A starting guide to root ecology: strengthening ecological concepts and standardising root classification, sampling, processing and trait measurements

Abstract

In the context of a recent massive increase in research on plant root functions and their impact on the environment, root ecologists currently face many important challenges to keep on generating cutting-edge, meaningful and integrated knowledge. Consideration of the below-ground components in plant and ecosystem studies has been consistently called for in recent decades, but methodology is disparate and sometimes inappropriate. This handbook, based on the collective effort of a large team of experts, will improve trait comparisons across studies and integration of information across databases by providing standardised methods and controlled vocabularies. It is meant to be used not only as starting point by students and scientists who desire working on below-ground ecosystems, but also by experts for consolidating and broadening their views on multiple aspects of root ecology. Beyond the classical compilation of measurement protocols, we have synthesised recommendations from the literature to provide key background knowledge useful for: (1) defining below-ground plant entities and giving keys for their meaningful dissection, classification and naming beyond the classical fine-root vs coarse-root approach; (2) considering the specificity of root research to produce sound laboratory and field data; (3) describing typical, but overlooked steps for studying roots (e.g. root handling, cleaning and storage); and (4) gathering metadata necessary for the interpretation of results and their reuse. Most importantly, all root traits have been introduced with some degree of ecological context that will be a foundation for understanding their ecological meaning, their typical use and uncertainties, and some methodological and conceptual perspectives for future research. Considering all of this, we urge readers not to solely extract protocol recommendations for trait measurements from this work, but to take a moment to read and reflect on the extensive information contained in this broader guide to root ecology, including sections I-VII and the many introductions to each section and root trait description. Finally, it is critical to understand that a major aim of this guide is to help break down barriers between the many subdisciplines of root ecology and ecophysiology, broaden researchers' views on the multiple aspects of root study and create favourable conditions for the inception of comprehensive experiments on the role of roots in plant and ecosystem functioning.

Keywords: below-ground ecology; handbook; plant root functions; protocol; root classification; root ecology; root traits; trait measurements.

Fig. 1

Map of trait categories included in this guide and approximate frequency at which these categories have been studied together. While not all categories are necessarily relevant to study together, this diagram can be used to identify the (lack of) connections between these ‘fields’ of research. The width of connectors depicts weak‐to‐strong linkages between categories. No connector, no or very few studies looking at both fields jointly; thin connectors, few studies; medium connectors, fields sharing substantial number of studies; thick connectors, fields that are frequently studied together. This diagram represents the authors’ expert assessment only and is imperfect as no exhaustive review of the literature was carried out.

Fig. 2

Schematic presentations of generic root nomenclature in monocotyledonous (left side) and dicotyledonous (right side) plants and the corresponding nomenclature as proposed by the International Society of Root Research (ISRR; centre); colours of nomenclature match the respective roots in the drawing; lines indicate superordinate terms. Roots potentially originating from the scutellar node (e.g. in wheat) are not drawn. See Box 1 for further information on root entities and synonyms; drawing not to scale.

Fig. 3

Schematic presentations of common root nomenclatures related to specific root morphological and anatomical traits. (a) Macrorhiza and brachyrhiza (fine roots) and woody coarse roots (example: Tilia sp.; modified after Kubíková, 1967). Macrorhiza, description of layers from periphery to centre: (1) rhizodermis, cortex, stage 0 endodermis, stele with four protoxylem groups; (2) rhizodermis, cortex, stage I endodermis, stele with first metaphloem and ‐xylem; (3) rhizodermis, cortex, stage I–II endodermis, pericycle, phloem with parenchyma, cambium, xylem; Brachyrhiza with ectomycorrhizal symbiont: (1) mycorrhizal mantle, rhizodermis, cortex, stage I endodermis, stele with two protoxylem groups; (2) mycorrhizal mantle, rhizodermis, cortex, stage I–II endodermis, phloem, cambium, xylem; Nonmycorrhized brachyrhiza: (1) rhizodermis, cortex, stage 0 endodermis, stele with two protoxylem groups; (2) rhizodermis, cortex, stage 1 endodermis, phloem, cambium, xylem. Woody coarse root: l) periderm, phellogen, secondary phloem, vascular cambium, secondary xylem. Dividing (cross‐hatch), lignified (filled) or suberised (horizontal hatch) tissues and hyphal mantel (diagonal hatch) are indicated. (b) Cluster root with two groups of abundant, short lateral roots (rootlets) with root hairs. (c) Taproot, sinker and horizontal roots in a schematic tree root system. See Box 1 and text for further information on root entities and synonyms; drawings not to scale.

