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Chapter 8

Nitrogen Physiology of Sugarcane

Nicole Robinson, Jessica Vogt, Prakash Lakshmanan, andSusanne Schmidt

SUMMARY

Nitrogen (N), which accounts for ∼80% of soil-derived essential nutrients acquired by plants,is critical for sugarcane productivity. With thenotable exception of Brazil, high N fertilizer ratescharacterize producer countries aiming to maxi-mize yields; up to nine times more N is appliedthan removed by sugarcane crops in China andIndia. Sustainable production is now a focus ofsugarcane agronomy and breeding. Developingimproved nitrogen use efficiency (NUE), i.e., pro-duction per unit nitrogen, is targeted as one way tomaximize yields in low-input agronomic systems.Additionally, improved NUE may minimize thedetrimental effects of N as a pollutant in water andair under high input agronomic systems. NUEis underpinned by three aspects or levels of con-trol: N acquisition from the soil (external, eNUE),use of acquired N by the plant (internal, iNUE),including remobilization of N within the plant,and N obtained from diazotrophic microbes.

Transgenesis has become a potential way toimprove NUE. Advances have been made in graincrops through tissue-specific upregulation of theN assimilation enzymes glutamine synthetase andalanine aminotransferase. This potential is beingexplored in sugarcane.

Selection of new cultivars under “below-recommended” and “recommended” N suppliesis being used to identify N-efficient and N-responsive genotypes that will help characterizethe underlying traits contributing to NUE.

INTRODUCTION

Nitrogen (N) is quantitatively the most importantsoil-derived nutrient accounting for 0.3 to 2% ofplant dry matter throughout the sugarcane cropcycle. At high-N supply (> 200 kg N ha−1 yr−1)trial sites in Australia and Brazil, crops yielding60–150 t DW ha−1 accumulated between 150 and250 kg N ha−1 in the shoots and up to 380 kgN ha−1 in the total sugarcane biomass during thecrop cycle (i.e., one year of plant or ratoon cropgrowth) (Chapman et al. 1994; Muchow & Robert-son 1994; Wood et al. 1996; Basanta et al. 2003).However, reports of much lower N accumulation(∼66 kg N ha−1) in sugarcane from farms usingconventional practices in Australia yielding 100–150 t ha−1 fresh weight have raised doubts aboutsugarcane’s crop N requirements (Thorburn et al.2009; Bell et al. 2010).

Both the responses of the crop to N appli-cation and the NUE of the crop can be relatedto biomass production and sugar yield. Althoughhigher N application rates generally increase caneyield (fresh weight) and crop biomass (dry weight),they can reduce the commercial cane sugar (CCS)content of the expressed sugarcane juice and thesugar quality (defined as the amount of nonsucrosecompounds in the recovered sucrose) (Stevensonet al. 1992; Muchow et al. 1996). Current N fer-tilization strategies in most sugarcane producernations have resulted in high rates of N fertilizerapplication and unbalanced N-output-to-N-inputratios (Table 8.1); for example, in China 512 Gg

Sugarcane: Physiology, Biochemistry, and Functional Biology, First Edition. Edited by Paul H. Moore and Frederik C. Botha.

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170 Sugarcane: Physiology, Biochemistry, and Functional Biology

Table 8.1 Sugarcane production and N fertilizer application of the main sugarcane producing countries, accounting for 86and 88% of global sugarcane production area and cane yield, respectively.

Country Areaa (ha)Productiona

(Tg)N appliedb

(Gg)N harvestedc

(Gg)Output/input

ratio

Reported Napplication rates(kg N ha−1)

1. Brazil 8 141 135 649 613 389 0.6 60–100d

2. India 5 055 200 348 732 209 0.3 150–400e

3. China 1 708 520 125 512 75 0.1 100–755f

4. Thailand 1 054 439 74 44 44 1.05. Pakistan 1 241 300 64 231 38 0.2 120–180g

6. Mexico 669 231 51 70 30 0.47. Colombia 383 388 39 n/a 238. Australia 390 000 34 70 20 0.3 160h

9. Argentina 355 000 30 25 18 0.710. USA 374 200 28 40k 17 0.4 78–146i,

247–280j

11. Philippines 397 991 27 11 16 1.412. Indonesia 415 578 26 52 16 0.3 125g

13. Guatemala 287 000 25 n/a 1514. South Africa 425 000 21 57 12 0.2 60–200g

Total 20 897982 1 541 2 457 922 0.47l

Source: Table from Robinson et al. (2011).Note: An output/input ratio of 1 indicates that N removal with the harvested product matches N application (high NUE); anoutput/input ratio of 0.1 indicates that 90% of fertilizer N is not used by the crop (low NUE).aFAO area and yield data 2008.bHeffer 2009 International Fertilizer Industry Association Assessment of fertilizer use by sugar crops at the global level2007/2008.cCalculated based on a stalk dry matter content of 30% and N content of 0.2% dry weight.dHartemink (2008).eDr. T.K. Srivstava, Indian Council of Agricultural Research, India (personal communication 2010).fDr. Jiang Xiong Liao, Guanxi Sugarcane Research Institute, China (personal communication 2010).gFAO Fertilizer use by crop (2005).haverage application rate (Wood et al. 2010).iLouisiana State University Agricultural Center, Fertilizer Recommendations-2009.jR. Rice, Institute of Food and Agricultural Sciences, University of Florida (personal communication 2010).kFigures for USA calculated from average application rate and area.lWeighted average output/input ratio.

N are applied nationwide, whereas only 75 Gg Nare removed at harvest.

Declining or plateauing sugarcane yields inAustralia, coupled with the need to reduce off-sitepollution, have stimulated critical evaluations ofN fertilizer use (Schroeder et al. 2008; Wood et al.2010; Thorburn et al. 2011). Negative effects ofexcess N fertilizer applications include higher pro-duction costs and environmental degradation dueto soil N losses and soil acidification. Single appli-cations of readily dissolvable synthetic N fertiliz-ers result in high concentrations of soil inorganic Nearly in the growing season (Fig. 8.1). Nitrate andammonium not acquired by the crop can be lostfrom soil in three ways: through microbial con-

version into gaseous N, by leaching deeper intothe soil profile, or through runoff. Nitrogen lossesare exacerbated in the two to three months follow-ing fertilizer application by excess water from theonset of a wet season or by over irrigation (Fig. 8.1)(Allen et al. 2010).

Improvements of N relations in sugarcane pro-duction have focused primarily on agronomicmanagement of fertilization (Chapter 5). How-ever, understanding the physiology of N uptakeand metabolism may help develop alternativeagronomic practices to better align soil N relationswith crop production and germplasm improve-ment. Obvious issues are the timing of N availabil-ity and the forms of N available in the soil. While

Nitrogen Physiology of Sugarcane 171

Fig. 8.1. Nitrogen relations throughout a crop cycle in Australian sugarcane production. a, Soil nitrous oxide flux, soil mineralN (ammonium and nitrate), and third ratoon shoot N content fertilized with urea (200 kg N ha−1, arrow). b, Dissolved andc, soluble soil N of a plant crop fertilized with urea (110 kg N ha−1, arrow). In b, dissolved nitrate and ammonium werequantified in the soil solution (0–20 cm depth); in c, soluble nitrate and ammonium were quantified in soil KCl extracts (0–20cm) (mean and standard error n = 5 and n = 3, respectively). From Robinson et al. (2011).

nitrate has long been considered the main N sourcefor crops, partially due to its high mobility in soil,there is evidence that ammonium is an equallyimportant, or even preferred, N source for manyplant species including sugarcane (Robinson et al.2011). Because plants can use a wide range of Nsources, the existing focus on inorganic N sourcesmay not be totally justified. Amino acids and pep-tides are increasingly considered as N sourcesfor plant N requirements. In Australian sugar-cane soils, the concentrations of free amino acidsspanned one order of magnitude (0.22–2.42 mg

amino acid-N kg−1 soil) and accounted for up to70% of soluble N (sum of amino acids, ammo-nium, and nitrate) throughout the crop season(Fig. 8.2).

Sugarcane differs from grain crops by the pro-duction of sucrose-storing culms as the primaryharvest product rather than N-rich kernels. How-ever, sucrose yield and physiological processessuch as photosynthesis are strongly linked to cropN status. NUE has been investigated predomi-nantly in grain crops, focusing on the central roleof genetic variability for NUE explored through


Fig. 8.2. Soluble (KCl-extractable) low molecular weight N compounds in topsoil (0–20 cm) under sugarcane (% of soluble N,amino acids, ammonium, nitrate N). Soluble N concentrations are shown as numbers above bars. a, Redoxic hydrosol (secondratoon sugarcane, first crop on virgin soil, with 100 kg urea-N ha−1 applied Nov 2007) in subtropical Australia. b, Oxyaquichydrosol (plant crop, �50 year sugarcane cropping, with 130 and 150 kg urea-N ha−1 applied in Nov 2007 and Oct 2008,respectively) in tropical Australia (Holst et al. 2012).

whole-plant physiology, quantitative genetics, andforward and reverse molecular genetics (Hirelet al. 2007). Similar approaches are now emerg-ing in sugarcane, with emphasis on identifyingand quantifying the genetic variation for NUE(Schumann et al. 1998; Robinson et al. 2007; Whanet al. 2010).

