Ecological Archives E093-061-A1

Emily Grman. 2012. Plant species differ in their ability to reduce allocation to non-beneficial arbuscular mycorrhizal fungi. Ecology 93:711–718.

Appendix A. Detailed methods describing greenhouse conditions, sampling procedure, statistical analyses, and references.


To test the hypothesis that a species' mycorrhizal responsiveness would determine its ability to reduce allocation to non-beneficial arbuscular mycorrhizal fungi and thus experience different levels of benefit or parasitism, I conducted a greenhouse experiment.  In a factorial design of five phosphorus levels, two light levels, and two AMF treatments (mycorrhizal and nonmycorrhizal), I grew four species of grass: two native C4 prairie bunchgrasses, big bluestem (Andropogon gerardii Vitman) and little bluestem (Schizachyrium scoparium (Michx.) Nash), and two introduced C3 grasses, smooth brome (Bromus inermis Leyss.) and quackgrass (Elymus repens (L.) Gould.).

Soil and fungal inoculum.  I grew the plants and AMF in 0.7 L pots containing a mixture of 90% sand and 10% field soil collected from an old-field in southwest Michigan and sieved through a 2 mm sieve.  I autoclaved the sand/soil mix for three hours in the nonmycorrhizal treatment and did not autoclave the mix in the mycorrhizal treatment.  The non-autoclaved sand and soil in the mycorrhizal treatment served as a source of AMF.  All pots also received 150 mL additional inoculum of field soil, either autoclaved (nonmycorrhizal treatment) or not (mycorrhizal treatment), and an additional 5 cm of autoclaved sand/soil mix on top of the inoculum to reduce cross-contamination among pots.  I watered the pots excessively to leach nutrients released during autoclaving.  To control for the abundance of other soil microbes, I added to each pot 40 mL of a microbial wash prepared by blending field soil with water in a 1:5 ratio and filtering through a 35 µm sieve.  This sieve size excluded mycorrhizae from the microbial wash, but allowed bacteria and other microbes to pass through (Corkidi et al. 2002, Johnson et al. 2008). 

Light and phosphorus treatments.  In December 2008, I randomly arranged these pots into six replicate blocks on benches in a heated greenhouse with supplemental lighting at the W. K. Kellogg Biological Station in southwest Michigan.  I germinated seeds of each species in petri plates and transplanted newly emerged seedlings into the pots to ensure that one individual grew in each pot.  To manipulate light availability, I placed half the pots in each block under a shade structure that blocked 30% of incoming light (low light treatment).  The other half of the pots received ambient light (high light treatment).  To manipulate phosphorus availability, I randomly assigned to each pot one of five fertilizer solutions with 0, 0.15, 0.31, 3.1, or 31.0 g/L NaH2PO4.  To balance the pH, I added a different amount of 1M NaOH to each fertilizer solution: 0.05, 0.06, 0.25, 1.5, or 12.5 mL/L, from lowest to highest phosphorus treatment.  To ensure that phosphorus was the only limiting nutrient, all of the fertilizer solutions included a mixture of 24 g/L KNO3, 20 g/L MgSO4 7H2O, 0.5 g/L KCl, 0.11 g/L H3BO3, 0.05 g/L ZnSO4 7H2O, 0.03 g/L MnSO4 H2O, 3.3 mg/L CuSO4 5H2O, and 0.6 mg/L MoO3.  The pots were watered to capacity 5–6 times a week with tap water and fertilizer was added once weekly, immediately after watering the pots.  To increase the chance of successful seedling establishment, I began fertilizer applications three weeks after transplanting all seedlings into pots.  I added 7.5 mL of the appropriate solution to the pots for the first four weeks of the fertilizer applications and 15 mL during weeks 5–8.  The highest level of phosphorus addition was intended to mimic the high phosphorus availability that can result from application of manure to agricultural soils (40–120 µg P/g soil; Andraski and Bundy 2003, Butler and Coale 2005). 

