Reviewers: Weigel R3, Ribbons R4
Measurement unit: percentage of colonisation; Measurement scale: plot or plant; Equipment costs: €; Running costs: €; Installation effort: medium to high; Maintenance effort: -; Knowledge need: medium to high; Measurement mode: manual
Mycorrhizae are plant–fungi symbiotic relationships in which plants benefit from fungi-derived nutrient supplies and protection from environmental stresses while fungi are provided with plant-synthesised carbon (Smith & Read, 2008). Arbuscular mycorrhizal fungi (AMF), with their namesake nutrient-exchange structures called arbuscules, are the most common type of mycorrhizal fungi, associating with more than 80% of vascular plant species including both herbaceous and woody plants (Bueno et al., 2017). The second most abundant mycorrhizal association is Ectomycorrhiza (EcM) which occurs in 2% of seed plants (Maherali et al., 2016), most of which are highly prevalent temperate tree species. Mycorrhizal fungi can strongly affect plant nutrient uptake, biomass, and photosynthesis as well as the carbon allocation (Bago et al., 2000). Thus, they can play an important role in global carbon and nutrient cycling processes (Hodge et al., 2001; Veresoglou et al., 2012; Soudzilovskaia et al., 2015). In many ecosystems, mycorrhizae, together with fine roots, provide the largest input of carbon into soils (Kramer et al., 2010; Verbruggen et al., 2016). Mycorrhizal symbionts hence determine the flow of vast quantities of carbon on global scales, where the impact of AMF alone may be as large as 4.5 billion tonnes of carbon annually (Bago et al., 2000). The intensity of plant root colonisation by AMF is expressed as a percentage of root length colonised by the fungi (Soudzilovskaia et al., 2015). This is the single best available measure that quantifies the “strength” of the plant–fungi relationship in situ and it can indicate the reliance of plants on mycorrhizae in a particular ecosystem (Soudzilovskaia et al., 2015). This is likely positively correlated with nutrient provision by, and C supply to, AMF, but it is not a direct measure, so it should be interpreted with care. AMF can also be isolated directly from the soil which can provide information about the “extraradical” part of AMF consisting of hyphae foraging for nutrients. EcM fungi can be quantified in an analogous process. Quantifying root colonisation by mycorrhizal fungi can be used as a measure of the strength of plant–fungal relationships and an indicator of nutrient and carbon flow between plants and fungi in both manipulation and observational experiments investigating the effect of environmental changes, such as warming (Wilson et al., 2016), elevated CO2 (Staddon et al., 1999), nitrogen enrichment (Jumpponen et al., 2005), and land-use change (Xiang et al., 2014) on ecosystem functioning.
188.8.131.52 What and how to measure?
Depending of the study question, either the roots of particular plant species are excavated to assess the mycorrhizal colonisation of these plant species, or, if the colonisation of all roots within a soil sample is of interest, a soil core is taken (ca. 10 g of soil) from the experimental plot usually at 0-5 or 5-10 cm depth. The samples can be stored in sterile plastic ziplock bags and, if not processed immediately, they should be kept in the freezer (at -20 °C). Typically, between 3 and 5 samples are pooled to represent an experimental unit. Root colonisation varies over time but is highest around the time of flowering, which represents an optimal time for sampling (e.g. Gosling et al., 2013), although earlier development can also be of interest as it is likely relevant for plant development. Subsequently, live-roots are picked out from soil following, for example, the root extraction-flotation method (Cook et al., 1988). Briefly, the soil with roots is washed repeatedly under running water over a sieve (with a mesh size < 1 mm) until the soil particles are completely removed (see also Pérez-Harguindeguy et al., 2013).
After extraction, roots are kept fresh at 4 °C or in 70% ethanol followed by (optional) determination of length of roots according to the method described by Tennant (1975). Alternatively, the roots can be scanned and analysed using dedicated software (for some examples see Cai et al., 2015). In order to measure root length colonisation of AMF, the roots are first cleared in 5–10% KOH in a 90 °C waterbath. Thicker and more coloured roots require more time than small and less coloured roots. Highly pigmented and lignified roots can additionally be bleached or subjected to H2O2 to remove any remaining phenolic compounds and pigmentation. In order to make AMF structures visible under the microscope, the roots need to be stained. This is commonly done using Trypan Blue as in Phillips & Hayman (1970) or ink and vinegar as in Vierheilig et al. (1998). Approximately 20 pieces of stained roots (ca. 2 cm long each) are randomly selected and aligned on a microscope slide. The percent of AMF colonisation can then be estimated using the magnified intersections method, described by McGonigle et al. (1990).