Fig. 4

Schematic presentations of systematic root classification approaches. (a) Developmental approach; (b) morphometric approach; and (c) topological approach. Root growth axis (a) and root order (b, c) levels are indicated by adjacent numbers; note that highest root orders are 5, 4 and 32, respectively (a–c), corresponding to 6, 4 and 12 classification levels (a–c). The ‘0’ root order in (a) can be replaced by root class (e.g. basal root, tap root); lower root order in (b) can be refined further, for example by adding information on the mycorrhizal status; the highest order number in (c) represents the numbers of root tips. See Box 1 and text for further information on systematic root classification approaches.

Fig. 5

Root branches of three architecturally diverse, co‐occurring subtropical species, demonstrating the functional classification approach (i.e. absorptive and transport fine roots) and the variable number of morphometric (centripetal) fine‐root orders that fall below different diameter cut‐offs (0.5, 1.0, and 2.0 mm). Schima superba (top left) with up to five root orders ≤ 0.5 mm, including both absorptive and transport fine roots, Choerospondias axillaris (middle) with three root orders ≤ 0.5 mm and only including absorptive fine roots, and Cinnamomum austrosinense (bottom right) with no roots ≤ 0.5 mm. Black, absorptive fine roots; grey, transport fine roots (modified after McCormack et al., 2015a).

Fig. 6

The main classes (rectangles) and properties (connections between the rectangles) of the observation and measurement ontology (OBOE) initially developed by Madin et al. (2007), and modified later by Saunders et al. (2011). An observation is made of an entity. The quality of an entity can be represented by a measure. Measures establish a relationship between the characteristics and a measurement standard via a value, and are obtained with a certain precision. Measures are carried out using a protocol in a certain place at a certain time. Observations can have multiple measures. Entities, characteristics and measurement standards constitute entry points for domain‐specific ontologies. The notations ‘1 : 1’ and ‘0 : n’ are called multiplicities: they indicate how many objects within a given class can be linked to objects of another class. For example in the relationship ‘of entity’, an Observation will be linked to only one entity (‘1 : 1’), while an entity could be linked to 0 or to n Observations (‘0 : n’).

Fig. 7

Soil sampling techniques using (a) a metallic frame (0.25 × 0.25 × 0.15 m); (b) a manual core or (c, d) mechanised augers. (e) Monolith excavation provides a large volume of soil, which increases the accuracy of root biomass estimates; (f) manual coring provides small cores usually sampled in successive steps of 10–20 cm depth with a 5–10 cm diameter core; (g) mechanised augers allow deep coring in harsh soils. Image courtesy of C. Roumet (a, b, c, f), M. Zhun (e), F. Khalfallah (g).

Fig. 8

Soil in‐growth core method. A soil core is taken and used for initial root biomass estimation. The core is then filled with similar sieved soil, free of roots, placed into a mesh cylinder. After few weeks of incubation, the mesh bag is removed to estimate the fine‐root biomass production rate.

Fig. 9

Trench profile method: (a) root mapping using a grid fixed on the wall of a 4‐m agroforestry trench to determine (b) the root intersection densities at different soil depths (RID, cm⁻²). Image courtesy of R. Cardinael and C. Jourdan.

Fig. 10

Rhizobox installation and observation. (a) Installation in the field, note that when installed, the windows may be tilted inward by c. 5–10° to facilitate better contact with added sieved soil; (b) diagram of rhizobox construction, an acetate sheet is fixed to the window frame; (c) observations of roots together with fungal hyphae. Images courtesy of J. Pippen (a), M. L. McCormack (b) and I. C. Meier (c).

Fig. 11

(a) Minirhizotron (MR) techniques with image acquisition devices (i.e. digital camera MR or scanner MR) and different options to install the MR tubes, that is angled or vertical from the soil surface or horizontally from trenches; (b) roots of Fagus sylvatica captured with a scanner MR system, CID 600; (c) roots of Pisum sativum taken using the Vienna Scientific MR camera MS‐190. Images courtesy of B. Rewald.

Fig. 12

Anatomical and morphological differences between clonality derived from roots (a) and stems (b).

Fig. 13

Examples of plants with long (a) and short (b) persistence of their below‐ground clonal organ. (b) Below‐ground plant parts during winter rest: black parts of the rhizome system are decaying while white parts are living and will sprout at the beginning of a growing season.

Fig. 14

Classification of root systems for trees and shrubs, according to Krasilnikov (1968). (a) Primary root systems, with 11 different types; (b) secondary and mixed root systems, with eight different types.