Uptake and molecular regulation of the inor-ganic N forms nitrate and ammonium have beenstudied intensively in Arabidopsis (Ludewig et al.2007; Miller et al. 2007). In addition, some func-tional characterization of the transporter proteinsthat facilitate nitrate and ammonium uptake intoroots has been made in cereals (Quaggiotti et al.2002; Glass 2003). The focus of NUE researchin these species has been on molecular and bio-chemical traits linked to seed or grain produc-tion, including N remobilization for grain fill-

ing (Masclaux-Daubresse et al. 2010). This con-trasts with sugarcane, where the link between theregulation of N uptake and vegetative growth ismore relevant, particularly as N fertilizer appli-cations occur in the early stage of crop develop-ment. Hence, desirable traits for sugarcane includeearly maximum N capture and storage by thecrop and subsequent use of the stored N forbiomass production throughout the relatively longcrop cycle.

A further consideration is the inevitability ofN deficiency stress under water limitation; thephysiological implications of both drought andN stresses require an integrated physiologicalapproach to improving the use of N and waterby the crop (see Chapter 16). Here, we synthesizeknowledge of NUE in sugarcane with a focus oncrop physiology.


SETTING THE SCENE: NITROGEN IN THESUGARCANE CROP SYSTEM

Australian sugarcane agricultural systems are dis-cussed as representative of high-production sys-tems to illustrate how N inputs are linked to cropN physiology (Color Plate 8.1). Nitrogen fertil-izer is applied as granulated urea or ammoniumnitrate, generally as a single application early inthe approximately 12-month crop cycle at a rateof 120–200 kg N ha−1 y−1 (Wood et al. 2010).Fertilizer N as urea is rapidly converted by soilmicrobes to ammonium and nitrate, which arereadily available for plant uptake. Soil N is mostlycontained in the soil organic N pool (SON) com-prising insoluble, soluble, and dissolved organicpolymers and oligomers (proteins, peptides, other)and monomers (amino acids, other). The SONpool generally exceeds crop applied fertilizer N byat least one order of magnitude. SON accountedfor 1500 and 3700 kg N ha−1 to a 30 cm depthin subtropical and tropical Hydrosols (Holst et al.2012) but can reach 10 tons ha−1 in some soils (to1 m depth). Soil N is subject to interconversion bysoil biota and plants; the resultant N forms differin their role as crop N sources due to chemicalstructure, interaction with soil and biota, and theplants’ ability to acquire and metabolize differentforms of N (Color Plate 8.1).

Application of N fertilizer results in stronglyheterogeneous spatial and temporal availabilityof ammonium and nitrate (Meier et al. 2006a;Bell et al. 2010). In the three months follow-ing fertilizer application, concentrations in thesoil solution reached 3.5 mM ammonium and8 mM nitrate (Fig. 8.1), which is in the upperrange for agricultural soils (ammonium aver-age 0.78±1.5 mM; nitrate average 4.5±9.8 mM)(Wolt 1994).

Additional N is provided via crop residues,including ca. 40 to 100 kg N ha−1y−1 through“green cane harvesting”, under which the imma-ture tops of culms with attached green leaves arereturned to the soil surface as a trash blanket (ColorPlate 8.1) (Robertson & Thorburn 2000). Approx-imately 6–15% of trash blanket N is recoveredby the crop in the first year, and a large portionremains immobilized in the soil microbial biomass

(Ng Kee Kwong et al. 1987; Meier et al. 2006b).Possible N losses from the trash blanket includegaseous emissions, which remain to be quantified.Thorburn et al. (2000) investigated the mid- tolong-term effects of trash blanketing on soil Nand C stocks using five Australian field sites undergreen cane trash blanketing or burn treatments,with the duration of treatments ranging from 4 to18 years. At four of the five sites, trash blanketingincreased soil N in the top 2 cm of soil. However,the degree of increased soil N was disproportion-ate to the duration of treatment, suggesting thateffects of residue retention are strongly site specific(Thorburn et al. 2000). A 69-year empirical studyat a South African burnt versus trash-managedexperimental site also suggests that soil N stocksbuild up over the long term (van Antwerpen et al.2002).

One avenue considered for improving N acqui-sition from the soil would be to establish bet-ter synchrony between soil N supply and plantN demand by shifting from readily dissolvable,rapid-turnover, inorganic N fertilizers to slow-turnover, organic N-based fertilizers. The effectof organic N-based fertilizers has been exploredin temperate crop and tree production systemsin combination with crop genotypes that havean improved acquisition capacity for organicN (T. Nasholm, Swedish Agricultural Univer-sity personal communication). Australian growersincreasingly use organic amendments, includingmill and distillery wastes, as sources of N andother essential nutrients.

However, organic amendments vary in com-position (Chapter 5) and turnover characteris-tics and do not necessarily equate to a shift fromnitrate-dominated to organic N-dominated solu-ble N pools. For example, application of 150 tha−1 of compost (1–1.5% N) resulted in four tofive times higher soil nitrate concentrations thanN-fertilizer at the standard recommended appli-cation (120 kg N ha−1) (Calcino et al. 2009). InAustralian sugarcane systems, legumes with activebiological nitrogen fixation (BNF) are rotated withsugarcane in 3- to 4-year cycles during which thelegumes contribute 75–120 kg N ha−1 as residues,or up to 300 kg N ha−1 if the legume seeds are notharvested (Park et al. 2010).


Net removal of N through harvested sugarcanecrops accounts on average for 0.5–0.9 kg N t−1

fresh cane (Thorburn et al. 2011). This removalrate amounts to 50–90 kg N ha−1 y−1 for a nor-mal yielding crop (100 t cane ha−1 y−1). Nation-ally, annual N fertilizer applications (excluding Ninput from crop residues) show that 30% of N isrecovered in the harvested Australian sugarcanecrop (Table 8.1). Site-specific analysis indicates23 to 40% of applied N is recovered in sugarcanebiomass in one season (Kingston et al. 2008).

Loss of N occurs via pathways that are not wellquantified, but include

� ammonia volatilization (∼30–40% N loss inbroadcast and banded N-fertilizer applica-tion) (Freney et al. 1991),

� N2O emission (1.1–20% loss of applied N-fertilizer) (Weier 1998; Wang et al. 2008;Allen et al. 2010),

� surface run-off (< 10kg N ha−1 y−1) (Prove& Moody 1997), and

� leaching (3% loss of applied N-fertilizer)(Chapman et al. 1994).

In the case of N loss to leaching, it should benoted that such losses are difficult to quantify sothat modeling has been used to complement thelimitations of extensive monitoring. Nitrate lostto deep drainage was estimated at 1.5% of appliedfertilizer using the agricultural productivity modelAPSIM-SWIM (Stewart et al. 2006). However,the authors acknowledge this as a possible under-estimation of nitrate loss due to inaccurate tem-poral patterns of leaching and an inability of themodel to capture the effect of preferential flowpathways (Stewart et al. 2006).

Tracking 15N-labeled fertilizer in Brazil andMauritius suggests low N losses via leaching (NgKee Kwong & Deville 1984; Trivelin et al. 2002).However, leaching losses are site-specific andlarger at sites with higher rainfall and leaching-prone soils. Considerable leaching losses occurin north Australian sugarcane soils where nitrateconcentrations in deep soil (4–10 m depth) weremore than two orders of magnitude higher (0–72.5 mg nitrate-N kg−1 soil) than in adjacent rain-forest soil (0–0.31 mg nitrate-N kg−1 soil) (Rasiahet al. 2003).

Whole system N budgets exist for grain crops(Dobermann & Cassman 2002; Ju et al. 2009).However, N budgets of sugarcane systems arelacking details about the fate of N fertilizer, includ-ing how much N is incorporated into soil organicmatter and lost as N, which together prevent res-olution of eNUE.

MICROBIAL ASSOCIATIONS ANDSYMBIOSES FOR NITROGENACQUISITION

Brazilian sugarcane researchers have studied BNFsince the 1960s and have isolated diazotrophic bac-teria of the genera Gluconacetobacter, Azospiril-lum, Burkholderia, and Herbaspirillum from sugar-cane. These bacteria colonize intercellular spacesin shoot and root tissues and in the rhizosphere(James 2000 and references therein; Fischer et al.2011). Gluconacetobacter diazotrophicus, originallydiscovered in Brazilian sugarcane, occurs in sugar-cane systems of the Philippines, Argentina, Cuba,Mexico, Mauritius, India, and Australia. How-ever, the role of G. diazotrophicus as a functionalendosymbiont has not been universally confirmed(Saravanan et al. 2008).

Despite evidence for sugarcane-diazotrophicbacterial associations (Urquiaga et al. 1992), thecontribution of BNF to the plant N budget is dif-ficult to ascertain. It remains to be substantiatedthat diazotrophic endophytes actually deliver Nto plant cells because it is difficult to differen-tiate BNF from the soil N pool versus crop-Nremoval budgets due to the large SON pool, soilheterogeneity, and N inputs from biomass burn-ing and aerial deposition. Surrogate techniquesfor quantifying BNF include acetylene reduction,15N-tracer dilution, and measurement of 15N nat-ural abundance. While each of these techniqueshas been used on sugarcane, all have been crit-icized for their inaccuracy (Baldani et al. 2002).Thus, the contribution of diazotrophs to the over-all crop N budget has yet to be quantified (James2000).

