Supplementary Material 2 - Nutrient uptake strategies

Almost all land plants form symbiotic associations with mycorrhizal fungi and (or) nitrogen fixing bacteria and relay on these plant symbionts for nutrient uptake (Franche et al. 2009; Lambers et al. 2009). Mycorrhyzial fungi and nitrogen fixing bacteria are responsible for ca. 5-20% (grassland and savannah) to 80% (temperate and boreal forest) of all nitrogen, and up to 75% of phosphorus, that is acquired by plants annually (e.g. van der Heijden et al. 2008). Moreover, some plants parasite on other plants or on fungi to obtain nutrients and carbon. Using the protocols describe below, we give a brief guide to identify major nutrient uptake strategies. See Table SM 2.1 for updated references on these strategies.

Assign preferably one category to each plant species selected. In cases where both a specialised N- and a specialised P-uptake strategy seem important (e.g. N fixing and hairy root clusters; or N-fixing and mycorrhizal associations), give both categories. If there is good evidence to classify a species in one of the categories 5 through 11, there is no need to further test for categories 1 through 4. For most strategies, useful data are also available in the literature for many species. For instance, Harley and Harley (1987a; 1987b; 1990), Wang and Qiu (2006) and Brundrett (2009) have information on mycorrhizal associations of many temperate European species belonging to different families. Mabberley (1987) has some general information for a huge number of genera and families (additionally Sprent 2001 gives a long list of nodulation in legumes).

What and how to collect? (Categories 1 through 4)

To check for N2-fixing capacity and mycorrhiza, dig up a minimum of five (preferably 10) healthy looking plants during the growing season, from typical sites for each of the predominant ecosystems studied. If possible, use the same plants used to determine belowground allocation (see main text 2.9. Root mass fraction). Plant roots need to be carefully washed and soil particles removed by rinsing or with fine forceps. It is important to use roots that are attached to the plant; otherwise there is the risk of mixing roots of different plant species.

Storing, processing and observations (Categories 1 through 4)

Washed roots can be stored at 4 °C for several days before further cleaning and staining procedures start. N fixing root nodules and ecto-mycorrhizal roots can be identified visually at lower magnification under a dissecting microscope (see description in strategies 1 and 3, below). Arbuscular, ericoid and orquidoid mycorrhizal fungi inhabit the inside of the roots and the procedures to determine root colonization by these fungi are more elaborate. Clear and more detailed descriptions of the procedures explained below are given by Brundrett et al. (1996). The species belongs to one of the categories 1 through 4 below if the relevant structures are clearly seen in at least a third of the plants, or in at least two plants if only five plants are sampled.

1) Nitrogen fixing species: Check for nodules or other specialized structures on washed root systems under the dissecting microscope. The roots of many legumes (Mimosaceae, Fabaceae/Papilionaceae, Caesalpiniaceae) contain mostly globose or semiglobose root nodules of diameters 2 - 10 mm (Corby 2000); however several legume families are rarely nodulated. Finger-like elongated forms also occur. The number of root nodules can vary greatly; some roots are almost 'covered' with nodules while on other roots they are sparsely distributed. Nodules tend to be clearly pink, or sometimes red or brown (rarely black) in colour while active N fixation is taking place.

Be aware that (a) some legume species do not form symbiotic root nodules; (b) root nodules with symbiotic rhizobia bacteria have also been reported from Ulmaceae (Trema cannabina) and Zygophyllaceae (Zygophyllum spp., Fagonia arabica, Tribulus alatus), while they have been suspected to occur in some other families as well (Becking 1975); and (c) some legume species (e.g. Sesbania in tropical forests) bear the nodules on the stem. Moreover, non- fixing root nodules can be colonized by mycorrhizal fungi (Scheublin and van der Heijden 2006) and some plant species even form “mycorrhizal root nodules” (Rusell et al. 2002).
Other root structures that host N fixers are the 'actinorhiza', found in some members of other vascular plant families. These symbiotic nodules, also called rhizothamia (Lambers et al. 2009), usually contain N-fixing Actinobacteriomycetes (or Actinomycetes) of the genus Frankia, and have a different morphology from legume nodules. Some taxa feature coralloid nodules (the Alnus type, which are similar to those found in Cyca), while other taxa have upward pointing nodules extending into upward pointing rootlets (the Casuarina / Myrica type). Good photographs of these types can be found in Becking (1975). Be aware that there are also plant taxa that feature nodule-like structures without N-fixing symbionts (Becking 1975).

