Next Article in Journal
Recovery of Degraded Urochloa Pasture: Effects of Polyhalite on Dry Mass Accumulation and Macronutrient Dynamics
Previous Article in Journal
A Mycorrhiza-Associated Receptor-like Kinase Regulates Disease Resistance in Rice
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Agronomy
Volume 15
Issue 10
10.3390/agronomy15102299
agronomy-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Potential Contributions of Residual Soil Nitrogen to Subsequent Ratoon Sugarcane Crops in the Wet Subtropics

by
Terry James Rose
1,*,
Joshua Rust
2,
Michael Timothy Rose
1 and
Lukas Van Zwieten
2
1
Faculty of Science and Engineering, Southern Cross University, 1 Military Rd, Lismore, NSW 2480, Australia
2
New South Wales Department of Primary Industries, Wollongbar, NSW 2477, Australia
*
Author to whom correspondence should be addressed.
Agronomy 2025, 15(10), 2299; https://doi.org/10.3390/agronomy15102299
Submission received: 31 August 2025 / Revised: 23 September 2025 / Accepted: 25 September 2025 / Published: 29 September 2025
(This article belongs to the Section Soil and Plant Nutrition)
Browse Figures
Versions Notes

Abstract

Nitrogen (N) fertiliser recommendations for ratoon sugarcane crops in the Australian subtropics do not specifically account for residual soil N. The present study was undertaken to determine whether residual soil N is sufficiently high to warrant incorporation into current N fertiliser guidelines in subtropical Australia. Nine soil cores were taken to 1 m depth (separated into 0–20, 20–40, 40–60, 60–80 and 80–100 cm layers) in 25 fields in the Australian subtropics after sugarcane harvest and assessed for soil pH, total carbon and nitrogen and mineral N (NO3− + NH4+) concentrations, and potentially mineralisable N (PMN) in the top 40 cm. Root weight in each soil layer was also measured in one core per field to determine rooting depth. When coupled with 14 d PMN in the top 40 cm, total available N ranged from 44–346 kg N ha−1, which could potentially contribute 30–100% of the typical 150 kg N ha−1 accumulated in shoots of ratoon cane crops in the region. Further field studies are required to determine the actual contributions that these N reserves can make to the N nutrition of ratoon cane crops, and the ramifications of those contributions to fertiliser recommendations.

1. Introduction

Sugarcane (Saccharum officinarum L.) is predominantly grown in tropical regions of the globe, and is a key crop in the Australian tropics and wet subtropics. Owing to the large quantities of biomass produced, large amounts of nitrogen (N) fertiliser—in the order of hundreds of kg N ha−1 [1,2]—are applied to crops to maintain biomass and sugar yields. With increasing N fertiliser rates the efficiency of fertiliser-N recovery tends to decline, leading to substantial N losses through nitrate (NO3−) leaching or ammonia (NH3) volatilisation in sugarcane farming systems [3]. This is particularly problematic when sugarcane is cultivated near environmentally sensitive areas. In Australia, sugarcane farming is a significant contributor to the anthropogenic loads of N entering the Great Barrier Reef lagoon [4]. In light of the proximity of Australian sugarcane growing regions to environmentally sensitive areas including rivers and coastal environments, there is an urgent need to improve to efficiency of N fertiliser use in these systems.
A number of management options including trash (cane residue) retention (e.g., [5]) or the use of enhanced-efficiency N fertilisers under specific climatic conditions (e.g., [6]) can improve N use efficiency in sugarcane systems. However, studies in the Australian wet subtropics have found limited or no improvements in N use efficiency from the use of enhanced-efficiency N fertilisers [7,8,9,10], and retention of trash is only likely to have minor impacts on N use efficiency in the long term [5]. Modelling studies suggest that a more promising approach to improving N use efficiency in cane farming systems and reducing N losses to the environment is the optimisation of N fertiliser rates [11]. One method proposed for refining N fertiliser recommendations is the N replacement system, which involves an estimate of expected cane yield coupled with an estimate of the N required to achieve each tonne of estimated yield [12]. These concepts are also incorporated into the Australian ‘Six Easy Steps’ guidelines, which further require an estimate of potentially mineralisable N from the soil [13]. Essentially, for a given expected crop yield and N demand, crop N fertiliser requirements are a function of the amount of N that will be provided to the crop from existing soil N sources over the season, and the estimated recovery of applied N fertiliser.
The Six Easy Steps guidelines use an estimate of the amount of N provided to the crop from existing soil reserves based on a N mineralisation index that uses soil texture/colour or soil total carbon (C) or total N as an indicator of mineralisable N [13]. However, in other cropping situations, quantifying soil mineral N (i.e., NH4+ and NO3−) reserves to 1 m depth, in addition to a N mineralisation estimate, can further refine the estimation of the potential amount of N that a crop may acquire from the soil over the course of the season [14]. The availability of any mineral N at depth to the ratoon cane crop also depends on the crop rooting depth. Whether sugarcane N fertiliser guidelines can be refined by including an estimate of mineral N to depth in soils prior to N fertiliser application is not known, largely due to a paucity of data on mineral N in soils following cane harvest and the distribution of cane roots of modern cultivars following harvest.
The present study was therefore undertaken to investigate root distributions and the quantities and location (depth) of mineral N pools in subtropical sugarcane fields following cane harvest and prior to N fertiliser application to ratoon crops. Mineral N pools, soil carbon (C) and root distribution were quantified in 0.2 m increments to a depth of 1 m, and potentially mineralisable N (PMN) to 40 cm, following the 2016 cane harvest in 25 fields in the Australian wet subtropics to determine whether available mineral N reserves are sufficient to warrant incorporation into current sugarcane N fertiliser guidelines. Refinement of current N fertiliser recommendations could lead to financial savings by reducing excess N fertiliser application as well an environmental benefit from a reduction in N losses from sugarcane farming systems.

