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Biology

Multipronged Phenotyping Approaches to Characterize Sugarcane Root Systems

Published: August 17, 2022 doi: 10.3791/63596

Abstract

Roots are the primary conductors of water and nutrients and play a vital role in sustaining growth and yield under stressful environments. The study of plant roots poses methodological difficulties in in situ assessment and sampling, which is especially true for sugarcane (Saccharum spp.). Traditional methods during the 1920s documented the genotypic variation in sugarcane root systems, after which few studies were reported on sugarcane root traits per se until recently. In addition to morphology, rhizosphere characteristics, including allelopathic effects and/or affinity for microbial symbiosis, determine plant establishment and survival.

Ultimately, root systems define the above-ground productivity of sugarcane. With the impetus for climate-resilient varieties, it is becoming more relevant to explore and utilize the variability in root system traits of sugarcane. This paper describes multipronged approaches for sugarcane root phenotyping, including field excavation by trench sampling, the use of a root core sampler, raised platforms for root sampling, and raising plants under hydroponic culture, employed by a team of scientists at the Indian Council of Agricultural Research-Sugarcane Breeding Institute (ICAR-SBI).

Field excavation by trench sampling is imperative to assess the plant roots in their natural growing environment. The use of raised platforms simulating field conditions and a root core sampler are alternative approaches, with a considerable reduction in time, uniform sample size, and less loss of root material. Hydroponic plant culture allows the study of morphology, anatomical features, and rhizosphere biology, including the exudation of organic compounds and microbial interactions. Data generated from different experiments using diverse sampling methods add to the wealth of information on the root system traits of sugarcane.

Introduction

Sugarcane (Saccharum spp.), an important food and bioenergy source, is a significant industrial crop suited for cultivation in many countries with tropical and subtropical climatic conditions. Owing to the C4 photosynthetic pathway for carbon assimilation, sugarcane is highly productive, efficiently using farm inputs such as water and fertilizers. Post-harvest processing of sugarcane yields economically valuable products such as sugar and jaggery, alongside its byproducts-molasses, ethanol, and energy. Sugarcane is produced by nearly 100 countries over an area of 25.97 Mha, which is approximately 1.5% of total arable land. India alone contributes to 16% of world sugarcane production (approximately 306 Mt), with an average productivity of 70 t·ha−1 1. The major abiotic stresses drastically affecting sugarcane production include water deficits, water logging, temperature extremes, and soil properties such as nutrient deficiency, salinity, sodicity, and alkalinity. The biggest challenge for developing abiotic stress-tolerant crops is to pinpoint specific traits that confer a substantial yield advantage under stress conditions.

Several aspects of sugarcane physiology are poorly understood, including the root-shoot relationship, which drastically affects cane productivity. Sugarcane root is not as well studied as shoots, although the different types, such as sett roots, shoot roots, and adult roots, may be developmentally distinct with varying functions. Genotypic differences have been observed with regard to the number and length of sett roots emerging during germination2. Sett roots are implicated in the germination of sugarcane buds, ensuring early crop establishment, and are later replaced by shoot roots, which are robust, emerging from the base of the developing shoot3. Fine branches observed in the sett roots help in anchoring the young plants and aid in the absorption of water and nutrients until they are replaced by shoot roots. Similar to sett roots, shoot roots also arise from the root primordia present in the lower, unexpanded internodes of the cane.

As shoot roots persist in the plant for a longer duration, they are 4x-10x thicker than sett roots. Shoot roots constitute the sole root system of sugarcane, with an important role in further growth and development. The vigor of the shoot roots is positively associated with the overall vegetative vigor of the plant. The continuous development of roots resulting from the turnover of sett roots and shoot roots gives rise to the "adult root system" of sugarcane, which is ever-adapting to the prevailing environmental conditions. In general, a deeper, more prolific root system makes more water and nutrients available for the crop than does a shallower distribution of roots. Periodic dissections revealed that, when the soil moisture content was high, shallow root systems were observed, whereas a much deeper root system developed as the water table dropped2. The root system in sugarcane remains active even after harvesting of the crop, contributing to the growth of the ratoon crop until new shoot roots emerge from underground buds4. Root angle and the level of root branching are two important factors determining the volume of soil explored by plant roots. Root angle, a genotypic trait, may be altered through conventional breeding or molecular approaches to improve tolerance to biotic and abiotic stresses. On the contrary, the level of root branching is mostly influenced by the environment, necessitating periodic monitoring of root development and its response to localized soil conditions.