Some further vascular plants host N-fixing bacteria (Nostoc, Anabaena) in looser structures, notably the water fern Azolla, Gunnera and some members of the Cycadaceae (cycads). Some tropical grasses also form loose associations with N-fixing bacteria (Wullstein et al. 1979; Boddey and Döbereiner 1982). Moreover, several plant species (e.g. sugar cane) contain endophytic bacteria that can fix nitrogen. It has been reported that such endophytic bacteria can fix up to 150 kg N per hectare per year (Boddey et al. 1995). Stable isotopes (15N) in combination with molecular tools (e.g. targeting nitrogen fixation genes) can be used to verify the presence and importance of endophitic N-fixers.

2) Arbuscular mycorrhiza (AMF) (SM 2. Fig. 1a-c): This type of mycorrhiza needs a more complex procedure to be detected:

(a) Clear the roots in a 10% potassium hydroxide (KOH) solution at 90°C in a water bath. Alternatively, cold staining could be achieved by clearing roots in 20% potassium hydroxide (KOH) for 24-48 h. Clearing time depends on root age and plant species and varies from 5 min for young herb roots collected in pot experiments, to 1 h for old roots from the field. If roots are left for too long they might disintegrate. Clearing is necessary to remove cell contents and pigments. Staining after clearing shows the fungal structures (when present) inside the root.

(b) Next, wash the roots with water and acidify with 10% hydrochloric acid for 10-30 min to remove the potassium hydroxide (this procedure is not always necessary). The washed roots can be stained with traditonal trypan blue solution (0.05% trypan blue in 2:1:1 lactic-acid:water:glycerol) or chlorazol black E solution (0.03% chlorazol black E 1:1:1 lactic-acid:water:glycerol). Pen ink vinegar (5% ink in 95% vinegar) and Aniline blue solution (0.25 g aniline blue in 25 ml water and 475 ml lactic acid) are equally effective and far less hazardous than the other two solutions. Staining needs to be done in a water bath at 90°C for 20 min or 24-48 h at ambient temperature (or shorter with young fragile roots from pot experiments).

(c) Wash the stained roots again with water and store and destain the roots in a glycerol solution. Trypan blue is carcinogenic and it needs to be recollected after use. Use gloves when clearing and staining!

(d) Cut thin longitudinal sections (app. 1-2 cm long) of 25 root pieces per root system. Note this needs to be done before the roots are processed.

(e) Examine the roots under the microscope at 100x magnification. The degree of mycorrhizal colonization varies depending on staining agent, plant species, soil type and soil nutrient availability (Gange et al. 1999; Vierheilig et al. 2005; Smith and Read 2008).
Hyphae typically spread longitudinally between cortical cells within the intercellular spaces. In some cases hyphae also penetrate cortical cells and spread from cell to cell. Usually many hyphae can be observed in a longitudinal section of a root under the microscope (SM 2. Fig. 1a-c). AMF are characterised by arbuscules, extensively branched tree-like structures that are formed within cortical cells of young roots and function as nutrient exchange organs. Arbuscules are often difficult to detect in field roots since they have, in most cases, a limited life span. Arbuscules have a granular appearance under the microscope (SM 2. Fig. 1a). Intracellular hyphal coils could also be common in roots of certain species and are believed to have analogous function to arbuscules (SM 2. Fig. 1b). Vesicles, swollen structures of variable size and shape within the intercellular spaces, are formed by some AMF fungi and are, when present, a good indicator for AMF infection. Vesicles are thought to have a storage function and contain small lipid droplets that sometimes can be detected under the microscope (SM 2. Fig. 1c). It may be difficult to distinguish AMF from other root colonizing fungi, for example dark septate endophytes (see Jumpponen 2001). Hyphae from members of the Basidiomycotaetes and Ascomycotaetes (two abundant phyla of fungi in soils) contain hyphal septa at regular distances, while septa are mostly absent in AMF.

3) Ecto-mycorrhiza (EMF) (SM 2. Fig. 1d-e): Parts of the root system of ecto-mycorrhizal plants are surrounded by a mantle of fungal hyphae, which have replaced any root hairs. Ecto-mycorrhizal roots are typically swollen and often, but not always, dichotomously branched (SM 2. Fig. 1d-e). Ecto-mycorrhizal fungi differ from AMF in that the largest part of the fungus remains outside the root. Many different ecto-mycorrhizal structures have been observed depending on the identity of fungus and plant host. The colour atlas of ecto-mycorrhizae (Agerer 1986-1998) shows many types and species of ecto-mycorrhiza. Ecto-mycorrhizal structures can be further examined under the microscope. A thin cross section of a plant root can be made with a sharp razor blade and subsequently be stained with chlorazol-black (see strategy 2 above). Such a section typically shows the mantle at the root surface and a Hartig net of fungal hyphae surrounding root cortex cells within the root. An additional useful (but not exclusive) trait is the clear 'fungal' smell that some ecto-mycorrhizal roots have. Also, many ectomycorrhizal fungi produce conspicuous epigeous fruiting bodies (including many of the well-known toadstools), which may give a first suspicion about the possible ectomycorrhizal status of neighbouring plants. Molina et al. (1992) listed the families and genera of such fungi.