Beneficial effects on plant growth have beenobserved with plant growth-promoting rhizobac-teria (PGPR), which augment plant performance


through enhanced resilience to abiotic and bioticstresses, root vigor, and nutrient uptake (Yanget al. 2009). These multiple factors confoundinterpretation of PGRP-improved crop growthbecause the growth responses may not beattributable solely to BNF. While BNF contri-butions of 25 to 60% of the crop N budgethave been proposed in Brazilian sugarcane sys-tems (Boddey et al. 2001), there was no evidencethat BNF contributes significant quantities of N inSouth African sugarcane systems (Hoefsloot et al.2005). This was true even though diazotrophicendophytes were detected in the tested sugarcanetissues and nitrogenase encoding nif genes wereexpressed. An initial assessment of Australian sug-arcane systems suggested that there was no notableN input via BNF (Biggs et al. 2002).

Symbiotic BNF is restricted to sugarcane sys-tems receiving low N fertilizer input as high soilconcentrations of nitrate and ammonium are asso-ciated with reduced bacterial infection and toxi-city to G. diazotrophicus strains (Saravanan et al.2008). In addition to these environmental con-straints, there is evidence that diazotrophic bac-teria have cultivar-specific relationships (Baldaniet al. 2002; Nogueira et al. 2005).

Inoculation of sugarcane with diazotrophic andother beneficial bacteria is gaining popularity(Govindarajan et al. 2006). Whether input fromBNF can be derived from endophytes, from rhi-zosphere bacteria, and/or from legume compan-ion crops remains to be established and may differbetween production systems. BNF is currently nota focus in all producer countries because syntheticN fertilizers can be used to increase yields suffi-ciently for a positive return on costs. The advent ofmetagenomics and metatranscriptomics for char-acterization of microbial communities (Hugen-holtz & Tyson 2008), combined with knowledgeof soil biological processes, can be expected toadvance BNF in sugarcane crop systems.

While BNF symbioses have been a researchfocus, mycorrhizal symbioses have received rel-atively little attention in sugarcane systems. Themajority of terrestrial plants form symbioses withmycorrhizal fungi that extend the root system,absorb nutrients from plant-distant soil, and exudechemicals and enzymes to assist organic mat-

ter decomposition. Arbuscular mycorrhizal fungi(AMF) form symbioses with most herbaceousspecies, including sugarcane. AMF facilitate theplant’s access to nutrients, especially phospho-rus and N via an effective hyphal network thatis, in terms of the plant’s carbon economy, 100times cheaper to construct than are foraging roots(Rooney et al. 2009).

AMF account for at least 5–10% of soil micro-bial biomass. This abundance highlights the possi-ble importance of such symbiosis for crop systemfunction; nevertheless, many fundamental ques-tions about AMF remain unanswered (Fitter et al.2011). Nitrate leaching was reduced 40-fold inN-pulsed pots containing arbuscular mycorrhizal(AM)-colonized tomato plants compared to non-mycorrhizal plants (Ashgari & Cavagnaro 2012).This increased ability of mycorrhizal plants to cap-ture nitrate could also be important in sugarcaneproduction systems. Plowing physically damageshyphal networks such that one benefit of reducedtillage includes improved supply of the crop’srequired N and P via mycorrhizal symbionts (Fit-ter et al. 2011).

Most research of AMF and N relations hasevaluated assimilation of ammonium and nitrate,although it is not clear how much and which formsof N are delivered to roots via mycorrhizal sym-bionts (Smith & Smith 2011). In addition, there isevidence that AMF decompose organic materialand retrieve organic N for the host plant (Hodge& Fitter 2010).

Among the improvements in performance ofAM-colonized plants compared with nonmycor-rhizal controls are higher rates of BNF in legumes,which has been attributed to better P nutrition(Smith & Smith 2011). The plant cost of AMFsymbiosis is estimated at 4–20% of photosyn-thetically fixed carbon, depending on plant geno-type and soil available nutrients. Plants achieve acost-benefit balance by modulating their invest-ment into AMF symbiosis. Generally, plants par-tition less fixed carbon and energy to AMF whennutrient availability is increased (Sawers et al.2008; Rooney et al. 2009). Modern crop breed-ing may have inadvertently selected against AMFsymbioses by selecting current cultivars underhigh fertilizer input conditions. For example, it


has been shown that older wheat varieties, whichwere selected under lower fertilizer conditions,are more responsive to AMF colonization thanare modern varieties (Sawers et al. 2008). Thisphenomenon has not been studied in sugarcane.

Increased AM-colonization of field-grown sug-arcane in South Africa increased the crop leaf Nand Ca contents (Jamal et al. 2004). Similar pos-itive effects were noted in Indian sugarcane colo-nized by AMF (Prasad & Bilgrami 2002). Growthpromotion was observed with four compatibleAMF species; at 80 days after inoculation, theAMF-inoculated sugarcane biomass was 4- to 5.8-fold greater than uninoculated controls (Reddyet al. 2004). In a survey of southern Indian sitesacross different soil types and sugarcane geno-types, 34 to 84% of the root system was colonizedby AMF species (Srikumar et al. 2009), supportingthe notion that AMF are important in sugarcaneproduction systems.

In Australia, the AMF species Glomus clarumwas examined in soils associated with yield declinebut was not confirmed as an agent responsible forthe negative trend (Kelly et al. 2001). Phosphorusnutrition of AMF-inoculated and noninoculatedsugarcane seedlings demonstrated that G. clarumsymbiosis enhanced plant growth and P nutri-tion at low P supply, while a lower colonizationof AMF at higher P supply was consistent withthe plants’ regulatory control over the cost-benefitof AMF symbiosis (Magarey et al. 2005). Indeed,Kiers et al. (2011) confirmed that AM symbiosisconsists of a fair, two-way transfer of resourcesin which plants reward the most beneficial fungito obtain optimum nutrient-transfer. Expandingknowledge of AMF and the role of AM symbiosishave potential to increase interception and storageof fertilizer N supplied to the crop.

NITROGEN AND SUGARCANEPRODUCTIVITY

Nitrogen use efficiency

Nitrogen use efficiency describes relationshipsbetween N supply and biomass production or har-vested yield. NUE is often separated into exter-

nal efficiency (eNUE, denoting the efficiency ofabsorption or uptake of N from soil) and inter-nal or utilization efficiency (iNUE, the measure ofhow well the acquired N is used for production ofbiomass or seed) (Good et al. 2004). Distinctionbetween the two components of NUE is importantwhen determining the physiological and biochem-ical processes controlling NUE.

Much initial research on NUE focused on pre-dicting crop N uptake as affected by climate,management, crop age, and phenology and deter-mining the crop N demand, including inter-nal requirements for maximum yield production.Biomass-to-N ratios (iNUE) of sugarcane tissuesvaried from 160 to 540 kg DW kg−1 N over a rangeof biomass from 26 to 150 t DW ha−1 (Muchow &Robertson 1994). With increasing N application,increases in N uptake by the crop were greaterthan was the accumulation of biomass, indicatingthat sugarcane has a capacity for luxury N uptake.Muchow et al. (1996) suggest a yield threshold of400 kg DW kg−1 N, with lower and higher ratiosindicative of oversupplied and N-limited crops,respectively. However, these ratios are based ononly few studies with few varieties and high Ninputs (Stanford & Ayres 1964; Muchow et al.1996). These limited studies may not reflect Nrelations of sugarcane genotypes globally. Indeed,a plant crop in Brazil increased biomass by 50% inresponse to 120 kg N ha−1, although both fertil-ized and unfertilized crops had identical iNUE of< 400 kg DW kg−1 N (Franco et al. 2011). Addi-tional studies of diverse genotypes and manage-ment systems are needed to improve our under-standing of the relationship between crop biomassand N content.

Genetic variation in iNUE

Sugarcane has not been extensively studied forgenetic variation of NUE, whereas grain cropsincluding maize (Hirel et al. 2001, 2007), rice(Obara et al. 2004), and wheat (Foulkes et al. 2009)have received considerable attention. Knowledgeof NUE in grain crops has steadily progressed;molecular mapping and screening for physiologi-cal parameters of NUE have examined genotypes


in both controlled and field conditions (Coque &Gallais 2007).

Sugarcane genetic variation of NUE was stud-ied in Australia on a biparental population of 61progeny genotypes. The male parent of this fam-ily was an Australian commercial cultivar derivedfrom several generations of breeding and selec-tion for high sugar yield under high N supply;the female parent was an unselected S. offici-narum accession. Screening of the 61 genotypesin glasshouse conditions at limiting (0.4 mM N)and high (10 mM N) N supply demonstrated thatbiomass production after three months’ growthvaried ninefold between genotypes under low Nand fourfold under high N supply (Robinson et al.2007). iNUE was on average twofold greater atlow than high N supply and genotypes varied upto twofold at both low (143 to 303 g DW g−1 N)and high (53 to 110 g DW g−1 N) N supply. TheiNUE of genotypes exhibiting low iNUE (∼150g DW g−1 N) at low N supply ranged widely athigh N supply, while genotypes with high iNUEat low N supply (∼300 g DW g−1 N) also hadhigh iNUE at high N supply (Fig. 8.3; Robin-son et al. 2007). These findings suggest that highiNUE occurs across N supply regimes and thatcultivation of genotypes under N-limiting condi-tions may prove effective for selecting genotypeswith high iNUE.