Anatomical features of sugarcane roots have been examined to ascertain differences with regard to genotype and environment. The anatomy of sett roots in sugarcane resembles that of mature roots in other grasses such as maize, wherein the cortex comprises well-differentiated cell layers in a regular pattern. The endodermis is suberized, followed by a single-layered pericycle. Metaxylem elements are the main conductors or water and nutrient ions, radially arranged and interspersed with groups of phloem, the latter comprising a sieve element with two companion cells. The large central mass of undifferentiated cells forms the root pith. Distinct anatomical features of sugarcane cultivars correspond to root hydraulic properties, thereby influencing water movement. Early studies on the differences in the root anatomical traits of sugarcane revealed that, under low moisture stress conditions, pronounced thickening of the cell wall was observed in the layers immediately inside of the endodermis, between the pith and the vascular region, and around the vessels5. Such thickened cells may be an adaptation to prevent the backward flow of sap and for mechanical strength during stress.

Some important traits implicated in the drought resistance of sugarcane include the relative thickness and number of exodermal layers, the ratio of cortex to stele, intercellular spaces in the cortex, and thickened root hair tips. The ratios of the area occupied by cortical cells to the area occupied by the stellar tissues of shoot roots are significantly different among sugarcane cultivars, with wide variability with respect to the area of the stele6. The hydraulic conductivity of sugarcane roots is related to the size and number of metaxylem elements in the shoot roots. Hydrophobic cell layers within the root are likely to define zones of apoplastic water movement. Suberized Casparian bands are found in the endodermis and in the hypodermis (termed as exodermis), which serve as hydrophobic barriers. The disintegration of cortical cells leads to the formation of lysigenous aerenchyma in older roots and in plants subjected to hypoxic conditions, irrespective of developmental age. The formation of aerenchyma during waterlogging stress is correlated with the maintenance of growth in resistant varieties7.

The morphology and anatomy of Erianthus arundinaceus [Retzius] Jeswiet (genera related to Saccharum spp.) roots are implicated in its strong tolerance to environmental stresses8. Erianthus arundinaceus roots exhibit nodal roots distributed at steep angles, with dense roots hairs to facilitate the uptake of water and nutrient ions from deeper soil zones. The deep-root system consists of many nodal roots growing with steep growth angles. The diameter of the nodal roots correlates with the size and number of large xylem vessels, the former varying widely from 0.5 mm to 5.0 mm. These nodal roots also form a rhizomatous sheath, with a hypodermis showing lignified sclerenchyma in the outer cortex (exodermis), lysigenous aerenchyma in the mid-portion of the cortex, and starch granules in the stele. In addition to architecture and morphological traits, root-exuded organic compounds play an important role in determining plant germination, establishment, and survival, with plausible allelopathic effects and/or affinity for microbial symbiosis.

Root enzymatic activity and the finer details of the morphology, including root cap pigmentation and rejuvenation potential upon injury, were documented in sugarcane varieties grown under hydroponic culture9. Root growth shows a highly plastic response to changes in the soil environment, both in terms of the form and size of the root system. The most efficient sugarcane variety would be one that has few or an optimal number of shoots, with a correspondingly lower number of roots, aiding better survival during stressful conditions. The systematic study of the root system should, thus, form an important component of any crop improvement program10. The majority of the experiments focusing on roots rely mostly on developmental aspects, while a focus on functional plasticity is often lacking11. Apart from the structural distribution, functional root plasticity plays a crucial role in survival under stress and would, therefore, support breeders in their efforts to include root system traits in the selection pipeline for abiotic stress tolerance and improve the robustness of sugarcane.

Considering its importance in sustaining growth and yield under stressful environments, it is essential to explore and utilize the inherent variability in root system traits of sugarcane. An emphasis on the selection of component traits and mechanisms imparting superior root systems is the way forward for better crop performance under changing climatic conditions. Phenotypic evaluation is a long and costly process; however, the integration of multipronged approaches would add tremendous value to its utility in crop improvement. In this manuscript, four different approaches for root phenotyping in sugarcane are described, each with its own set of merits and demerits, implying that a concerted effort is required to arrive at comprehensive and holistic results.

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Protocol

1. Field excavation by trench sampling

  1. Raise commercial sugarcane hybrids (hereafter called "varieties") in the field from two-budded setts planted at a row spacing of 120 cm, with 90 cm within rows, following the recommended package of practices (POP) to ensure good crop establishment and growth.
  2. At the end of the maturity phase of the crop, employ an excavator to dig a trench (1.5 m deep and 1.0 m wide) in the field. Through continuous water jetting, ensure that the soils from root zones are cleared without damaging the roots (Figure 1).
  3. When the adhering soil loosens, uproot the cane along with the root system and take it to the laboratory for manual measurement of the number of roots, root length, volume, and weight.

Figure 1
Figure 1: Trench sampling method for the field excavation of roots. (A) Sideways trench dug along the field, (B) water jetting, and (C) inner view showing the depth of the trench. Please click here to view a larger version of this figure.