Fig. 15

Picture of a piece of a root of Typha latifolia L. (Typhaceae), an herbaceous wetland monocot. This root illustrates the potential root order dimorphism and its consequences for measurements of morphological traits. The pictured root, grown in water, has a total length of 6.33 m, out of which 0.11 m (1.7%) is taken by the thick basal root. Average diameter of the fine lateral roots is 0.21 mm, that of the basal root is 1.66 mm. Average diameter of the whole sample is 0.23 mm. Specific root length of the basal root, lateral roots and total sample are 6 m g⁻¹, 230 m g⁻¹ and 138 m g⁻¹, respectively. The porous basal root with a root tissue density (dry mass per fresh root volume; RTD) of 0.08 g cm⁻³ takes up 53% of the sample volume, while the less porous lateral roots with a RTD of 0.125 g cm⁻³ take up 47% of the volume, resulting in an average RTD of 0.10 g cm⁻³.

Fig. 16

Typical greyscale images used for root morphological trait analysis. Roots from three herbaceous species with contrasting root types are displayed, a grass (a) Bromus erectus Huds. (Poaceae); a forb (b) Sanguisorba minor Scop. (Rosaceae); and a legume (c) Lotus corniculatus L. (Fabaceae). Note that nodules are visible on the legume roots but are typically removed for morphological analyses of roots (and nodule biomass should be separately assessed, see section XX. 2. Nodule investment ). Also, note that roots should not be allowed to overlap too much for accurate length, diameter and volume estimations: this can be achieved by clipping the roots into subunits.

Fig. 17

Root cross‐section through different growth zones showing primary (a–c) and secondary (d–f) development of Populus trichocarpa roots. Roots were fixed and embedded in Technovit, sectioned using rotary microtome and stained with toluidine blue using the protocol described under section XIII. 1. a. Generalities of microtechniques used for the analysis of root anatomy . Traits that can be assessed through such serial images include: root diameter, cortex thickness, cortex and stele area fractions, cortex‐to‐stele area ratio, primary/secondary xylem differentiation distance from root tip, archic structure (here: triarch), the number of xylem poles in primary xylem, number of conductive elements per root section or per xylem pole, conduit diameter, conduit wall thickness, cell wall thickness ratio to conduit diameter, root specific hydraulic conductance (Lx), critical tension for conduit collapse (t/d)². cx, cortical parenchyma cells; px, primary xylem (within ellipses); sx, secondary xylem; sr, secondary growth of root. Bars, 100 µm.

Fig. 18

Visualisation of exodermis in Liriodendron tulipifera and Populus trichocarpa roots; exodermis autofluorescence (a, b) and root sections stained with berberine and aniline blue (c, d). Cs, Casparian strips; sl, suberin lamellae; pc, passage cells (asterisks). Bars, 100 µm.

Fig. 19

(a) A section of root is clamped between the jaws of a Universal Testing Machine and stretched until failure occurs in tension; (b) during a tensile test, stress (σ) and strain (ε) increase quasilinearly and the root has an elastic behaviour. Modulus of elasticity (MoE) can be calculated from the slope of the linear relationship between stress and strain. Once permanent deformation starts (yield stress), the root undergoes nonlinear plastic behaviour. The point at which ultimate failure occurs is used to calculate tensile strength. Note that yield stress can be difficult or even impossible to observe during a test and usually only a small inflection point occurs in the slope of stress and strain.

Fig. 20

Survivorship curve and comparison between small and large diameter roots observed with the minirhizotron technique. Note that the point at which the survival probability reaches 0.5 (i.e. 50%) represents the estimated median lifespan for the population. Therefore, the median lifespan for small and large diameter roots would be interpreted as c. 460 and 630 d, respectively. Survivorship curves were generated using Kaplan–Meier regression. Each circle represents the timing of death of an individual root while roots that have unknown death dates treated as censored and estimated. Log‐Rank and Wilcoxon tests are used to identify significant differences in the survivorship behaviour between groups. Figure taken from McCormack et al. (2010).