The activity of N assimilation enzymes glu-tamine synthetase (GS) and nitrate reductase(NR) has been implicated in NUE of maize, show-ing positive and negative correlations, respec-tively, with yield (Hirel et al. 2001). However,GS was not a determinant of iNUE in young sug-arcane plants (Robinson et al. 2007), and the roleof NR in sugarcane NUE has not been investi-gated systematically in relation to genotypic NUE.Quantitative trait loci (QTL) analysis of iNUEin sugarcane was performed in glasshouse exper-iments with the same mapping population thatwas used for the iNUE study. Significant marker-trait associations (MTAs) accounted for 3–12%of phenotypic variation in the male parent and 3–19% of phenotypic variation in the female parent.MTAs were detected for biomass, shoot N contentand iNUE, leaf protein content, and GS activity(Whan et al. 2010).

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Fig. 8.3. Impact of N supply on (a) biomass accumulation (gdw) and (b) iNUE (g dw g−1N) for studied genotypes. Sixty-one genotypes were grown for 12 weeks with low (0.4 mM)or high (10 mM) N. Data are means of three to five plants.Q165 is represented by solid symbol (5), siblings by opensymbols, KQ99-1302 (1), KQ99-1345 (2), KQ99-1367 (3),KQ99-1447 (4), KQ99-1454 (6), KQ99-1459 (7), KQ99-1286(8), KQ99-1308 (9) and KQ99-1391 (10). Population averagebiomass (3.7, 28.4 g dw) and iNUE (190, 90 g dw g-1 N) atlow and high N supply, respectively, are indicated by linesin each panel. Confidence intervals (CI) of 95% for averagebiomass and iNUE for genotypes at low and high N sup-ply shown by bars. Reproduced from Robinson et al. (2007),http://www.publish.csiro.au/nid/102/paper/FP07183.htm.

The significant MTAs identified for iNUEmay reflect different selection pressures on theparental lines because the male parent was a cul-tivar selected under high N and the female par-ent was an accession that evolved under low N.


No positive MTAs were detected under high Nsupply for iNUE among markers from the malecultivar, whereas six of ten MTAs detected inthe female S. officinarum accession were positive.At low N supply, 13 MTAs, five of which werepositive for iNUE, were detected in the parentalfemale accession compared with four of sevenmarkers that were positive in the parental malecultivar (Whan et al. 2010). Markers with thegreatest effect were inherited from the S. offici-narum accession. Variation for iNUE under highN may be lower in commercial varieties than inunimproved genotypes, indicating that greatestgains for improving iNUE could occur by intro-gression of unselected germplasm. These findingshighlight the importance of screening genotypes atlower N supply to characterize variation in NUE(Whan et al. 2010).

Experiments in South Africa compared NUE,based on biomass and sucrose yield, of seven culti-vars grown over a period of >12 months in outdoorsand cultures supplied with a continuous nutrientbath at three N rates (2.1 to 6.4 mM N) (Schu-mann et al. 1998). The two cultivars with thehighest biomass and iNUE (40–50 g sugar g−1

N), at medium N supply, had an average iNUE(18 g sugar g−1 N) at the high N supply. In con-trast, the least efficient cultivar increased biomassmore than twofold from medium to high N sup-ply, but had lower iNUE (Schumann et al. 1998).Based on subsequent field trials, fertilizer recom-mendations for the least efficient cultivar wereincreased by 30 kg N ha−1, while a reduction of20 kg N ha−1 was recommended for the most effi-cient cultivar (Meyer et al. 2007). These exam-ples illustrate that screening of genotypes in con-trolled conditions is a useful approach to identifygenetic variation for iNUE and to inform N fertil-izer application. Discerning the traits conveyingN efficiency will allow more targeted screeningand breeding for optimum production under fieldconditions.

Field screening of genotypes for NUE

Field trials in South Africa have indicated dif-fering genotypic responses to changing N sup-ply (Inman-Bamber 1984; Meyer et al. 2007),

although the traits conferring NUE remain uncer-tain. Further evaluation of the genotypes would berequired to establish the significance of the geno-type and N supply interaction.

A comparison of four sugarcane varieties grow-ing on sandy soils in Florida showed that theone cultivar that had been selected on sandy, N-deficient soils, produced 1.5- to 2.6-fold greateryields across a wide range of N supply rates (0to 896 kg N ha−1) than did the set of threecultivars that had been selected on N-rich soils(Gascho et al. 1986). However, results did notallow determination of the cultivar by N inter-action, which could be determined only in con-trolled hydroponic culture. The cultivar selectedon N-deficient soils had the highest iNUE produc-ing more biomass while having the N content inculms similar to that of the other cultivars. Thus,it is possible that several NUE traits contributedto the enhanced performance of this cultivar in thefield trials (Gascho et al. 1986).

Wood et al. (1996) compared growth of twoleading Australian cultivars that received high Nfertilizer application and additional side dressingto alleviate potential N limitation throughout thegrowing season. Both cultivars produced similarlevels of biomass (Robertson et al. 1996). How-ever, one cultivar accumulated more N than didthe other, which had a greater ability to diluteN throughout the crop cycle. Thus, maximum Naccumulation was unrelated to maximum biomassproduction. However, maximum N accumulationwas greater in the plant than ratoon crops ofboth cultivars (Wood et al. 1996). The questionsof whether the cessation of N accumulation bythe crop was due to depletion of soil N sources,reduced N uptake activity of roots, lower crop Nrequirement, or some combination of these factorswere not resolved. The difficulty of distinguishingamong these factors remains an ongoing problemwith field screening of genotypes.

N allocation to aboveground plant tissues

To ensure maximal crop productivity, leaf N lev-els must be maintained at concentrations that donot limit radiation use efficiency (see Chapter 6).Early in crop growth, leaves constitute 40% of the


biomass (∼10 t DW ha−1) so that leaf N accountsfor most of the plant N pool. Later in crop devel-opment, leaves constitute 10% of plant biomass,which is approximately 50 to 70 t DW ha−1 atharvest (Robertson et al. 1996). In addition, leafarea index (LAI) increases in the first five to sixmonths of crop growth and decreases thereafterwith the senescence of leaves near the culm base.New leaves are produced throughout the cropcycle, but the N content of successive green leavesdeclines with crop age and potentially decreasestotal photosynthesis of the leaf canopy (Allisonet al. 1997). Culm N biomass becomes far greaterlater in crop development and in South Africaand Australia, applications of additional N sup-ply over the growing season resulted in increasedculm N, rather than increased foliar N content orleaf biomass (Stevenson et al. 1992; Catchpoole &Keating 1995).

Variation in canopy N due to uptake and par-titioning of N has been assessed in different cul-tivars and different crops. Cultivars could be dis-tinguished by different levels of N accumulated,and the plant crop accumulated more N than didthe ratoon crop (Wood et al. 1996). Differencesin foliar N content among plant and ratoon cropscould be explained by differences in total N accu-mulation for a given biomass. Although total Naccumulation differed across crops, the plant andratoon crops had a similar proportion of green leafbiomass and total amount of plant N containedin the green leaves (Robertson et al. 1996; Woodet al. 1996).

Whether N allocation to different tissues is asignificant trait for enhancing NUE was investi-gated in field plots involving five genotypes of anunselected mapping population contrasted witha single common cultivar in the second ratoonat 0 and 100 kg N ha−1 supply (Robinson et al.2008, 2009). Despite more than twofold differ-ences in biomass accumulation within and betweenN treatments, whole culm biomass and trash atharvest contained on average 0.27% N across allgenotypes and N supply rates (Fig. 8.4). The geno-type that accumulated most biomass at low N rel-ative to high N supply had <40% of N allocatedto culms at both N supply rates, which was sig-nificantly less (P < 0.05) than in all other geno-

types (except for one where the reduction wasnot statistically significant) (Robinson et al. 2009).This result suggests that greater N allocation toleaves contributes to better growth at low N sup-ply. Nitrogen allocation to different tissues in con-text of NUE is being investigated in the field over awider range of genotypes (Robinson et al. unpub-lished observations).

Leaf N and photosynthesis

Leaf N concentration and photosynthetic capac-ity are understandably correlated since N is amain component of chlorophyll and the CO2-assimilating enzymes phosphoenolpyruvate (PEP)carboxylase and ribulose bisphosphate carboxy-lase/oxygenase (Rubisco). The importance of N inthese fractions is seen in the relationship betweenleaf area and adequacy of N supply. Plants reduceleaf area, leaf N concentration, or both in responseto N deficiency, which in turn decreases radiationinterception and radiation use efficiency. There isevidence that grasses maintain their leaf area butdecrease leaf N concentration in response to low Nsupply (Sinclair & Horie 1989; Pilbeam 2011). Adecrease in specific leaf N content (SLN) from 1.7to 0.8 g N m−2 leaf area resulted in a linear declinein photosynthetic rate in a single cultivar studyin South Africa (Allison et al. 1997). Similarly,a different cultivar studied in Australia showed adecline in SLN, from 2 to 0.8 g N m−2, accompa-nied by a near linear decline in its net assimilationrate from 40 down to 8 μmol m−2 s−1 (Ludlowet al. 1991).