2. Root core sampler to reduce sampling errors

  1. Use a cylindrical root core sampler of 61 cm height and 16 cm diameter weighing 8 kg, fabricated using mild steel (MS) material for sugarcane root sampling in the field. Provide a sharp edge at the bottom edge of the sampler to facilitate easy penetration while inserting it into the soil. Provide the top with collars of 3 cm diameter to lift the sampler (Figure 2).
  2. Raise sugarcane varieties in the field from two-budded setts planted at a row spacing of 120 cm, with 90 cm within rows, following the recommended POP to ensure good crop establishment and growth.
  3. At the maturity phase of the crop, fasten the top edge of the sampler to the primary shoot/cane and hammer continuously to reach the desired soil depth (45 cm). Lift the entire soil mass into the sampler and wash carefully under running water to separate the adhering roots.
  4. After thorough washing of the roots, record the volume, surface area, length, and weight by manual measurement, as well as by spreading the roots on transparent trays to scan and analyze the corresponding digitized images using the referenced software (see the Table of Materials).

Figure 2
Figure 2: Root core sampler. (A) Dimensions, (B) top view, and (C) site of sampling. Please click here to view a larger version of this figure.

3. Root phenotyping structure to facilitate sampling at different phenophases

  1. Construct a root phenotyping structure comprising three adjacent compartments of dimensions 4.5 m x 10.1 m for sampling of the sugarcane roots, with provisions for manually dismantling the side walls to reveal the underground root system (up to a depth of 80-100 cm) (Figure 3). Construct the side walls with precast slabs of dimensions 1.8 m in length, 30 cm in width, and 4 cm in thickness.
  2. Fill and compact the structure with field soil, leaving a headspace of ~20 cm, with adequate drainage holes to facilitate soil aeration.
  3. Sow the bud chips of germplasm clones comprising Saccharum officinarum L., Saccharum spontaneum L., Saccharum barberi Jesw., Saccharum sinense Roxb., and Saccharum robustum Brandes and Jeswiet ex Grassl. (hereafter called "Saccharum spp. clones") and allow them to germinate for 30 days in protrays comprising rooting media (red soil: farm yard manure: sand = 2:2:1). Transplant uniform and healthy settlings to the structure at a row spacing of 90 cm, with 60 cm within rows, following the recommended POP to ensure good crop establishment and growth.
  4. During the formative (60-120 days after planting [DAP]) and grand growth (120-150 DAP) phases, manually remove the side walls made of the precast slabs, followed by continuous spraying of the water jet to expose the roots.
  5. Uproot the entire root system and take it to the laboratory for manual measurement of the length, volume, and weight, and spread the roots on transparent trays to scan and analyze the corresponding digitized images in the referenced software (see the Table of Materials).
  6. Impose drought stress by withholding irrigation in one of the compartments, and plug the drainage holes in the second compartment to maintain soil saturation to simulate waterlogging stress. Irrigate the third compartment according to the recommended POP to maintain the field capacity and to serve as a control.

Figure 3
Figure 3: Root phenotyping structure. (A) Dimensions, (B) overview of the three compartments, and (C) view of one compartment. Please click here to view a larger version of this figure.

4. Hydroponic culture of plants to study rhizosphere biology

  1. Fabricate an in-house hydroponic system in an environment-controlled glasshouse conducive for growing sugarcane to study the finer details of root biology. Add ~15 L of modified Hoagland's nutrient solution (Table 1) to glass tanks of dimensions 20 cm x 20 cm x 50 cm, with aeration provided by aquarium pumps (Figure 4).
  2. Sow the bud chips of sugarcane varieties and Saccharum spp. clones and allow them to germinate for 30 days in protrays comprising composted coir pith. Transplant uniform and healthy settlings to hydroponic tanks at the frequency of three settlings per tank, taking care to place the entire root in the nutrient solution. Cover the tanks with black cloth to ensure that roots are not exposed to light. Use a plastic mesh (20 cm x 20 cm) at the brim of the glass tanks to support the plants upright.
  3. At the end of the germination phase (60 days), collect the root exudates by immersing the roots of intact plants in 50 mL of trap solution (sterile double-distilled water) for 4 h (0800 h to 1200 h) during the time of peak photosynthetic activity. Filter the collected solution through Whatman filter paper, then pass it through glass columns filled with anion-exchange followed by cation-exchange resins12. Evaporate the eluted fractions to dryness and store at -20 °C until further processing.
  4. Analyze the processed root exudate samples by HPLC for the determination of organic acids12 and by spectrophotometry for the estimation of total phenolics13, proteins14, sugars15, and amino acids16 according to standard protocol.
  5. Monitor the root growth at weekly intervals to record the root tip pigmentation and root hair density. Assess the activity of the enzymes, peroxidase17 and superoxide dismutase18, and total phenolic content19 at the 3rd month according to standard protocol.
  6. Assess the response to root injury by inflicting a longitudinal slice in the primary root up to the root tip using a sterile surgical blade, and monitor the changes periodically.
Chemical Final concentration
Potassium nitrate 0.608 g·L-1
Calcium nitrate 1.415 g·L-1
Potassium dihydrogen phosphate 0.164 g·L-1
Magnesium sulphate 0.560 g·L-1
EDTA-ferric monosodium salt 6.00 g·250L-1
Boric acid 1.43 g·250L-1
Manganese chloride tetrahydrate 0.91 g·250L-1
Zinc sulphate 0.11 g·250L-1
Cupric suphate 0.04 g·250L-1
Molybdic acid 0.01 g250L-1

Table 1: Composition of the modified nutrient solution for the hydroponic culture of sugarcane.