Harvesting plants and fungi. I harvested the experiment nine weeks after initiating the fertilizer treatments.  To determine plant aboveground biomass, I clipped seedlings at the soil surface and placed the material in coin envelopes or small paper bags.  To determine belowground plant biomass, I placed the contents of each pot in a 2 mm sieve and gently agitated the soil and roots to remove as much soil as possible from the roots.  I saved and air-dried this soil for later analysis of phosphorus availability and AMF abundance.  I then submerged the roots in water and gently agitated them to remove the remaining soil.  I placed the wet, cleaned roots in coin envelopes or small paper bags.  I dried both root and shoot biomass in an oven at 65°C to constant mass (at least 48 hours). 

Assessing AMF abundance in plant roots.  To determine root colonization by arbuscular mycorrhizal fungi, I cleared haphazardly-chosen subsamples of dried roots in 2.5% KOH for 90–120 minutes in an oven at 90°C, soaked roots in 1% HCl overnight, and stained for 90 minutes at 90°C in trypan blue stain, a solution of 500 mL/L glycerol, 450 mL/L water, 5 mL/L 1% HCl, and 0.5 g/L trypan blue (modified from Koske and Gemma 1989).  I then mounted roots on microscope slides with water and assessed percent root colonization using the gridline intercept method (Giovannetti and Mosse 1980) at 100x magnification.  I quantified root colonization in all pots in the mycorrhizal treatment and one third of the pots (two of six blocks) in the nonmycorrhizal treatment to test for contamination.  Because the mean of root colonization in the nonmycorrhizal treatment was less than 1%, I decided it was not necessary to quantify colonization in the nonmycorrhizal pots in the remaining four blocks. 

Assessing AMF abundance in the soil.  To determine the abundance of AMF hyphae in the soil (extra-radical hyphae), I thoroughly mixed all the air-dried soil saved from each pot.  I extracted the hyphae from a 5 g subsample of each sample by stirring it into 90 mL of 20 g/L sodium hexametaphosphate on a stir plate, sonicating, filtering a subsample through 20 µm mesh, resuspending the hyphae trapped on the mesh in trypan blue stain, staining overnight, filtering through 1.2 µm pore size nitrocellulose filters, mounting the filters with immersion oil on microscope slides, and quantifying AMF hyphal abundance using the gridline intercept method at 400x magnification (modified from Miller et al. 1995).  To increase accuracy of quantifying extra-radical hyphae, I counted each slide twice and used the averaged value. 

Soil phosphorus analysis.  To determine water-extractable phosphorus in the soil at the end of the experiment, I extracted 10 g of air-dried soil in 50 mL water by shaking for 2 minutes, allowing to settle overnight, and filtering through glass fiber filters (modified from Olsen and Sommers 1982).  I froze the extract until analysis.  To determine the phosphorus content of the extract, I developed color using malachite green and determined intensity of color development on a microplate reader (SpectraMax M5, Molecular Devices, Sunnyvale, CA, USA; modified from D'Angelo et al. 2001).

Mycorrhizal responsiveness (MR).  To determine plant response to AMF, I summed root and shoot biomass, paired plants of the same species grown in the same resource environment (phosphorus and light) in each block (and differing only in AMF treatment), and calculated an index of mycorrhizal responsiveness (van der Heijden 2002).  When plant biomass with AMF (bAMF) was greater than biomass without (bN), MR = 100(1 - bN / bAMF); when bAMF < bN, MR = 100(bAMF / bN - 1).  MR ranges from -100 to 100; MR > 0 indicates that AMF acted as mutualists to increase plant biomass and MR < 0 indicates that AMF acted as parasites to reduce plant biomass.  Other indices of mycorrhizal responsiveness (Graham and Eissenstat 1994, Johnson 1998, Kaeppler et al. 2000) gave qualitatively similar results. 

Statistical analysis.  I conducted all statistical analyses in R (2.10.1, The R Foundation for Statistical Computing), package nlme.  I analyzed the data as a split-plot ANCOVA.  The plant species (4 levels), AMF (2 levels), and phosphorus treatments (5 levels) were completely randomized within the whole-plot level light treatment (2 levels); all treatment combinations were present in each of six replicate blocks (480 pots in total).  I used water-extractable soil P as a continuous predictor instead of the discrete phosphorus treatment variable because the phosphorus treatment explained 98% of the variation in water-extractable soil P.  To better examine responses across the range of tested phosphorus availabilities, which spanned three orders of magnitude, I log-transformed this continuous independent variable.  I used ANOVA and ANCOVA models to test for the effects of resources on log-transformed total plant biomass, plant mycorrhizal responsiveness, square-root transformed percent root colonization by AMF, and log-transformed extra-radical hyphal abundance.  When there were significant interactions between species and resource availability, I conducted separate ANOVA or ANCOVA on each species to determine how individual species responded to phosphorus or light. 