Interestingly, the photosynthesis-limitingthresholds of SLN differ among C4 crops. Inmaize and sorghum, maximum net assimilationrates of 45–50 μmol m–2 s−1 were achievedat SLN ∼1.0–1.5 g N m−2 and declined onlywhen the SLN was below this threshold level(Muchow & Sinclair 1994). In sugarcane the SLNthreshold is considered to be higher at 1.7–2.0 gN m−2 (Ludlow et al. 1991) (see Chapter 6 for adiscussion on the relationship between leaf N andassimilation).

Sugarcane grown under a standard crop N fer-tilizer rate displayed a decline in SLN from theupper to lower canopy leaves and throughout the


0.0

1387

1317

1355

Q117

1484

1409

1387

1317

1355

Q117

1484

1409

0.5

N A

lloca

tion

Bio

mas

s A

lloca

tion

1.0

0.0

0.5

1.0

0.0

0.5

1.0

0.0

0.5

1.0

0 kg N ha–1 100 kg N ha–1

Leaf

Cabbage

Stalk

Trash

Fig. 8.4. Proportion of biomass and nitrogen allocated to leaf, tops (shoot top including leaf sheath, but no lamina), stalk,and trash (dead leaves) in individual stalks sampled prior to final harvest in September 2008 after 12 months growth withoutfertilizer or with 100 kg N ha−1. Values are averages of six individual stalks with bars indicating standard error. Genotypesinclude five unselected clones (KQ99-clones mapping population) and Australian commercial variety Q117.

crop season. SLN ranged from 1.7 to 1.0 g N m−2

in the upper and lower canopy at week 15 andfrom 1.2 to 0.8 g N m−2 at week 45. This seasonaldecline was not prevented by an additional 100kg N ha−1 top dressing of N fertilizer applied at15, 25, and 35 weeks after planting (Allison et al.1997). In more than 50% of high N-input (>160kg N ha−1) systems studied in Australia, the SLNof canopy leaves was <1.2 g N m−2 at 4000 g m−2

biomass, equating to approximately 40% of thefinal yield (Park et al. 2005). This result indicatesthat photosynthesis was limited by low SLN dur-ing much of the crop life cycle, irrespective of Nfertilizer supply.

Suboptimal SLN along with negative feedbackof high sucrose concentrations on photosynthesishave been proposed as causes for reduced growthin the second half of the crop cycle (Park et al.2005; van Heerden et al. 2010, and see Chap-ter 18). Developing cultivars with both enhancedability to acquire N later in the cropping cycleand improved allocation of N to canopy leaves to

avoid N limitation of photosynthesis is thereforeimportant for maintaining high assimilation andgrowth rates throughout the crop cycle. Similarly,lowering the photosynthesis-limiting threshold ofSLN should be an avenue to improve growth andNUE (see Chapter 6).

Glasshouse-grown Hawaiian cultivars differedin NUE (biomass produced per unit leaf N) andPNUE (photosynthetic NUE defined as the CO2

fixation rate per unit leaf N) (Ranjith et al. 1995;Ranjith & Meinzer 1997). NUE was positivelycorrelated with allocation of N to foliar chloro-phyll and Rubisco. These findings suggest thatselecting genotypes with greater investment intoessential components of the photosynthesis appa-ratus under lower N supply may be a promisingapproach for improving NUE.

Root systems

Plants modulate root growth in response to thesupply and form of N (e.g., Reynolds & D’Antonio


1996). Root phenology, especially vigorous rootgrowth early in the growing season when N avail-ability in fertilized soils is high, was important forefficient N acquisition by wheat (Liao et al. 2006).Few studies have evaluated genotypic variation ofsugarcane root systems, and none have integratedroot traits from overall morphology to molecu-lar control of N uptake with a view of improvingNUE. Smith et al. (2005) concluded that the lackof information on root traits of sugarcane cultivarshas prevented targeted selection for root vigor andenhanced efficiency for acquisition of soil-derivedresources.

Sugarcane root length densities in the uppersoil profile vary considerably, ranging from 1.3to 5.3 cm/cm−3 (root length per volume soil),but the effects of soil properties, crop develop-ment, and cultivars on root growth are poorlyunderstood (Smith et al. 2005). Root distribu-tion in response to N supply was investigated in amedium-textured soil in Brazil (Otto et al. 2009).The absence of fertilizer resulted in a more evendistribution of roots throughout the profile in acultivar having 50% of its roots occurring between20-60 cm depth compared to the fertilized plotswhere only 30% of root occurred below this depth.Evensen et al. (1997) determined biomass alloca-tion of two Hawaiian cultivars over a 2-year growthcycle and showed that root growth patterns weresimilar in both cultivars. Maximal rooting depth ofboth cultivars was 1.3 m and the majority of rootsoccurred in the upper 50 cm. However, use of afertigation drip system likely reduced deeper rootgrowth by providing a constant supply of waterand nutrients on the surface, which alleviated theneed for plant partitioning of resources to rootgrowth to seek water and nutrients (Evensen et al.1997).

Genotypic differences in root growth havebeen identified in controlled conditions in soiland hydroculture. In hydroculture, shoot growthincreased four- to sixfold in response to threefoldincreased N supply. In contrast, root growth wasunaltered in one cultivar, but increased two- andthreefold in two other cultivars (Schumann et al.1998), demonstrating that the root response candiffer from shoot response to N supply.

The impressive size of sugarcane root systemswas evidenced in Mauritius-grown crops in the1930s that extended to 5 m depth and 2.4 m width(Evans 1935). More recently, rooting depths of4.2 and 4.7 m were observed in Brazil for a cul-tivar grown in irrigated and rainfed, well-drainedferrasol soil having a clay content ranging from36 to 59% (Laclau & Laclau 2009). Greater rootlength density in the upper soil increased N uptakein wheat (Liao et al. 2006), and deep roots to2.5 m enabled extraction of water and nutrientsfrom the deeper soil in fodder radish (Kristensen& Thorup-Kristensen 2004). There is generallylittle recognition that deeper roots do play a rolein nutrient acquisition in sugarcane, so much sothat deep roots beyond 2.25 m are not consideredin sugarcane crop modeling (Stewart et al. 2006).However, deep roots could be important for nutri-ent acquisition by sugarcane.

In a pilot field study in Australia with 0 or 100 kgN-fertilizer ha−1, root growth of one cultivar andtwo unselected genotypes was assessed at the endof the growing season (Robinson et al. 2008). Inboth N treatments, the cultivar had more vigorousroot and shoot growth than did either of the uns-elected genotypes. Relative root growth and thedistribution of roots in the soil profile in responseto N supply also differed between genotypes. With0 kg N, the cultivar and the more sensitive of theunselected genotypes reduced root growth by ca.50% compared with N-fertilized plants, while theother unselected genotype had only a 30% reduc-tion (Fig. 8.5). Under 0 N, this less sensitive geno-type preferentially partitioned root production tothe deeper profile. In both N treatments, the sen-sitive genotype had >90% of root mass in the top40 cm, while the less sensitive genotype and thecultivar produced 60 and 83% of their roots in thetop 40 cm.

Glasshouse experiments in Hawaii showed thata drought-resistant cultivar invested more biomassto roots at low N supply than did a more drought-susceptible cultivar. This partitioning of biomassresulted in a root:shoot ratio of 0.48 for the resis-tant cultivar compared to a ratio of 0.38 forthe susceptible (Ranjith 2006). Rooting differ-ences among commercial cultivars obviously exist;


Fig. 8.5. Root distribution (number of roots per 100 cm2)scored on a trench face to 1 m depth for variety Q117 andunselected genotypes KQ99-1409 and KQ99-1484 at harvestof the second ratoon grown without (-N) or with 100 kg Nha−1 (+N). Values are averages for a 3 m row within a fieldexperiment (Robinson et al. 2008).

however, whether cultivars selected for root vigorcan enhance eNUE remains to be established.

Storage of N in belowground tissues

There is an increasing focus on the role of thebelow ground organs in developing cultivars withimproved resource use efficiency (Matsuoka &Garcia 2011). Efficient N remobilization fromshoots to below ground tissues prior to har-vest (“remobilization NUE”, often considereda component of iNUE) reduces the amountof N removed with biomass and can provideN for subsequent ratoon crops. Below groundbiomass, including stubble, rhizomatous mate-rial, and coarse roots excavated to 1.5 m depth,accounted for 15 to 60% of above ground biomassand 20 to 41% of plant N (2.4 to 4.7 mg N gDW−1) in the second ratoon crop of a cultivarstudied under two levels of N (0 or 100 kg N-fertilizer ha−1). Unfertilized plants allocated 67%and fertilized plants allocated 72% of N to shoots,suggesting that cultivars allocate ca. 30% of plantN to below ground tissues irrespective of N supply(Table 8.2) (Robinson et al. 2009).