Figure 4
Figure 4: Hydroponics setup. The setup (A) customized for growing sugarcane and (B) 5-month-old crop (black cloth removed for photography purpose only). This figure has been modified from Hari et al.9. Please click here to view a larger version of this figure.

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Representative Results

Variation in the root morphological traits of sugarcane varieties
Representative images of the root system in Co 62175, excavated from the field by trench sampling and grown in a hydroponic setup, are presented in Figure 5A,B. Long roots (~100 cm) were observed in the varieties Co 62175 and Co 99006, while Co 99006 recorded the highest root weight (127 g·clump−1). Root traits were recorded using a root core sampler, which revealed that Co 62175 had a superior root system compared to Co 99006 with regard to cumulative root length, root surface area, and root volume (Figure 5C-E). However, the average root diameter was higher in Co 99006 than in Co 62175 (Figure 5F).

Variation in the root morphological traits of Saccharum spp. clones
Morphological traits in the formative phase showed significant variation among the Saccharum spp. clones, with the highest cumulative root length of 39615.72 cm per clump observed in S. spontaneum (IND 85-490), while the lowest was observed in S. sinense (Oshima) (Figure 6A). The highest root surface area per clump (4700.31 cm2) was observed in S. spontaneum (IND 85-490), followed by S. officinarum (Awela Green Sport), and the lowest was recorded in S. sinense (Oshima) (Figure 6B). A similar trend was observed in the case of root volume and average root diameter (Figure 6C,D). The morphology of the roots sampled at the grand growth phase was measured manually and revealed that the longest roots (131 cm) were present in S. robustum (NG 77-237) (Figure 6E), whereas the root dry weight was the highest in S. spontaneum (IND 85-490: 57.4 g·clump−1) and lowest in S. sinense (Oshima: 11.2 g·clump−1) (Figure 6F). Root volume and root-to-shoot ratio followed a similar trend as the root dry weight (Figure 6G,H).

Variation in the root anatomical traits of Saccharum spp. clones
The cross-sectional area of roots at 10 mm from the tip varied significantly among the Saccharum spp. clones, with S. spontaneum (IND 85-490) showing the largest area (27646.8 mm2), while S. sinense (Oshima) recorded the lowest (7539.1 mm2) (Table 2). The major area in S. sinense (Oshima) was occupied by cortical cells, whereas in S. spontaneum (IND 85-490), area occupied by the stele tissue was prominent, which is reflected in the data on the ratio of cortex-to-stele. The number of metaxylem elements ranged from 6 (S. officinarum: Awela Green Sport) to 13 (S. spontaneum: IND 85-490), while the number of air spaces (lysigenous aerenchyma) varied from 12 (S. barberi: Putli Khajee) to 32 (S. spontaneum: IND 85-490). The area occupied by metaxylem elements was highest (791.3 mm2) in S. barberi (Putli Khajee) and lowest (254.3 mm2) in S. sinense (Oshima). The area occupied by aerenchyma was highest (4921.3 mm2) in S. spontaneum (IND 85-490), while S. sinense (Oshima) and S. robustum (NG 77-237) roots were devoid of aerenchymatous cells under the normal irrigated condition. Saccharum sinense (Oshima), S. spontaneum (IND 85-490), and S. barberi (Putli Khajee) exhibited superior root anatomical phenes under drought and waterlogging stress conditions (Figure 7A-C). Long root hairs, sclerenchymatous exodermis, reduced cortical cell layers, increased cortical aerenchyma, increased stele area, and xylem vessel number with large diameter were favorable under drought, whereas increased root diameter and higher cortical cell area with increased aerenchymatous cells enhanced mechanical strength, root porosity, and oxygen diffusion under waterlogging stress.

Variation in the type and quantity of root exudates of Saccharum spp. clones
Among the organic compounds exuded by Saccharum spp. clones during the germination phase, the proportion of total protein was the highest, followed by carboxylates, sugars, phenols, and free amino acids. Saccharum robustum (NG 77-221) showed the highest (5.11 mg·g−1·root fresh weight [RFW]·h−1) total carboxylate exudation, while the lowest (0.65 mg·g−1·RFW·h−1) was recorded in S. spontaneum (IND 85-490), with significant variation in the type and quantity of individual carboxylic acid exuded (Figure 8A). Saccharum officinarum (Awela Green Sport) exuded the highest total phenol levels (0.034 mg·g−1·RFW·h−1), while the lowest (0.002 mg·g−1·RFW·h−1) were observed in the case of S. spontaneum (IND 85-490) (Figure 8B). The highest total protein exudation (3.23 mg·g−1·RFW·h−1) was observed in S. officinarum (Awela Green Sport), while the highest total sugar (0.22 mg·g−1·RFW·h−1) and total free amino acids (0.04 mg·g−1·RFW·h−1) were recorded in S. sinense (Oshima) (Figure 8C-E). The lowest total protein exudation (2.08 mg·g−1·RFW·h−1) was observed in S. spontaneum (IND 85-490), while S. robustum (NG 77-221) exuded the lowest amounts of total sugar (0.05·mg·g−1·RFW·h−1) and total free amino acids (0.01 mg·g−1·RFW·h−1).