One E. repens individual and 35 S. scoparium individuals died before the end of the experiment.  To maintain a balanced design, I included the biomass collected from those pots in the analyses below.  Removing them would not qualitatively change the results.  However, root samples of five S. scoparium individuals were too small to assess percent root colonization so these data were omitted from analyses of root colonization.  I used marginal sums of squares in all analyses including this unbalanced root colonization dataset.

Model simplification.  I performed model simplification as suggested in Crawley (2007) to better understand treatment effects and species differences.  I removed non-significant interactions.  Because I was interested in comparing the responses of C3 and C4 grasses, I combined species into the appropriate functional group (C3 or C4) and tested whether this simplification (removal of extra terms) affected model fit.  A result of no difference (P > 0.05) between the complex model (containing terms for individual species) and the simple model (containing only terms for functional group) indicated that species within functional groups were not significantly different, either in mean response or in interactions with other treatment variables.

Literature Cited

Andraski, T. W. and L. G. Bundy. 2003. Relationships between phosphorus levels in soil and runoff from corn production systems. Journal of Environmental Quality 32:310–316.

Butler, J. S. and F. J. Coale. 2005. Phosphorus leaching in manure-amended Atlantic Coastal Plain soils. Journal of Environmental Quality 34:370–381.

Corkidi, L., D. L. Rowland, N. C. Johnson, and E. B. Allen. 2002. Nitrogen fertilization alters the functioning of arbuscular mycorrhizas at two semiarid grasslands. Plant and Soil 240:299–310.

D'Angelo, E., J. Crutchfield, and M. Vandiviere. 2001. Rapid, sensitive, microscale determination of phosphate in water and soil. Journal of Environmental Quality 30:2206–2209.

Giovannetti, M. and B. Mosse. 1980. An evaluation of techniques for measuring vesicular arbuscular mycorrhizal infection in roots. New Phytologist 84:489–500.

Graham, J. H. and D. M. Eissenstat. 1994. Host genotype and the formation and function of VA mycorrhizae. Plant and Soil 159:179–185.

Johnson, N. C. 1998. Responses of Salsola kali and Panicum virgatum to mycorrhizal fungi, phosphorus and soil organic matter: implications for reclamation. Journal of Applied Ecology 35:86–94.

Johnson, N. C., D. L. Rowland, L. Corkidi, and E. B. Allen. 2008. Plant winners and losers during grassland N-eutrophication differ in biomass allocation and mycorrhizas. Ecology 89:2868–2878.

Kaeppler, S. M., J. L. Parke, S. M. Mueller, L. Senior, C. Stuber, and W. F. Tracy. 2000. Variation among maize inbred lines and detection of quantitative trait loci for growth at low phosphorus and responsiveness to arbuscular mycorrhizal fungi. Crop Science 40:358–364.

Koske, R. E. and J. N. Gemma. 1989. A modified procedure for staining roots to detect VA mycorrhizas. Mycological Research 92:486–488.

Miller, R. M., D. R. Reinhardt, and J. D. Jastrow. 1995. External hyphal production of vesicular-arbuscular mycorrhizal fungi in pasture and tallgrass prairie communities. Oecologia 103:17–23.

Olsen, S. R. and L. E. Sommers. 1982. Phosphorus. Pages 403-430 in A. L. Page, editor. Methods of Soil Analysis, Part 2.  Chemical and Microbiological Properties. American Society of Agronomy, Soil Science Society of America, Madison, WI.

van der Heijden, M. G. A. 2002. Arbuscular mycorrhizal fungi as a determinant of plant diversity: in search of underlying mechanisms and general principles. Pages 243–265 in M. G. A. van der Heijden and I. Sanders, editors. Mycorrhizal Ecology. Springer-Verlag, Berlin.

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