Remobilization of N is desirable for improv-ing sugar yield and quality, but whether N remo-bilization from the culm exists in sugarcane hasnot been explored. Remobilization of N from theshoot of the related temperate giant grass Miscant-hus x giganteus is reported to have transported upto 280 kg N ha−1 into rhizomes (Amougou et al.2011) and supported successive high yields withlittle or no N fertilizer input (Beale & Long 1997).However, perennial growth habit, variable har-vest times, and a continuous growth climate posea challenge for realizing efficient N remobilizationin tropical sugarcane.

NITROGEN ASSIMILATION ANDAGRONOMIC GAINS

Crop N uptake, particularly the acquisition andassimilation of nitrate and ammonium and thetransport and remobilization of N, has receivedconsiderable attention in grain crops and themodel plant Arabidopsis (Masclaux-Daubresse


Table 8.2 Biomass and N allocation of the second ratoon of Q117 in southeastern Australia.

Q117 2nd ratoonYielda (t freshcane ha−1)

Totalplant N(kg Nha−1)

Aboveground N(stalks and leaves

kg N ha−1)% of

total N

Belowground N(stubble, rhizomeand coarse roots

kg N ha1)% of

total N

Unfertilized 49 39 26 67 13 33Fertilized (100 kg

N ha−1)109 103 74 72 29 28

Note: Data were calculated from averages of three individual stools (Robinson et al. 2009).aYield includes leaves, tops, and stalk but not belowground material.

et al. 2010; Lea & Miflin 2011). However, lessis known about these aspects of N physiology insugarcane. Understandably, the physiology andagronomy of N in sugarcane differ from the better-studied species in part at least because sugarcane isa tropical perennial species that produces a carbon-based yield. Nevertheless, even in those speciesthat have been well-characterized for their N phys-iology, agronomic gains have rarely been directlylinked to crop N physiological traits. Identificationof molecular markers for complex N physiologytraits is beginning to bridge this gap (Hirel et al.2007).

Nitrogen source preferences of sugarcane

There is general agreement that many plantspecies use nitrate as their main source of N dueto the high concentrations and mobility of nitratein agricultural soils and plants’ ability to take up,store, and metabolize nitrate. However, we do nothave a very good understanding of plant N usesources because the complex turnover processesin the soil-microbe-root interface produce organicforms of N (e.g., amino acids, small peptides) thatmay be more important than has generally beenconsidered (Nasholm et al. 2009). Our under-standing of plant N source is still largely deducedfrom the presence of soluble inorganic N in the soiland experiments in controlled hydroponic culturethat show plants grow best in response to a mixtureof nitrate and ammonium (Raven et al. 1992).

One Saccharum officinarum accession grewequally well in hydroponic culture with eithernitrate or ammonium as its sole N source (deArmas et al. 1992). On the other hand, commer-

cial hybrid cultivars, accessions of wild S. spon-taneum, and other S. officinarums discriminatedagainst nitrate when well supplied with equimolarconcentrations of nitrate and ammonium (Robin-son et al. 2011). This behavior of Saccharum spp.contrasts with that of related Poaceae. Sorghumand maize took up similar amounts of nitrate andammonium; Erianthus took up 80% the amount ofnitrate compared to ammonium (Robinson et al.2011). Possible reasons for the differences amongthese species include a lower ability of sugar-cane hybrids and sugarcane’s progenitor speciesto acquire, transport, and store nitrate.

The ability of plant species to use nitrate didnot appear to be directly linked with NR activitiesof leaves. A sugarcane cultivar had an in vitro NRactivity of 100 nmol g−1 DW min−1 in contrastto that of S. spontaneum, Erianthus, and maize,which all had greater leaf NR activity of 212-250nmol g DW−1 min−1 (N. Robinson unpublishedobservations). NR activities in sugarcane roots aregenerally lower (ca. two- to tenfold) than in leaves(Maretzki & Dela Cruz 1967; de Armas et al. 1992;Biggs 2003), indicating that leaves, rather thanroots, are the main site of nitrate reduction. Thisnotion is supported by the increase in nitrate con-tent of stem xylem sap with an increasing externalsupply of nitrate, which also led to the suggestionthat nitrate content in xylem sap may be a moresuitable indicator of N status and availability thanis foliar N content (Ranjith 2007). However, inter-pretation of tissue nitrate levels requires caution aslack of nitrate can also be attributed to alternativeN sources and decreased water availability.

Similar differences in nitrate storage as reportedby Robinson et al. (2011) were observed in


field-grown Erianthus and sorghum, which had upto 30-fold higher nitrate concentrations in stemsthan did sugarcane cultivars (Ishikawa et al. 2009).The giant grass species Andropogon gayanus, aclose relative of Saccharum, acquired ammoniumat a higher rate than nitrate and reduced nitrifi-cation in the soil (Rossiter-Rachor et al. 2009).Similarly, Brachiaria genotypes with enhancedexudation of the biological nitrification-inhibiting(BNI) compound brachialactone were associatedwith reduced N2O emissions from soil (Subbaraoet al. 2009). Thus, increasing nitrate use and eitherreducing the presence of nitrate via BNI com-pounds or agronomic measures or both are possi-ble avenues to increase plant N capture and uptakeefficiency. Selecting sugarcane genotypes with theability to exude BNI compounds could advanceeNUE in addition to improving nitrate use viaimproved uptake, storage, and remobilization.

When three sources of N (ammonium, nitrate,and amino acid glycine) were supplied simultane-ously, roots of field-grown sugarcane incorporatedN sources in the order of preference as ammonium> amino acid > nitrate (Vinall 2011). Amino acidscan account for a substantial proportion of lowmolecular-mass N in agricultural soils (Jamtgardet al. 2010), and similar to nitrate and ammonium,the amino acids enter roots via membrane trans-porters (Nasholm et al. 2009) (see Chapter 11).

Organic oligomers and polymers are alsoacquired by roots (Komarova et al. 2008;Paungfoo-Lonhienne et al. 2008). How muchorganic N contributes to crop N supply is poorlyunderstood because methodology to quantify Nturnover in the rhizosphere is lacking (Nasholmet al. 2009).

Membrane transporters facilitatingnitrogen uptake from soil

Membrane-spanning transporter proteins facil-itate the transport of N compounds from thesoil solution into root cells. Numerous trans-porters for inorganic and organic N compoundshave been identified and categorized accord-ing to transported substrate, substrate affinity,and whether the transporter is constitutive or

inducible (more fully discussed in Chapter 11).Constitutive transporters are present irrespectiveof substrate availability, whereas inducible trans-porter systems respond to presence and demand ofsubstrate. Low-affinity transporters (LATS) facil-itate uptake at high (>1 mM) substrate concen-trations, and high-affinity transporters (HATS)do so at low (μM) substrate concentrations.

In the model plant Arabidopsis thaliana, sixsequences encode for ammonium transporters(AMT), of which AtAMT1;1 and AtAMT1;3have key roles, typified by an approximate 60%reduction in ammonium influx in AMT1;1/3-deficient mutants under N-deficient growth con-ditions (Ludewig et al. 2007).

Also, in Arabidopsis, two types of nitrate trans-porters, NRT1 (53 NRT1 genes encoding mostlyLATS) and NRT2 (seven NRT2 genes encod-ing HATS), enable transport of nitrate acrossthe plasma membrane and tonoplast to enablexylem and phloem loading and unloading (Tsayet al. 2007). Two NRT1 members, AtNRT1.1 andAtNRT1.2, and two NRT2 members, AtNRT2.1(together with a second protein termed NAR2)and AtNRT2.2, facilitate nitrate uptake into roots(Tsay et al. 2007; Dechorgnat et al. 2011). Othermembers of the NRT1 family are involved in load-ing and unloading of nitrate to and from xylemvessels (NRT1.5, NRT1.8) and phloem (NRT1.9),while chloride channel (CLC) and NRT2 mem-bers facilitate nitrate transport into the vacuoles(Tsay et al. 2007; Dechorgnat et al. 2011).

Uptake of organic N, including amino acidsand di- and tripeptides, into Arabidopsis rootsoccurs mainly through the amino acid transportersAtLHT1 and AtAPP5 (Nasholm et al. 2009) andthe peptide transporters AtPTR1 and AtPTR5(Komarova et al. 2008). Overexpression of peptidetransporter genes enhanced growth of Arabidop-sis on peptide N (Komarova et al. 2008), whileoverexpression of the OsAMT1-1 gene increasedammonium uptake into rice roots (Hoque et al.2006).

N transporters have not been characterizedin sugarcane, although they have been in otherplants, where they show considerable homologyacross plant species (Color Plate 8.2). Nitratetransporters for uptake into roots and xylem


loading are of particular interest for crop pro-duction in soils with high nitrifying potential.As discussed above, sugarcane may differ fromrelated grain crop species, which have a greaterpreference and ability for uptake and storage ofnitrate (Robinson et al. 2011). How much knowl-edge on nitrate transporters can be deduced fromArabidopsis remains to be established as analy-ses of maize, rice, sorghum, and Brachypodiumhave shown that Arabidopsis may not be the idealmodel for interpreting nitrate transport in grasses.For example, there is a significant separation inphylogeny of the nitrate transporter gene NRT2between dicotyledonous and moncotyledonousplants (Plett et al. 2010). Analysis of transportertranscripts in expressed sequence tag (EST) col-lections of sugarcane will advance knowledge onregulatory aspects of N uptake in this crop.