Variation in the root tip pigmentation, enzymatic activity, and total phenolic content of sugarcane varieties
The root cap shapes ranged from simple triangle (Co 62175) to dome (Co 86032), while root cap pigmentation varied from light pink (Co 62175) to deep violet (Co 86032), with significant differences in root hair density (Figure 9A-C). Peroxidase activity was highest (40 units·g−1·h−1) in Co 86032 and lowest (26 units·g−1·h−1) in Co 62175, while the trend was reversed in the case of superoxide dismutase activity in roots (Figure 9D,E). Similarly, the total phenolic content was highest (18 µg·g−1) in Co 86032 and lowest (15 µg·g−1) in Co 62175 (Figure 9F).

Variation in the developmental phases of rejuvenation upon root injury of sugarcane varieties
Rejuvenation upon root injury was tested in sugarcane varieties CoC 671 and Co 06022 raised in a hydroponic setup (Figure 10A-I). In CoC 671, secondary roots appeared on the third day of injury, whereas in Co 06022, secondary root initiation occurred only on the tenth day. Further, the primary root of Co 06022 showed symptoms of senescence on the third day, while CoC 671 roots remained relatively healthy. Tertiary roots were also observed on the tenth day in CoC 671.

Figure 5
Figure 5: Root systems of Co 62175 and different morphological traits. Root system of (A) Co 62175 excavated by trench sampling, (B) Co 62175 grown in a hydroponic setup, and morphological traits (C) cumulative root length, (D) root surface area, (E) root volume, and (F) average root diameter recorded using the root core sampler. Data represent mean ± SEM (n = 3). Please click here to view a larger version of this figure.

Figure 6
Figure 6: Root morphological traits. (A) Cumulative root length, (B) root surface area, (C) root volume, and (D) average root diameter at the formative phase, and (E) length of the longest root, (F) root dry weight, (G) root volume, and (H) root-to-shoot ratio at the grand growth phase of Saccharum spp. clones recorded from phenotyping structure (S. officinarum: Awela Green Sport, S. spontaneum: IND 85-490, S. sinense: Oshima, S. barberi: Putli Khajee, S. robustum: NG 77-237). Data represent mean ± SEM (n = 3). Please click here to view a larger version of this figure.

Figure 7
Figure 7: Root anatomical traits of S. sinense clone Oshima. (A) Normal growing conditions and in response to (B) drought and (C) waterlogging stress. Scale bars = 100 µm. Abbreviations: rh = root hairs; ae = lysigenous aerenchyma; mx = metaxylem; ex = exodermis; ep = epidermis. Please click here to view a larger version of this figure.

Figure 8
Figure 8: Root exudation. (A) Carboxylates, (B) total phenol, (C) total protein, (D) total sugar, and (E) total free amino acids at the germination phase in Saccharum spp. clones (S. officinarum: Awela Green Sport, S. spontaneum: IND 85-490, S. sinense: Oshima, S. barberi: Putli Khajee, S. robustum: NG 77-221). Data represent mean ± SEM (n = 3). Abbreviation: RFW = root fresh weight. Please click here to view a larger version of this figure.

Figure 9
Figure 9: Rhizosphere traits in sugarcane varieties. (A) Triangle root cap in Co 62175, (B) dome-shaped root cap in Co 86032, (C) deep violet root tip pigmentation in Co 86032, (D) peroxidase activity, (E) superoxide dismutase activity, and (F) total phenolic content of plants raised in the hydroponic setup. Data in D, E, and F represent mean ± SEM (n = 3). This figure has been modified from Hari et al.9. Please click here to view a larger version of this figure.

Figure 10
Figure 10: Developmental phases of rejuvenation upon root injury in sugarcane varieties raised in the hydroponic setup. (A-E) CoC 671 and (F-I) Co 06022; (A,F) uninjured roots; (B,G) roots injured by longitudinal slicing; (C,H) 3 days after injury; (D,E,I) 10 days after injury. This figure has been modified from Hari et al.9. Please click here to view a larger version of this figure.