EMF are particularly common in a range of plant families, including for instance Betulaceae, Caesalpineaceae, Dipterocarpaceae, Fagaceae, Myrtaceae, Nyctaginaceae, Pinaceae and Salicaceae.

4) Ericoid mycorrhiza: Virtually all genera and species belonging to the families Ericaceae (except Arbutus and Arctostaphylos, which are usually arbutoid Mycorrhizal, see strategy 6 below, and Smith and Read 2008, for an extensive discussion), Empetraceae and Epacridaceae can be assumed to host ericoid mycorrhizal fungi under natural conditions. These mycorrhizas are not yet known from other families. Most of the genera are ericaceous (dwarf) shrubs linked with strongly organic soils such as those found in tundra, heathland, Mediterranean-type shrubland and boreal forest. The same staining methods described for arbuscular mycorrhyzas apply for ericoids to observe the typical hyphae and intracellular coil hyphae complexes (SM 2. Fig. 1f).

5) Orchid roots: All species of orchid (Orchidaceae) appear to depend strongly on association with orchid mycorrhizal fungi for their establishment under natural conditions. Therefore, any Orchidaceae species can be assumed to form these mycorrhizas and belong to this category. The same staining methods described for arbuscular mycorrhyzas apply for orchid to observe the typical intracellular hyphal coils or pelotons (SM 2. Fig. 1g).

6) Rarer types of mycorrhiza, e.g. arbutoid mycorrhiza (Arbutus, Arctostaphylos), ecto-endo-mycorrhiza (certain gymnosperms) and pyroloid mycorrhiza (Pyrolaceae). Details on these particular types and their detection are provided in Smith and Read (2008).

7) Myco-heterotrophs: If a plant species does not contain chlorophyll (i.e. shows no sign of greenness, during any phase in its life cycle), it can safely be classified as a heterotroph. Myco-heterotrophs derive carbon and nutrients from dead organic matter or living green plants via mycorrhizal fungi of various types. They should not be confused with holoparasites that directly parasites plant roots or shoots (see below, strategy 9). Since Myco-heterotroph plants have been studied well, we recommend to revise the comprehensive overview of plant families and genera worldwide with myco-heterotrophic members presented in the literature.

8) Root/stem hemiparasites: These are green plants whose roots tap into the roots of a host plant. Careful microscopic examination of the root system of a plant may reveal connections with a host plant, but this is very hard to verify without digging up hemiparasite and host plant simultaneously. Therefore, given that this group has been reasonably well studied, it may be wise to only check for parasite-host connections within the Scrophulariaceae, and particularly the subfamily Rhinanthoideae. This is the only higher taxon that has both parasitic and non-parasitic members. Within this subfamily, Bartsia, Buchnera, Castilleja, Euphrasia, Melampyrum, Pedicularis, Rhinanthus and Tozzia are safely classified as hemiparasitic, while Digitalis, Hebe and Veronica are not parasitic. Other known root hemi-parasitic families are Olacaceae, Opiliaceae, Santalaceae, Loranthaceae and Krameriaceae. Some species belonging to these families that are not shoot parasites have been found to be root hemiparasites.

9) Holoparasites: Holoparasites directly parasitise the roots or shoots of other species. For instance, all achlorophyllous plant belonging to: a) families: Balanophoraceae, Orobranchaceae (also classified as Scrophulariaceae) and Rafflesiaceae, and b) genera: Cuscuta, Hydnora, Prosopanche, Cassytha, Ammobroma, Lennoa, Pholisma, Mitrastemon, Harveya, Lathraea, Striga, are known to be holoparasites (but see SM 2 Table 1 for more references).

10) Carnivorous plants: Look for obvious specialised organs to capture preys (external digestive glands (often sticky), as well as showy appendages or other features to attract invertebrate animals), or the captured preys themselves, Utricularia is a specialised aquatic genus. New carnivorous species are being discovered even in this century, among families with no other known carnivorous species (e.g. Philcoxia sp., Plantaginaceae).

11) Hairy root clusters (proteoid roots, cluster roots, capillaroid roots or dauciform roots): Under the (dissecting) microscope, look for 'distinct clusters of longitudinal rows of contiguous, extremely hairy rootlets' (Lamont 1993; Shane et al. 2005), or 'a region of the primary or secondary root where many short rootlets are produced in a compact grouping, giving the appearance of a bottle brush' (Skene 1998). Examples for hairy root clusters can be found in Lambers et al. (2006). Careful examination is especially recommended for species belonging to families known to feature members with hairy root clusters. First get familiar with their appearance by checking roots of plants known to contain them (e.g. see Shane et al. 2005 for pictures and a list of Australian rush, reed and sedge species that form dauciform roots).

