–16. Kashulina, G., Reimann, C., Finne, T.E., Halleraker, J.H., Äyräs, M., Chekushin, V.A., 1997. The state of the ecosystems in central Barents region: scale, factors and mechanism of disturbance. Sci. Total Environ. 206, 203–225. Kattsov, V.M., Källén, E., Cattle, H., Christensen, J., Drange, H., Hanssen-Bauer, I., Jóhannsen, T., Karol, I., Räisänen, J., Svensson, G., Vavulin, S., Chen, D., Polyakov, I., Rinke, A., 2005. Future climate change: modeling and scenarios for the Arctic. In: Arctic Climate Impact Assessment. Cambridge University Press, New York, pp. 99–150. Kelly, R.H., Burke, I.C., Lauenroth, W.K., 1996. Soil organic matter and nutrient availability responses to reduced plant inputs in shortgrass steppe. Ecology 77, 2516–2527. Klanderud, K., Tøtland, Ö., 2004. Habitat dependent nurse effects of the dwarf shrub Dryas octopetala on arctic and alpine plant community structure. Ecoscience 11, 410–420. Kramer, C., Gleixner, G., 2008. Soil organic matter in depth profiles: distinct carbon preferences of microbial groups during carbon transformation. Soil Biol. Biochem. 40, 425–433.

Kramer, C., Gleixner, G., 2006. Variable use of plant- and soil-derived carbon by microorganisms in agricultural soils. Soil Biol. Biochem. 38, 3267–3278. Kytöviita, M.-M., 2005. Asymmetric symbiont adaptation to arctic conditions could explain why high arctic plants are non-mycorrhizal. FEMS Microbiol. Ecol. 53, 27–32. Loeffler, J., 2002. High mountain ecosystems and landscape degradation in northern Norway. Mt. Res. Dev. 20, 356–363. Mack, M.C., Schuur, E.A.G., Bret-Harte, M.S., Shaver, G.R., Chapin, F.S., 2004. Ecosystem carbon storage in arctic tundra reduced by long-term nutrient fertilization. Nature 431, 440–443. Maffia, B., Nadkarni, N.M., Janos, P., 1993. Vesicular-arbuscular mycorrhiza of epiphytic and terrestrial Piperaceae under field and greenhouse conditions. Mycorrhiza 4, 5–9. McGonigle, T.P., Miller, M.H., Evans, D.G., Fairchild, G.L., Swan, J.A., 1990. A new method which gives an objective measure of colonization of roots by vesiculararbuscular mycorrhizal fungi. New Phytol. 115, 495–501. McKane, R.B., Rastetter, E.B., Shaver, G.R., Nadelhoffer, K.J., Giblin, A.E., Laundre, J.A., Chapin, F.S., 1997. Climatic effects on tundra carbon storage inferred from experimental data and a model. Ecology 78, 1170–1187. Miller, R.M., Jastrow, J.D., 1992. The application of VA mycorrhizae to ecosystem restoration and reclamation. In: Allen, M.F. (Ed.), Mycorrhizal Functioning: An Integrative Plant-Fungal Process. Chapman and Hall, New York, pp. 438–467. Moorman, T., Reeves, F.B., 1979. The role of endomycorrhizae in revegetation practices in the semi-arid west. II. A bioassay to determine the effect of land disturbance on endomycorrhizal populations. Am. J. Bot. 66, 14–18. Nara, K., 2005. Ectomycorrhizal networks and seedling establishment during early primary succession. New Phytol. 169, 169–178. Newsham, K.K. A meta-analysis of plant responses to dark septate root endophytes. New Phytol., in press, doi:10.1111/j.1469-8137.2010.03611.x. Olsson, P.A., Johansen, A., 2000. Lipid and fatty acid composition and spores of mycorrhizal fungi at different growth stages. Mycol. Res. 104, 249–434. Olofsson, L., Moen, J., Oksanen, L., 1999. On the balance between positive and negative plant interactions in harsh environments. Oikos 86, 539–543. Olofsson, J., Hulme, P.E., Oksanen, L., Suominen, O., 2005. Effects of mammalian herbivores on revegetation of disturbed areas in the forest-tundra ecotone in northern Fennoscandia. Landscape Ecol. 20, 351–359. Padilla, F.M., Pugnaire, F.I., 2006. The role of nurse plants in the restoration of degraded environments. Front. Ecol. Environ. 4, 196–202. Peacock, A.D., Macnaughton, S.J., Cantu, J.M., Dale, V.H., White, D.C., 2001. Soil microbial biomass and community composition along an anthropogenic disturbance gradient within a long-leaf pine habitat. Ecol. Indic. 1, 113–121. Phillips, J.M., Hayman, D.S., 1970. Improved procedures for clearing roots and staining parasitic and vesicular-arbuscular mycorrhizal fungi for rapid assessment of infection. Trans. Br. Mycol. Soc. 55, 158–161. Pennanen, T., Liski, J., Bååth, E., Kitunen, V., Uotila, J., Westman, C.J., Fritze, H., 1999. Structure of the microbial communities in coniferous forest soils in relation to site fertility and stand development stage. Microb. Ecol. 38, 168–179. Pietikäinen, A., Kytöviita, M.-M., Husband, R., Young, P., 2007. Diversity and persistence of arbuscular mycorrhizas in low arctic meadow habitat. New Phytol. 176, 691–698. Pietikäinen, A., Kytöviita, M.-M., 2007. Defoliation changes mycorrhizal benefit and competitive interactions between seedlings and adult plant. J. Ecol. 95, 639–647. Pietikäinen, A., Kytöviita, M.-M., Vuoti, U., 2005. Mycorrhizal symbiosis and seedling establishment in a subarctic meadow: effects of fertilization and clipping. J. Veg. Sci. 16, 175–182. Read, D.J., Haselwandter, K., 1981. Observations on the mycorrhizal status of some alpine plant communities. New Phytol. 88, 341–352. Reeves, F.B., Wagner, D., Moorman, T., Kiel, J., 1979. The role of endomycorrhizae in revegetation practices in the semi-arid west I. A comparison of incidence of mycorrhizae in severely disturbed vs. natural habitats. Am. J. Bot. 66, 6–13. Rillig, M.C., Mummey, D.L., 2006. Mycorrhizas and soil structure. New Phytol. 171, 41–53. Ruotsalainen, A.-L., Väre, H., Vestberg, M., 2002. Seasonality of root fungal colonization in low-alpine herbs. Mycorrhiza 12, 29–36. Schadt, C.W., Martin, A.P., Lipson, D.A., Schmidt, S.K., 2003. Seasonal dynamics of previously unknown fungal lineages in tundra soils. Science 301, 1359–1361. Schimel, J.P., Weintraub, M.N., 2003. The implications on exoenzyme activity on microbial carbon and nitrogen limitation in soil: a theoretical model. Soil Biol. Biochem. 35, 549–563. Schmidt, N., Bölter, M., 2002. Fungal and bacterial biomass in tundra soils along an arctic transect from Taimyr Peninsula, central Siberia. Polar Biol. 25, 871–877. Sjögersten, S., Turner, B., Mahieu, N., Condron, L.M., Wookey, P.A., 2003. Soil organic matter biochemistry and potential susceptibility to climatic change across the forest-tundra ecotone in the Fennoscandian mountains. Global Change Biol. 9, 759–772. Smith, S.E., Read, D.J., 1997. Mycorrhizal Symbiosis. Academic Press, London, p. 605. Stark, S., Kytöviita, M.-M., 2006. Simulated grazer effects on microbial respiration in a subarctic meadow: implications for nutrient competition between plants and soil microorganisms. Appl. Soil Ecol. 31, 20–31. Suding, K.N., Gross, K.L., Houseman, R.G., 2004. Alternative states and positive feedbacks in restoration ecology. Trends Ecol. Evol. 19, 46–53. Tamminen, P., Starr, M.R., 1990. A survey of forest soil properties related to soil acidification in southern Finland. In: Kauppi, P., Anttila, P., Kenttämies, K. (Eds.), Acidification in Finland. Springer-Verlag, Berlin, pp. 234–251.