Cross sectional area of root (mm2) Ratio of cortex-to-stele Number of exodermal layers Number of metaxylem elements Average diameter of metaxylem (mm) Number of air spaces Area occupied by metaxylem (mm2) Area occupied by air spaces (mm2)
S. officinarum 6936.3±200.2 0.508±0.015 1.00±0.03 6.00±0.17 8.5±0.3 14.00±0.40 340.3±9.8 1993.0±57.5
S. spontaneum 27646.8±957.7 0.481±0.017 2.00±0.07 13.00±0.45 6.6±0.2 32.00±1.11 444.5±15.4 4921.3±170.5
S. sinense 7539.1±261.2 0.727±0.025 2.00±0.07 9.00±0.31 6.0±0.2 - 254.3±8.8 -
S. barberi 18859.6±435.6 0.692±0.015 1.00±0.02 7.00±0.16 12.0±0.3 12.00±0.28 791.3±18.3 4454.6±102.8
S. robustum 8328.1±288.5 0.563±0.019 2.00±0.07 10.00±0.35 7.0±0.2 - 384.7±13.3 -

Table 2: Root anatomical traits observed in Saccharum spp. clones. Data represent mean ± SEM (n = 3) (S. officinarum: Awela Green Sport, S. spontaneum: IND 85-490, S. sinense: Oshima, S. barberi: Putli Khajee, S. robustum: NG 77-237).

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Discussion

Root systems define the above-ground productivity of sugarcane, necessitating that all its facets be explored and understood thoroughly for the development of climate-resilient varieties. A team of scientists at ICAR-SBI comprising plant physiologists, a microbiologist, an agricultural engineer, a biochemist, and plant breeders employed multipronged approaches for sugarcane root phenotyping, including field excavation by trench sampling, the use of a root core sampler, raised platforms for root sampling, and raising plants under hydroponic culture. Given that an individual methodology may not be exhaustive and/or exclusive, the integration of information is required to reveal the component traits and mechanisms of an ideal root phenotype. Earlier methods of root sampling were isolated in their efforts but were useful for developing the strategy described herein.

Venkatraman and Thomas5 pioneered the study of sugarcane roots by describing three suitable methods for investigation. In the first method, a deep pit was dug at the side of a plant, which permitted the roots to be dissected from the soil, whitewashed, and photographed in their natural positions. In the second method, deep pits were dug on the opposite sides of a plant. Frameworks of 2 in wire netting were placed against the sides of the pits adjacent to the plant, with iron rods inserted through the breadth of the interposed soil to hold the roots in their respective positions. The soil was then washed with water, and the roots were photographed. A third method was to make special plantings for root study. Wire netting was placed at certain levels in an excavated plot, and the excavation was refilled, replacing the soil in its respective layers as before. The plants to be studied were grown in the plot. At the time of examination, the soil around the plants was washed with water, leaving the root systems suspended in position in the meshes of the wire netting.

Rege and Wagle20 developed a method of root study under irrigated conditions by digging a pit vertically down the edge of the cane and exposing the root system by washing the soil in a specified zone. It caused the least wastage of the plant material and enabled researchers to study the root system in situ both qualitatively and quantitatively. A special study was conducted to investigate the structure, development, and behavior of sugarcane roots21. "The root box method" and "the soil excavation method" proved quite useful in studying the distribution of sugarcane roots. It was found that there were more shallow roots in heavy soils than in mixed or lighter soils. The loose texture and aeration prevalent in the latter favored root penetration into deeper soil zones, making it more amenable for root development The first roots to develop grew directly downward in the soil, penetrating to depths of more than 30-36 in, followed by shallow-spreading roots observed in the surface layers (~14 in from the topsoil), wherein ~85%-90% of the roots are distributed. The deeper roots undoubtedly furnished moisture for the plants during dry periods. The rapidity of the growth of the roots and shoots varied according to the climatic conditions of the cropping year and with the variety. Maintaining roots in active physiological conditions is important for optimal plant growth, as it is often disturbed by cultivation practices, which must be avoided if possible. Root system variability in selected sugarcane varieties was studied in Cuba by the "direct examination method"22. Descriptions of the root development were documented together with illustrations of the root distribution in the soil.

The field excavation of roots by trench sampling was based on the work of Venkatraman and Thomas5, Rege and Wagle20, and Ryker and Edgerton21. Despite the fact that it is the most laborious and time-consuming method, in situ field sampling is imperative to understand root phenotypes in their natural growing environment. A comparison of root measurement methods in sugar beet and cereals revealed that data obtained from the trench profile wall must be interpreted with caution, wherein the calculated root length densities (RLD) (ratio of cumulative root length to soil volume) were 4x-10x lower than those obtained from the root core method, depending upon the soil type23. An appropriate and uniform distance from the trench to the stem is important to ensure minimal sampling error and minimize the loss of roots and resources, including labor. Monitoring root growth via trench sampling may not be feasible for long-term experiments, as it is destructive in nature, and only a small part of the root is visible based on the plane of observation. However, trench sampling provides an overall estimate of the root distribution and interaction with the soil profile24. This investigation on the spread of root systems in commercial sugarcane hybrids revealed that ~70% of physiologically active roots were found within 50 cm of the soil profile.