12) Specialised strategies (mostly in epiphytes)

a) Tank plants (ponds): Within the tropical Bromeliaceae family, look for rosettes of densely packed leaves that, together, create a 'pond' in which rain or run-off water collects. Different species may feature roots growing into these tanks or trichomes (see point d) below) on the surface of the inner leaf bases. See Martin (1994) for details. Most tank bromeliads are epiphytes, but there are also terricolous species, for instance in salinas (where the tanks may keep salt water out).

b) Baskets: Diagnostic are big leaf rosettes of epiphytic plants (often in big tree forks) that capture humus effectively. There are important representatives of this strategy within the ferns (Pteridophyta) and the Araceae family.

c) Ant nests: This form results from symbiotic relationships between plants and ants. Usually the ants transport seeds of ant nest plants to the 'nests', where these germinate and benefit from nutrients in other materials imported by the ants, and their faeces. In return, the plants may offer nectar, fruit and accommodation to the ants. Ant nest plants are found in several families, including Orchidaceae, Bromeliaceae, Asclepidiaceae, etc. In other cases, plants host ants inside special organs such as pseudobulbs, tanks or pitchers (Longino 1986; Heil and McKey 2003). Often more than one plant species inhabit an epiphytic ant nest. Moreover, ant also protect plants against herbivores (e.g. Wimp and Whitham 2001)

d) Trichomes: These are specialised epidermal water-absorbing organs on the leaves of various Bromeliaceae (Benzing et al. 1985) and members of some other families with poorly developed root systems (e.g. Malpighiaceae). Trichomes are usually recognisable as conspicuous whitish scales. Their main function is probably to absorb water and nutrients, but they may also prevent overheating by reflecting sunlight in exposed habitats, deter invertebrate herbivores and/or promote gas exchange. Note that not all trichomes have this function.

e) Root velamen radiculum: Look for a conspicuous spongy, white (especially when dry) or sometimes green cover of the aerial roots of certain light-exposed epiphytic orchids (Orchidaceae) and aroids (Araceae), but also check other families (Porembski and Barthlott; 1995).

12) No specialised mechanism: Only assign this category after careful checking for categories 1 through 11.

Special cases or extras

(1) Natural abundance of 15N. The natural abundance of the nitrogen isotope 15N can be used to assess the fraction of nitrogen which plants obtain from nitrogen fixing symbionts (see Hogberg 1997 for a discussion; see van der Heijden et al. 2006 for an example).

(2) Rarer types of mycorrhiza. For some rarer types of mycorrhiza, e.g. arbutoid mycorrhiza (Arbutus, Arctostaphylos), ect-endomycorrhiza (certain gymnosperms) and pyroloid mycorrhiza (Pyrolaceae) consult Molina et al. (1992) or Smith and Read (2008).

(3) Nutrient uptake in non-mycorrhizal species. There are also non-mycorrhizal vascular higher plant species capable of uptake of organic nutrient forms (e.g. Chapin et al. 1993), but these cannot be identified without detailed investigation involving element isotopes.
Hemiparasites with haustoria tapping into tree branches are treated under 2.3. Growth form (main text).

(4) The following list of plant families which are never or rarely mycorrhizal may be helpful: Aizoaceae, Amaranthaceae, Brassicaceae (Cruciferae), Caryophyllaceae, Chenopodiaceae, Comelinaceae, Cyperaceae, Fumariaceae, Juncaceae, Nyctaginaceae, Phytolacaceae, Polygonaceae, Portulacaceae, Proteceae, Urticaceae. However, exceptions may occur! See Wang and Qiu (2006), Akhmetzhanova et al. (2012). Updated information regarding this topic can be found at

References on theory, significance and large datasets: Kuijt (1969); Benzing (1976); Lüttge (1983); Sprent and Sprent (1990); Read (1991); Lamont (1993); Leake (1994); Pennings and Callaway (1996); Michelsen et al. (1998); Press (1998); Skene (1998); Spaink et al. (1998); Gutschick (1999); Hector et al. (1999); Aerts and Chapin (2000); Gualtieri and Bisseling (2000); Cornelissen et al. (2001); Grime (2001); Squartini (2001); van der Heijden and Sanders (2002); Lamont (2003); Quested et al. (2003); Read and Moreno (2003); van der Heijden et al. (2006); Lambers et al. (2008); Smith and Read (2008); Marschner (2012).

More on methods: Böhm (1979); Agerer (1986-1998); Somasegaran and Hoben (1994); Brundrett et al. (1996);

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