M.-M. Kytöviita et al. / Applied Soil Ecology 48 (2011) 142–151 Treu, R., Laursen, G.A., Stephenson, S.L., Landolt, J.C., Densmore, R., 1996. Mycorrhizae from Denali National Park and Reserve, Alaska. Mycorrhiza 6, 21–29. Vandenkoornhuyse, P., Ridgeway, K.P., Watson, I.J., Fitter, A.H., Young, J.P.W., 2003. Co-existing grass species have distinctive arbuscular mycorrhizal communities. Mol. Ecol. 12, 3085–3095. Vrålstad, T., Myhre, E., Schumacher, T., 2002. Molecular diversity and phylogenetic affinities of symbiotic root associated ascomycetes of the Helotiales in burnt and metal polluted sites. New Phytol. 155, 131–148.

151

Walker, L., Chapin, F.S., 1986. Physiological controls over seedling growth in primary succession on an Alaskan floodplain. Ecology 67, 1508–1523. White, J.A., Tallaksen, J., Charvat, I., 2008. The effects of arbuscular mycorrhizal fungal inoculation at a roadside prairie restoration site. Mycologia 100, 6–11. Zabavski, J., Zuravska, M., 1977. Quantitative studies of fungi and Actinomycetes in the primary soils of West-Spitzbergen. In: Szegi, J. (Ed.), Soil Biology and Conservation of the Biosphere. Akademiai Kiado, Budapest, pp. 3343–3353. Zak, D., Kling, G.W., 2006. Microbial community composition and function across and arctic tundra landscape. Ecology 87, 1659–1670.