The trench sampling method allowed the differentiation of various root classes according to their depth of occurrence in the soil profile. In general, roots penetrating deeper into the soil were usually thinner than those found in the shallow zones. Wide genotypic variability was observed with respect to root length, volume, and weight. Advances in crop modeling and simulation aid in refining the data obtained from conventional sampling methods by developing algorithms to minimize sampling errors and experimental heterogeneity. Sugarcane RLD data from an in situ soil profile was compared with that of soil coring, wherein it was observed that counting root intersections in a soil profile and estimating RLD using a model gave RLD values similar to those obtained with the soil core method25. A comparison of data obtained from soil excavation and coring methods in soybean revealed similar estimates of root biomass, and excavation required less total time and more labor26. Core-based samples were more precise, allowing sampling in deeper rooting zones with minimal disturbance to the experimental plot. Given the advantage of studying plant roots in their natural growing environment, trench sampling may be useful for combining soil profile data and the RLD profile, but it does not facilitate sampling on an extensive area27. In this context, the root core sampler has proven quite useful for quantifying root distribution and allowing more samples on a greater area.

The root core sampler was designed to facilitate sub-sampling of roots from the field, which warrants uniformity in sampling with a lower coefficient of variance; it has considerably reduced sampling errors. The use of the root core sampler has been adopted in several crops; it requires a large amount of post-sampling processing time because much care needs to be given to avoid the loss of roots during washing28. The core sampler facilitated the sampling of roots in a fixed volume of soil and brought out the genotypic variation among the tested clones for root system traits, allowing more samples to be tested in a given timeframe. RLD, which is an important parameter to assess plant health, may be derived precisely with the help of a root core sampler. In general, systematic variation is attributed to coring location in terms of distance from the stem, which may result in significantly different root data. However, in the case of sugarcane, the sampler was fabricated based on the crop's unique morphology, and the primary shoot/cane is fastened to it during sampling. Hence, a uniform core of 16 cm diameter around the primary shoot/cane is ensured for all the varieties to minimize sampling error.

A significant correlation was observed between the root volume data recorded using the root scanner method and the volume displacement method29. As the sample size is much lower than that of the trench method, the variability among genotypes may be assessed using the root core sampler. Random variation due to phenotypic plasticity, soil heterogeneity, and other unknown factors affects the reliability of data obtained from a root core sampler, which may be overcome by repeated measurements and the use of simulation software to narrow down functionally relevant differences27. A virtual assessment of the variation among different soil coring strategies in maize, based on three-dimensional models of root system architecture and an area weighting algorithm30, revealed that such cost-efficient ways to obtain reliable RLD estimates may be the way forward for root studies.

Root phenotyping structures with adjacent compartments serve the purpose of simulating the field, and at the same time, maintain the homogeneity required for screening the inherent variability of sugarcane germplasm. The ease of destructive sampling with significantly less damage and wastage of roots than trench sampling facilitates the analysis of the entire root system at different phenophases. Customized root boxes or chambers based on crop morphology and root distribution have proven to be effective in other crops such as sorghum31, soybean, and buckwheat32. Similarly, the size and depth of compartments have been optimized to support the growth of sugarcane. Further, the side walls have been constructed using precast slabs, which can be easily lifted manually during root sampling. Root samples are taken by manually lifting the sidewall, followed by water jetting to expose the roots. The sidewalls are then replaced, and the soil is refilled, ensuring minimal damage to the adjacent plants. The variation in root traits in sugarcane and its relationship with shoot growth may be documented in plants raised in the root phenotyping structure. Excess irrigation in sugarcane produces a superficial root system, while water deficit encourages lateral spread with slight deep penetration33,34. With increasing nitrogen application, the root weight is reduced, while phosphatic fertilizers produce a favorable effect on root development35,36. Adjacent compartments in the structure allow the study of such responses of sugarcane to varying abiotic stresses and soil nutrient levels by manipulating the amount of irrigation water and using nutrient-depleted soil, respectively.

Hydroponic plant culture is the method of choice to evaluate nutrient stress responses in a large and diverse population of genotypes in soybean12, rice37, wheat38, and maize39. Simulated root studies in sugarcane date back to 192340, wherein a column of rings of earthenware with wire netting was employed to study the relative depth and plan of the roots. Hydroponic culture using earthenware cooking pots filled with dilute Knop's solution was also attempted for continuous monitoring of root development. An in-house hydroponic facility enables the study of the finer details of root development and changes in the rhizosphere biology9. The container size and planting density were optimized for sugarcane, wherein plants were grown for up to 10 months with minimal lodging. Rhizosphere characteristics, including root cap shape and root tip pigmentation, may be assessed only under hydroponic conditions, which facilitates the ease of sampling and the absence of soil adherents. The evaluation of developmental and stress responses at the root level may not be possible when plants are grown in the field, whereas hydroponic culture allows such measurements. It facilitates the collection and characterization of root exudates, which may not be possible under field conditions. Although the absence of physical barriers to root penetration is a major disadvantage of this method, the regular monitoring of root growth is an advantage when plants are raised under hydroponic conditions.