Fig. 21

Typical workflow of specific root respiration (RR_S) measurements with closed‐chamber, gas‐phase CO₂ (upper) and aqueous‐phase O₂ (lower) gas‐exchange systems. (1) Roots are retrieved, rinsed and allowed acclimate to temperature. (2) Roots (or root segments) are inserted into temperature‐controlled, closed chambers to record changes in relative CO₂/O₂ concentrations, using either an infrared gas analyser (IRGA) or an O₂ electrode. A ‘dead band’ illustrates the importance of chamber equilibration (mixing and temperature); subsequently, the slope of the gas concentration curve (δCO₂ or δO₂/δt; ppm s⁻¹) is calculated by either linear or exponential regressions methods. For gas‐phase measurements, linear regressions often underestimate the slope; for aqueous‐phase O₂ measurements, a quasilinear area of the curve is frequently visually selected for slope calculations. (3) The absolute gas concentration (mol m⁻³) is calculated by using either the ideal gas law (gas‐phase measurements; in retrospect) or Henry’s law (for dissolved gas; often integrated in O₂ electrode software) together with information on chamber/solute volume (m³), air pressure (Pa) and temperature (K). (4) Absolute flux rates (µmol s⁻¹) are divided by root dry mass (g), or other size parameters, to calculate RR_S (e.g. µmol g⁻¹ s⁻¹). See text for details.

Fig. 22

Different methodological approaches are used to quantify soil resource uptake, as a physiological root trait. Roots can be excised from the plants and incubated ex situ or still attached to the plant in nutrient solutions in situ to characterise the properties of the uptake system. Tracer addition to the soil allows estimates of nutrient uptake of plants within the soil system. *The radioisotope method is not covered in this handbook.

Fig. 23

Experimental design adopting a multiple tracer study to quantify resource uptake rates along spatial (soil depth), temporal (season) and chemical (water, nitrogen, cations) niche axes (Jesch et al., 2018). The spatial and chemical axes are quantified in the same subplot, whereas different subplots are used to assess temporal niche differentiation. Reproduced with permission from the British Ecological Society.

Fig. 24

Two examples of tracer application by injection into the soil around target plants (a) or within a specific area (b). Images courtesy of M. Scherer‐Lorenzen (a) and A. Jesch (b).

Fig. 25

(a) Roots of Molinia caerulea colonised by arbuscular mycorrhizal (AM) fungi (species unknown) with visible hyphae; and (b) arbuscules and vesicules; captures with stereomicroscope (×3–5 magnification). (c) Wheat roots (Triticum aestivum L.) colonised by AM fungi Rhizophagus irregularis with visible hyphae, arbuscules and vesicules; and (d) spores and extraradical (i.e. outside of roots) mycelium of Rhizophagus irregularis. Note that vesicules (c) are very similar to fungal spores, but the latter can be distinguished because they are typically round and not connected to the hyphal network. Images courtesy of M. Bakker (a, b, stereomicroscope, ×3–5 magnification); and M. Rebeca Cosme (c, d, compound microscope, ×200 magnification).

Fig. 26

(a) Ectomycorrhizal fungi (EMF) structures on roots of Pinus pinaster Aiton showing ectomycorrhizal fungi (ECM) root tips; (b) mantle and extraradical hyphae; and (c) mantle. Images courtesy of C. Guérin and M. Bakker; stereomicroscope, ×3–5 magnification.

Fig. 27

Microscopic examination of roots to quantify arbuscular mycorrhizas (AM) (adapted from Brundrett et al., 1996).

Fig. 28

Schematic view of a root tip and suggested measurements. LANZ, length of apical nonbranched zone; LAHZ, length of apical hairless zone. Redrawn from Pagès et al. (2010).

Fig. 29

Stages of root hair development in Arabidopsis thaliana. Root hairs emerge in the root differentiation zone (numbers 1–4 of the middle panel) and progressively increase in length the further they are from the root cap. Numbers in the outer panel are high magnification differential interference contrast micrographs of root hairs corresponding to the numbers in the middle panel.

Fig. 30

Protocols for phenotyping root hairs in a large number of plants. (a) Imbibed wheat seeds are aligned in a row on moist germination paper placed on top of a tray; or (b) the paper can be rolled and inserted into cones. (c) After 10 d of vertical growth, root hairs are ready to be imaged with a stereomicroscope. (d) Representative images of unstained wheat root hairs for length and density measurements. Bar, 500 µm.

Fig. 31

Protocol for preparing Arabidopsis seedlings for live root hair imaging. (a) All material for handling seeds is sterilised and prepared in a laminar flow hood. (b, c) Sterilised seeds are picked individually with a pointed toothpick and planted on nutrient‐supplemented gel layered onto a coverslips. (d) Coverslips with seeds are placed inside Petri dishes and kept at an inclined angle in a growth chamber. (e) After 3–5 d, the coverslips with seedlings can be transferred directly onto a stage of a microscope for image acquisition.