In order to avoid biases, it is very important that the selection of root pieces and the location of the cross-hair (see McGonigle et al., 1990 for description) are random. It is also necessary to check the success of root staining, for instance by examining if the stele (vascular tissue) is stained, which is a good indication that the staining is successful. If the whole root is translucent, this indicates that the staining was not efficient and these pieces should not be counted as non-infected. The roots that only have a stele, an incomplete cortex, or those which are heavily colonised by dark-septate fungi (as a signal of potential root mortality) should be excluded from the analysis. Furthermore, given that the method requires recognising AMF hyphae, arbuscules, and vesicles, some training is required, for instance starting with pictures/descriptions at https://mycorrhizas.info/. One distinguishing feature of AMF contrasted against most other fungi is that their hyphae are aseptate, meaning that they are not regularly subdivided by septa (crosswalls within the filaments). The same website can be used to get acquainted with the appearance of EcM fungi. Because these fungi form a sheath around tree roots, fresh roots can be examined submerged in a petri dish under a dissecting microscope and EcM root tips can be expressed as the percentage colonisation of total root tips.
The hyphae are extracted from 4 g of soil using the aqueous extraction method as described in Jakobsen et al. (1992). This method involves the use of strong mechanical force (i.e. blending at high speeds) which could cause hyphae to break (thereby possibly hindering the identification of different hyphal types). To avoid this, the method above can be modified as in Rillig et al. (1999), using sodium hexametaphosphate to break up soil aggregates and bring all hyphae into suspension. The soil suspension is carefully decanted over a 38 µm sieve and gently sprayed to remove clay particles. Then, material left on the sieve is flushed in a known volume of water (e.g. 200 ml) and a subsample is brought on a nitrocellulose filter on a vacuum manifold to isolate the hyphae. The filter is stained with an ink and vinegar solution (or 0.02% Trypan Blue in lactoglycerol) and inspected using a microscope at x200 magnification. Hyphal length can be determined using the visual gridline intersection (VDI) method, which is based on the examination of the frequency of hyphal intersections with gridlines on a microscope eyepiece. The hyphal length is obtained by incorporating the number of hyphae crossing with gridlines into the formula by Newman (1966) (as in, for example, Camenzind & Rillig, 2013). Hyphal extraction efficiency is determined by re-extracting hyphae based on the method by Miller et al. (1995) and this can be used to adjust the values of hyphal lengths accordingly.
Where to start
Camenzind & Rillig (2013), McGonigle et al. (1990), Miller et al. (1995), Rillig et al. (1999), Vierheilig et al. (1998)
184.108.40.206 Special cases, emerging issues, and challenges
Measurement of hyphal length in soil is simple and cost efficient, but rather time-consuming since it requires an experienced eye in distinguishing AMF hyphae from non-AMF hyphae. Alternative methods, based on photomicrography and image processing offer semi-automated analysis of hyphal length where observer biases are minimised (Shen et al., 2016) but do not distinguish AM and non-AM fungi.
To estimate AMF biomass in the roots and soil, the neutral lipid fatty acid (NLFA) 16: 1ω5 (Olsson, 1999) can be used as an AMF specific biomarker (see protocol 2.2.1. Soil microbial biomass – C, N, and P for a small description of PLFA/NLFA analysis). There is no specific biomarker for EcM fungi, although when EcM tree roots are colonised, the majority of fungal PLFA markers or ergosterol will generally originate from EcM fungi.
Theory, significance, and large datasets
Bago et al. (2000), Bueno et al. (2017), Maherali et al. (2016), Smith & Read (2008), Soudzilovskaia et al. (2015)
More on methods and existing protocols
Cai et al. (2015), Cook et al. (1988), Pérez-Harguindeguy et al. (2013), Phillips & Hayman (1970), Shen et al. (2016), Tennant (1975)
Bago, B. Pfeffer, P. E., & Shachar-Hill, Y. (2000). Carbon metabolism and transport in arbuscular mycorrhizas. Plant Physiology, 124(3), 949-958.
Bueno, C. G., Moora, M., Gerz, M., Davison, J., Öpik, M., Pärtel, M., … Zobel, M. (2017). Plant mycorrhizal status, but not type, shifts with latitude and elevation in Europe. Global Ecology and Biogeography, 26(6), 690-699.
Cai, J. Zeng, Z. Connor, J. N. Huang, C. Y. Melino, V. Kumar, P., & Miklavcic, S. J. (2015). RootGraph: a graphic optimization tool for automated image analysis of plant roots. Journal of Experimental Botany, 66(21), 6551-6562.
Camenzind, T., & Rillig, M. C. (2013). Extraradical arbuscular mycorrhizal fungal hyphae in an organic tropical montane forest soil. Soil Biology and Biochemistry, 64, 96-102.
Cook, B. D., Jastrow, J. D., & Miller, R. M. (1988). Root and mycorrhizal endophyte development in a chronosequence of restored tallgrass prairie. New Phytologist, 110(3), 355-362.
Gosling, P., Mead, A., Proctor, M., Hammond, J. P., & Bending, G. D. (2013). Contrasting arbuscular mycorrhizal communities colonizing different host plants show a similar response to a soil phosphorus concentration gradient. New Phytologist, 198(2), 546-556.