It is essential to redesign the architecture of new sugarcane cultivars based on a strong and ideal root phenotype that can meet the demand for sustainable agriculture. High-throughput root phenotyping platforms developed for rice and maize include components adaptable to a wide range of growth systems, such as soil and hydroponics, advanced root imaging techniques, and software tools41; upscaling such platforms for sugarcane offers much scope for improving productivity in the near future. Despite the emphasis on root biology as an important determinant of the growth and productivity of sugarcane and the availability of advanced root phenotyping techniques, few studies have reported on sugarcane root traits per se42,43,44,45. As roots are the most dynamic plant organs, a single methodology to evaluate variation in root phenotypes may not be conclusive, especially in a crop such as sugarcane. The application and integration of suitable permutations of methodologies in line with the objectives of study should enhance the understanding of sugarcane underground systems.

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Disclosures

All authors declare that there are no conflicts of interest.

Acknowledgments

The authors acknowledge the infrastructure and support extended by the Director, ICAR-Sugarcane Breeding Institute, Coimbatore, for establishing root phenotyping facilities for sugarcane. Funding provided by the Science and Engineering Research Board, Department of Science and Technology, Government of India, in the form of Early Career Research Award to KV (ECR/2017/000738), is duly acknowledged. The authors acknowledge Brindha, Karpagam, Rajesh, Sivaraj, and Amburose for their assistance in generating data in a meticulous manner.

Materials

Name Company Catalog Number Comments
Aeration pump with pipeline accessories Purchased from local sources NA Used for hydroponic culture of sugarcane
Boric acid Sisco Research Laboratories, India 80266 Preparation of modified Hoagland's solution
Calcium nitrate Central Drug House, India 27606 Preparation of modified Hoagland's solution
Composted coir pith Purchased from local sources NA Used for germinating sugarcane setts
Cupric sulphate Sisco Research Laboratories, India 38869 Preparation of modified Hoagland's solution
DEAE-cellulose Sisco Research Laboratories, India 10529 anion exchange resin for processing of root exudates
EDTA-ferric monosodium salt Sisco Research Laboratories, India 59389 Preparation of modified Hoagland's solution
Farm yard manure Purchased from local sources NA Used for germinating sugarcane setts
Glass tanks Fabricated in-house NA Used for hydroponic culture of sugarcane
HPLC Agilent Technologies 1200 Infinity Quantification of organic acids in root exudates
Magnesium sulphate Sisco Research Laboratories, India 29117 Preparation of modified Hoagland's solution
Manganese chloride Sisco Research Laboratories, India 75113 Preparation of modified Hoagland's solution
Molybdic acid Sisco Research Laboratories, India 49664 Preparation of modified Hoagland's solution
Potassium dihydrogen phosphate Central Drug House, India 29608 Preparation of modified Hoagland's solution
Potassium nitrate Central Drug House, India 29638 Preparation of modified Hoagland's solution
Protrays Fabricated in-house NA Used for germinating sugarcane setts
Red soil Purchased from local sources NA Used for germinating sugarcane setts
Root core sampler Fabricated in-house NA Used for in situ root sampling
Sand Purchased from local sources NA Used for germinating sugarcane setts
Seralite-120 Sisco Research Laboratories, India 14891 cation exchange resin for processing of root exudates
Supporting frame Purchased from local sources NA Used for hydroponic culture of sugarcane
Water motor pump Purchased from local sources NA Used for hydroponic culture of sugarcane
Whatman filter paper grade 1 Universal Scientific 1001090 Processing of root exudates
WinRhizo PRO (software) Regent Instruments Inc., Canada STD4800 Two-dimensional root scanner with software for analysis of roots
Zinc sulphate Sisco Research Laboratories, India 76455 Preparation of modified Hoagland's solution

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Tags

Multipronged Phenotyping Approaches Characterize Sugarcane Root Systems Water Conductors Nutrient Conductors Growth And Yield Stressful Environments Plant Roots In Situ Assessment Sampling Difficulties Genotypic Variation Sugarcane Root Traits Rhizosphere Characteristics Allelopathic Effects Microbial Symbiosis Above-ground Productivity Climate-resilient Varieties Variability In Root System Traits Sugarcane Root Phenotyping Field Excavation Trench Sampling Root Core Sampler Raised Platforms Hydroponic Culture
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Vengavasi, K., Raja, A. K.,More

Vengavasi, K., Raja, A. K., Srinivasavedantham, V., K., H., T., A., G. S., S., A., A. D., K., C., M., N., D., P. Multipronged Phenotyping Approaches to Characterize Sugarcane Root Systems. J. Vis. Exp. (186), e63596, doi:10.3791/63596 (2022).

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