In addition to transporters, anatomical andmorphological differences of roots may influ-ence nitrate acquisition by sugarcane. N-limitedplants in solution culture with 200 μM nitratehad twofold greater maximal nitrate uptake rates(Vmax) than plants well-supplied with nitrate. Forexample, one Hawaiian cultivar had higher initialuptake rates and higher Vmax for nitrate absorptionthan did a different cultivar (Ranjith 2007). Thisdifference was interpreted as possibly attributableto the shorter radial distance for ion transportacross the thinner root cortex of the cultivar withthe higher uptake and absorption rates (Saliendra& Meinzer 1992).

In summary, the fundamental knowledge gen-erated for N transporters in the last decadenow enables targeted research on sugarcane inthe context of soil-relevant N composition andconcentration.

Nitrogen assimilation enzymes andnitrogen transport compounds

Transporters for N compounds and N-assimilating enzymes are main contributors tothe uptake, assimilation, and remobilization of Nthat underlie overall NUE (Masclaux-Daubresseet al. 2010) (Color Plate 8.2). Several key enzymesare involved in the assimilation of acquired N.

The most important enzymes involved in nitrateand ammonium assimilation include NR, nitritereductase (NiR), GS, asparagine synthetase (AS),glutamate synthase (GOGAT), alanine amino-transferase (AlaAT), and aspartate aminotrans-ferase (ASAT) (Color Plate 8.2).

Early investigations of N metabolism in sug-arcane utilized cell culture supplied with 13C-labelled amino acids in an effort to explain obser-vations of increased sett germination and growthof sugarcane plants receiving arginine appli-cation (Nickell & Kortschak 1964). Sugarcanecells demonstrated rapid uptake of arginine andincreased growth (Maretzki et al. 1969a, b). Argi-nine additions increased production of polyamineprecursor N-carbamylputrescine (Maretzki et al.1969b), and endogenously supplied arginine wasnot limiting to protein synthesis, suggesting thatarginine was acting as a regulatory signal ratherthan a source of N (Maretzki et al. 1969a).

The stable isotope 15N has been used to trace Nmetabolism pathways and to determine the trans-port and storage forms of N in sugarcane plants.Following application of 15N-labelled ammoniumto sugarcane plants in solution culture, the solubleN pool of the roots accumulated most of the 15N-label in glutamine and asparagine; only a minorproportion of 15N-label was incorporated into ala-nine, aspartate, and other amino acids (Color Plate8.3) (Biggs 2003). The small but highly labeledaspartic acid pool showed rapid flux through thismetabolite in the pathway to asparagine synthe-sis. Following xylem transfer of N into leaves,the 15N-label was transferred to glutamine andglutamate, resulting in higher proportions of 15N-label, but relatively smaller amino acid pools com-pared with the larger but less 15N-labelled poolof alanine (36% of 44 μg N g−1 FW present as15N) (Color Plate 8.3). This suggests the mainflux of N occurs through the enzymes AS andGOGAT.

Low asparagine and glutamine concentrationsin leaves compared to roots indicate rapid incor-poration of root-derived N into other N pools.Compound- and flux-specific analysis of the fateof labeled N could advance knowledge of Nphysiology of sugarcane. It might reveal whetherenhanced activity of alanine amino transferase in


roots might improve N fertilizer use in sugarcaneas has been demonstrated for other crops (Shrawatet al. 2008).

Enzymes GS2 and GS1, which facilitate assim-ilation of newly acquired and remobilized ammo-nium from senescing tissues, respectively, have acentral role in NUE. QTL for increased GS activ-ity colocate with QTL for NUE-linked physiolog-ical traits and harvest measures in maize, rice, andwheat. For these three cereal crops, GS is used asa genetic marker for NUE (Hirel et al. 2001, 2007;Yamaya et al. 2002; Habash et al. 2007). However,there was not a direct relationship between wholeleaf or root GS activity and vegetative biomassor iNUE in young, glasshouse-grown sugarcane(Robinson et al. 2007). A possible reason that sug-arcane responds differently to N may be that graincrops have a positive relationship between GSactivity, N remobilization, and kernel filling, butnot for biomass production. A negative correla-tion was discovered between root GS activity andbiomass production in Lotus japonicus (Limamiet al. 1999).

In sugarcane, one gene (scGSI) encodes fordodecameric GS, while four genes (scGS1.a,scGS1.b, scGS1.c) encode for octameric GS in thecytosol and (scGS2) in the plastids (Nogueira et al.2005). Expression of cytosolic GS in a Braziliancultivar that has good performance with low Ninput was compared with that in S. barberi, whichperforms poorly with low N input. The expres-sion of scGS1.b was more responsive to ammoniumaddition in modern cultivar plantlets (upregulatedfourfold after 12 h) compared with S. barberi,which maintained consistent scGS1.b expressionthroughout all ammonium treatments (Nogueiraet al. 2005). Further characterization of GS activ-ity and N uptake rates and feedback mechanismmay clarify the relationship between ammoniumassimilation, NUE, and yield.

In addition to the well characterizedGS/GOGAT cycle, AS has been proposed asan enzyme contributing to ammonium assimi-lation, not only in catalyzing the generation ofasparagine from the glutamine precursor, but alsoin the assimilation of ammonium (Color Plate 8.3)(Masclaux-Daubresse et al. 2010). An increasedsupply of ammonium and amino acids induces

expression of AS (Lea & Miflin 2011) to gener-ate asparagine (ASN), which is the main form ofN that is transported in numerous plant species.Accumulation of ASN in root, shoot, and xylemsap with increasing N supply by young glasshouse-grown sugarcane and mature field-grown sugar-cane indicates that ASN is a significant transportand storage form of N in sugarcane (Fig. 8.6;Fig. 8.7). Whereas leaf tissue of fertilized andunfertilized field-grown plants had similar aminoacid profiles, the xylem sap of N-fertilized plantshad 20-fold higher soluble N concentration with75% of the N in the form of ASN (Fig. 8.7). Thehigher xylem sap content than leaf tissue contentof N is consistent with the notion that total leaf N isless sensitive to N supply than is stem N (Steven-son et al. 1992; Keating et al. 1997). The aminoacid profiles of different aged leaf and internodeat harvest were similar; it remains to be shownwhether greater variation occurs during early cropgrowth.

Nitrate content increased in the roots and xylemsap, but not in leaves, of glasshouse-grown sug-arcane plants in response to increasing nitratesupply (Fig. 8.6). Field-grown sugarcane plantscontained minimal nitrate (Fig. 8.7). Determin-ing whether the low nitrate content in tissues wasdue to low nitrate in the soil at harvest, a consis-tently low rate of nitrate uptake, a lack of long termstorage of nitrate, or a combination of these fac-tors would require sampling throughout the entirecrop cycle.

Amino-N content and impurity of extractedculm juice are key parameters affecting color ofraw sugar and, ultimately, refining costs. Jacksonet al. (2006) examined genotypes in context ofculm juice color and N fertilizer rates over twocrop cycles. Genotypes differed up to twofold injuice color and amino-N content but ranked dif-ferently between years due to inter-year differ-ences in N status. With increasing N application,amino N concentration of juice ranged from 122to 425 mg N kg−1 in the plant crop and 68 to184 mg N kg−1 in the ratoon crop, indicating thatgenotypes differ in their capacity to accumulateexcess N as amino acids (Jackson et al. 2006).Evaluating genotypic differences in juice N con-tent and the links with biomass production could


Fig. 8.6. Soluble N composition of Q141 sugarcane grown in solution culture for 49 days on either nitrate or ammonium atthree N concentrations, 0.5, 2, and 8 mM. Bars represent amino compositions of root, xylem sap, and leaf. Data representpooled samples (n = 2). Values above bars are total soluble N concentrations (μg N g−1 FW in roots and leaf, μg N ml−1 xylemsap). Reproduced from Biggs (2003).

advance knowledge of crop N requirements andother growth limitations.

Nitrogen and carbon metabolism

Despite recent analyses of plant N and Cmetabolism, a better integration from the subcel-lular to whole plant levels is required for achiev-ing crop yield benefits from this knowledge. Forexample, it has been shown for C3 plants grownunder conditions that lead to temporary deficit ofC compounds that amino acids are catabolized as asource of C for the tricarboxylic acid cycle (Nunes-Nesi et al. 2010). In addition, several lines of evi-dence demonstrate that sugars stimulate aminoacid and protein biosynthesis. These results led tothe statement by Nunes-Nesi et al. (2010, 973) that“current knowledge indicates that plants possessintricate regulatory machinery that coordinatesthe capacity of N assimilation with C metabolism,nutrient availability, and other environmental fac-

tors, and the demands placed by plant growth anddevelopment.”

Nitrate reductase, a key enzyme in the Nmetabolism of nitrate-using plants, is a well-studied component of N and C interactions. NRis induced by the presence of nitrate, repressed byamino acids, and not induced in C starvation con-ditions (Nunes-Nesi et al. 2010). It is unknownwhether sugarcane, due to its unusual sucrose-dominated metabolism, differs from other Poaceaein the regulation of N and C metabolism as well asmetabolic regulation of starch synthesis and degra-dation (Sulpice et al. 2009).