Hodge, A., Campbell, C. D., & Fitter, A. H. (2001). An arbuscular mycorrhizal fungus accelerates decomposition and acquires nitrogen directly from organic material. Nature, 413(6853), 297-299.
Jakobsen, I., Abbott, L. K., & Robson, A. D. (1992). External hyphae of vesicular-arbuscular mycorrhizal fungi associated with Trifolium subterraneum L. 1. Spread of hyphae and phosphorus inflow into roots. New Phytologist, 120(3), 371-380.
Jumpponen, A., Trowbridge, J., Mandyam, K., & Johnson, L. (2005). Nitrogen enrichment causes minimal changes in arbuscular mycorrhizal colonization but shifts community composition—evidence from rDNA data. Biology and Fertility of Soils, 41(4), 217-224.
Kramer, C., Trumbore, S. E., Fröberg, M., Dozal, L. M. C., Zhang, D., Xu, X., … Hanson, P. J. (2010). Recent (< 4 year old) leaf litter is not a major source of microbial carbon in a temperate forest mineral soil. Soil Biology and Biochemistry, 42(7), 1028-1037.
Maherali, H., Oberle, B., Stevens, P. F., Cornwell, W. K., & McGlinn, D. J. (2016). Mutualism persistence and abandonment during the evolution of the mycorrhizal symbiosis. The American Naturalist, 188(5), E113-E125.
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 vesicular-arbuscular mycorrhizal fungi. New Phytologist, 115(3), 495-501.
Miller, R. M., Jastrow, J. D., & Reinhardt, D. R. (1995). External hyphal production of vesicular-arbuscular mycorrhizal fungi in pasture and tallgrass prairie communities. Oecologia, 103(1), 17-23.
Newman, E. I. (1966). A method of estimating the total length of root in a sample. Journal of Applied Ecology, 3(1), 139.
Olsson, P. A. (1999). Signature fatty acids provide tools for determination of the distribution and interactions of mycorrhizal fungi in soil. FEMS Microbiology Ecology, 29(4), 303-310.
Pérez-Harguindeguy, N., Diaz, S., Garnier, E., Lavorel, S., Poorter, H., Jaureguiberry, P., … Cornelissen, J. H. C. (2013). New handbook for standardized measurement of plant functional traits worldwide. Australian Journal of Botany, 61(34), 167-234.
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. Transactions of the British Mycological Society, 55(1), 158-161.
Rillig, M. C., Field, C. B., & Allen, M. F. (1999). Soil biota responses to long-term atmospheric CO2 enrichment in two California annual grasslands. Oecologia, 119(4), 572-577.
Shen, Q., Kirschbaum, M. U. F., Hedley, M. J., & Camps Arbestain, M. (2016). Testing an alternative method for estimating the length of fungal hyphae using photomicrography and image processing. PloS One, 11(6), e0157017.
Smith, S. E., & Read, D. J. (2008). Mycorrhizal Symbiosis. Academic Press.
Soudzilovskaia, N. A., Douma, J. C., Akhmetzhanova, A. A., van Bodegom, P. M., Cornwell, W. K., Moens, E. J., … Cornelissen, J. H. C. (2015). Global patterns of plant root colonization intensity by mycorrhizal fungi explained by climate and soil chemistry. Global Ecology and Biogeography, 24(3), 371-382.
Staddon, P. L., Fitter, A. H., & Graves, J. D. (1999). Effect of elevated atmospheric CO2 on mycorrhizal colonization, external mycorrhizal hyphal production and phosphorus inflow in Plantago lanceolata and Trifolium repens in association with the arbuscular mycorrhizal fungus Glomus mosseae. Global Change Biology, 5(3), 347-358.
Tennant, D. (1975). A test of a modified line intersect method of estimating root length. Journal of Ecology, 63(3), 995-1001.
Verbruggen, E., Jansa, J., Hammer, E. C., & Rillig, M. C. (2016). Do arbuscular mycorrhizal fungi stabilize litter-derived carbon in soil? Journal of Ecology, 104(1), 261-269.
Veresoglou, S. D., Chen, B., & Rillig, M. C. (2012). Arbuscular mycorrhiza and soil nitrogen cycling. Soil Biology and Biochemistry, 46, 53-62.
Vierheilig, H., Coughlan, A. P., Wyss, U., & Piche, Y. (1998). Ink and vinegar, a simple staining technique for arbuscular-mycorrhizal fungi. Applied and Environmental Microbiology, 64(12), 5004-5007.
Authors: Verbruggen E1, Radujković D1
Reviewers: Weigel R3, Ribbons R4
1 Centre of Excellence PLECO (Plants and Ecosystems), Biology Department, University of Antwerp, Wilrijk, Belgium
2 Experimental Plant Ecology, Institute of Botany and Landscape Ecology, University of Greifswald, Greifswald, Germany
3 Plant Ecology, Albrecht-von-Haller Institute for Plant Sciences, University of Goettingen, Goettingen, Germany
4 Biology and Geology Departments, Lawrence University, Appleton, USA