A break-through toward understanding theregulation of N and C interactions was madewith discovery of the PII signal transduction pro-teins in prokaryotes (Arcondeguy et al. 2001). PIIproteins, found in bacteria, Archaea, and plants,bind reversibly to ATP and 2-oxoglurarate andhelp coordinate carbon and nitrogen assimilationby regulating the activity of signal transduction


Fig. 8.7. Composition of soluble N in stem, xylem sap exudate, and leaf of cv. CP51-21 at harvest 300 days after applicationof 0 or 450 kg N ha−1. Plants were grown in subtropical Bundaberg. Stem tissue includes immature internodes 5 to 6 (Y),mature internodes 9 to 10 (M), and older internodes 15 to 16 (O); leaf tissue includes youngest fully expanded leaf (Y) andoldest green leaf (O). Bars represent soluble N components and total soluble N concentration in stem (μg N g−1 FW), xylemsap exudate (μg N ml−1), and leaf (μg N g−1 FW) indicated above each bar for pooled samples of four replicates. Reproducedfrom Biggs (2003).

enzymes in response to diverse signals (Nuns-Nesiet al. 2010 and references therein).

In higher plants PII is a chloroplastic protein.N-acetyl glutamate kinase (NAGK), a chloroplas-tic arginine biosynthesis enzyme, has been identi-fied as a target for PII binding (Chen et al. 2006).High N supply and low levels of 2-oxoglutaratefavor the formation of a PII–NAGK complex,which reduces feedback inhibition and stimu-lates arginine synthesis and N storage (Chenet al. 2006). In addition to its roles regulating Nmetabolism, PII has also been implicated in regu-latory function of chloroplastic fatty acid synthesis(Feria Bourrellier et al. 2010).

Glutamate is also involved in a signaling net-work that allows adaptation to changes in N rela-tions (Forde & Lea 2007). Neither PII nor glu-tamate has been explored for having a role in Nand C metabolism of sugarcane. Whole-genome

transcriptome analysis aimed at elucidating C andN sensing showed that ca. 50% of the Arabidop-sis transcriptome is controlled by C, N, or C-N interactions. Gene regulation of these path-ways occurs at several levels including posttrans-lational control by microRNAs (Gutierrez et al.2007). Metabolome analysis can provide insightinto N and C interactions; however, several hur-dles, including use of standard protocols and com-prehensive screening of metabolites, have to beovercome (Fukusaki & Kobayashi 2005). Progresshas been made in understanding the relationship ofC metabolites and sucrose accumulation (Glassopet al. 2007), C partitioning (Rae et al. 2007), andaccumulation of alternative C compounds (Chonget al. 2007). The International Sugarcane GenomeSequencing Project will facilitate research aimedat integration of data from genome, transcriptome,and metabolome analysis (Manners & Casu 2011).


IMPROVING NITROGEN USEEFFICIENCY THROUGH GENETICENGINEERING

Manipulation of gene function and engineeringnew traits through transgenesis is practiced inmany crops including sugarcane (Lakshmananet al. 2006; James 2010). Over 30 transgenes havebeen introduced into sugarcane (Chapter 24), butnone to date have been aimed at improving NUE.Crop species and Arabidopsis have been used tostudy the physiology of N in plant growth anddevelopment (Masclaux-Daubresse et al. 2010).The relative importance of N uptake, assimilation,transport, storage, and remobilization for improv-ing NUE vary with growth conditions, phenologi-cal stage, and plant species. All processes involvingN are potential targets for genetic engineering toimprove NUE. To date most efforts to improve Nrelations have been directed at increasing N uptakeby modifying N transporter activities or manipu-lating early steps of N metabolism through NR orGS. Improving NUE by upregulating nitrate andAMT and manipulating GS and other N assimi-lation enzymes have shown promise (reviewed byGood et al. 2004 and Xu et al. 2012) and can beused to guide current research.

Although constitutive expression of ahigh affinity nitrate transporter in Nicotianaplumbaginifolia increased nitrate uptake underlow N growth condition, it did not translateinto improved nitrate use or enhanced NUE(Fraisier et al. 2000). Rogato et al. (2010) ectopi-cally expressed the Lotus japonicus ammoniumtransporter genes AMT1;1 and AMT1;3 in N.plumbaginifolia to increase ammonium uptake,but the obtained increase in ammonium uptakedid not improve growth of the transgenic lines.Collectively, such studies suggest that upregula-tion of nitrate and AMT alone do not improveNUE. However, this should not be surprisingconsidering the diverse roles of transporters andthe large number of transporter genes (Rogatoet al. 2010).

With nitrate considered the dominant source ofN for crops, efforts have focused on manipulatingNR activity (Vincentz & Caboche 1991; Ferrario-Mery et al. 1998). Constitutive overexpression of

the NR gene in N. plumbaginifolia increased NRactivity up to 150% but did not affect growth ortotal N content (Quillere et al. 1994). The effectsof upregulation and manipulation of NR or NiRgenes in several species included increased tol-erance to short-term drought stress by delayingloss of NR activity; however, this did not improvegrowth or NUE (Ferrario-Mery et al. 1998; Taka-hashi et al. 2001).

iNUE has been improved via overexpression ofcytoplasmic and plastidic GS genes (Masclaux-Daubresse et al. 2010). In addition, expressionof Medicago sativa GS1 in tobacco enhancedgrowth under low N conditions (Fuentes et al.2001). Constitutive expression of heterologousGS1 genes in tobacco, poplar, and Lotus japon-icus resulted in higher biomass, leaf soluble pro-tein, and chlorophyll content (Gallardo et al. 1999;Oliveira et al. 2002; Suarez et al. 2003). Similarly,overexpressing the Phaseolus vulgaris GS1 genein transgenic wheat increased crop growth, grainyield, and grain N levels (Habash et al. 2001).Grain yield increased by as much as 30% inmaize when its own GS1 gene was upregulated[patent AU2007306040(A1)]. Numerous trans-genic studies have manipulated GS but few havetargeted GOGAT. In transgenic rice lines overex-pressing NADH-GOGAT, grain yield improvedby as much as 80% (Yamaya et al. 2002), butother studies did not show such promising results(Masclaux-Daubresse et al. 2010). Functionalcharacterization of GS and GOGAT genes in dif-ferent plant species, including sugarcane, mayhelp identify enzymes as targets for successfultransgenic manipulation.

Transgenic manipulation of downstream Nmetabolism with AlaAT has produced the mostpromising results to date for increasing yield.Tissue-specific upregulation of AlaAT with root-specific promoters in rice and canola improvedgrowth, tissue N content, and yield under N-limiting supply in the field (Good et al. 2007;Shrawat et al. 2008) and is being explored inother species. Although the biochemical reasonsfor the positive effect of AlaAT on NUE remainunresolved, its action via synthesis and degrada-tion of alanine and the role of alanine as intercel-lular N and C shuttle suggest that downstream


manipulation of N metabolism can improve NUE(Shrawat et al. 2008). Effects of increased AlaATexpression in roots and shoots include enhancedN uptake, larger root systems, and up to 40%reduced N fertilizer demand (Good et al. 2007;Strange et al. 2008). Increased NUE of sugar-cane through overexpression of AlaAT is cur-rently under development (Aracadia Biosciences,http://www.arcadiabio.com/).

AS is another N assimilation enzyme targetedfor NUE improvement and is considered impor-tant in ammonium assimilation when GS is lim-iting. Overexpression of AS in tobacco and Ara-bidopsis improved the fitness of plants under N-limiting conditions and increased total proteincontent (Lam et al. 2003; Wong et al. 2004).Improvement of NUE through manipulation ofthe transcription factor Dof1, which regulates sev-eral C-metabolism enzymes, has been exploredfor carbon skeleton production in maize. Thegenes upregulated by Dof1 are N-responsivegenes that improve NUE through a coordi-nated regulation of N and C metabolism genes.Upregulation of Dof1 in Arabidopsis resultedin an approximate 30% increase in N content,higher concentrations of soluble amino acids, andbetter growth under N limitation (Yanagisawaet al. 2004).

The examples mentioned earlier illustrateopportunities and limitations of transgenesis toimprove NUE. Limitations of the reported exam-ples include that (1) most of the studies did notquantify NUE of tested lines; (2) most observa-tions reported are based on laboratory research andremain to be confirmed in more mature plants infield settings; and (3) a mechanistic understandinghas to be established because constitutive upreg-ulation of individual N metabolism genes or Ntransporters may not yield the desired outcome inall situations. This last point is particularly rel-evant for sugarcane as our understanding of Ntransport and metabolism genes, especially at thestructural and functional diversity level, is rudi-mentary. With such knowledge gaps it remainsdifficult to develop an effective transgenic strat-egy to improve NUE under crop production con-ditions. Contrasting N source preference of sugar-cane compared to grain crops might be advanced

by examining the effects of manipulating nitrateuptake and transport genes.

CONCLUSIONS

Although knowledge of sugarcane N physiologylags behind that in more heavily researched graincrops, it is growing. Research aimed at improvingNUE in sugarcane will increasingly be informedby field-based research as breeding efforts targetresource use efficiency in addition to productiv-ity. With nutrients and water limiting yield inmost sugarcane production systems, selection ofgenotypes having combined efficiencies is partic-ularly attractive. Concerted efforts across sugar-cane growing nations will advance knowledge ofN physiology and its application for improvingNUE.

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