2. Materials and Methods

The location of each of the sampled fields is given in Table 1, along with dates of the previous cane harvests and dates of soil coring.

2.1. Determination of Soil Indicators

Soil samples were obtained from 25 commercial sugarcane fields from the Clarence, Richmond and Tweed River valleys in northern NSW, Australia, following cane harvest in 2016. For soil C and mineral N assessment, cores were taken from three locations within a field (A, B and C in a straight line transect 40 m apart), and at each location three cores were taken (i.e., 9 cores per field in total) to 1 m depth using a hydraulic rig. Cores were separated in the field with a knife into 0–20 cm, 20–40 cm, 40–60 cm, 60–80 cm, 80–100 cm soil zones then stored in a portable cooler and transferred back to the laboratory for further analysis. Samples of the same depth from each of the three cores taken at a given location in a field were pooled to form a composite sample. One additional core was taken from each field root assessment, which was transferred back to the laboratory as an intact core. Bulk density from each depth was determined by drying soil cores (above) of known length and dividing the dry mass by the volume of the sample (determined using the internal diameter of the corer). These measurements were done for each depth (0–20 cm, 20–40 cm, 40–60 cm, 60–80 cm, 80–100 cm) of all nine cores from each site. After weighing, the three samples from each location (A, B, C) for each depth were pooled to form three composite samples for each depth for each site.
The dried composite soil samples were sieved to 2 mm. The samples were then analysed for pH (CaCl2) and 2 M KCl-extractable NH4+ and NO3− using methods from Rayment and Lyons [15]. For total N and C, a subsample of sieved soil was ground to a powder and a 0.2 g subsample of ground material was analysed for total N and C by Dumas combustion using an vario MAX CN analyser (Elementar, Langenselbold, Germany). Potentially mineralisable N in the 0–20 cm and 20–40 cm depth layers was quantified using the ‘short term’ (14 d) method described by Allen et al. [16]. Total PMN to 40 cm depth was calculated by summing the PMN from the 0–20 cm and 20–40 cm depth layers.

2.2. Determination of Root Indicators

Root material was obtained by sectioning the intact cores in the lab with a knife into 0–20 cm, 20–40 cm, 40–60 cm, 60–80 cm, 80–100 cm soil zones, then washing soil from roots with tap water over a 2 mm sieve. Root material was then dried in an oven for 5 d at 40 °C and weighed.

2.3. Statistical Analysis and Graphing of Data

For soil mineral N, total N and total C concentrations, mean values were calculated from the three replicate samples from each field, and data are presented with SEM in Supplementary Table S1. These means were then used to derive mean values and standard errors for each catchment (n = 8 for the Clarence catchment, n = 16 for the Richmond catchment and n = 3 for the Tweed catchment) using the ‘summarySEwithin’ function from the Rmisc package (version 1.5.1) [17] in the R statistical environment (version 4.4.0) [18]. Total NH4+ and NO3− in each depth layer (kg ha−1) were calculated by multiplying the respective concentration (mg kg−1) of each of the three in-field replicates by the single measurement of bulk density (kg m3) and the volume of soil in the layer per ha (2000 m3). Total mineral N per ha to a depth of 1 m was calculated by summing the NH4+ and NO3− from all five depth layers in each of the three locations in the field. Total available mineral N per field was calculated by summing the mean mineral N contents from each depth layer where roots were present. Total available N was calculated by summing the Total available mineral N and the PMN to 40 cm depth. Figures were prepared using the ggplot2 package (version 3.5.1) [19] within the R statistical environment.

3. Results

3.1. Soil pH and Concentrations of Total C, Total N and Mineral N Throughout Soil Profiles Across the Clarence, Richmond and Tweed River Catchments

Means of soil pH (CaCl2) in the top 0–20 cm layer of soil ranged from around 4.2 in the Tweed River catchment to over 4.7 in the Richmond River catchment (Figure 1). There was a gradual increase in pH with soil depth in the Clarence and Richmond River catchments, although subsoils were still acidic, with pH means in the 80–100 cm layer of approximately 5.3 and 5.8 for the Richmond and Clarence River catchments, respectively. The mean soil pH did not change substantially with depth in the Tweed River catchment and was <4.5 in all depth layers (Figure 1). However, there was large variability between fields in the Tweed River catchment, with soil pH in the 40–60, 60–80 and 80–100 cm depths < 4 at Murwillumbah and Stott’s Creek but >5 at Tyalgah (Supplementary Table S1).
Total C concentrations were highest in the 0–20 cm layer in all catchments, ranging from around 2.7% in the Clarence catchment to around 5.5% in the Tweed catchment, and generally declined with depth (Figure 2). Mean total C concentration was still >4% in the 20–40 cm soil layer in the Tweed River catchment, and >1% in the 60–80 cm and 80–100 cm layers in both the Tweed and Richmond River catchments. Mean total N was also highest in the 0–20 cm layer in all catchments and declined with depth (Figure 2). Below 40 cm depth total N concentrations were less than 0.15%, with the exception of the 80–100 cm depth at the Teven 1 field (0.18%; Supplementary Table S1).

3.2. Mineral N to a Depth of 1 m, Root Depth and Plant-Available Mineral N

Soil NO3− concentrations were highest in the 0–20 cm layer for all catchments but were still <5 mg kg−1 (Figure 3). The concentrations generally declined with depth in all catchments, with negligible amounts present below 40 cm in all catchments (Figure 3). Soil NH4+ concentrations in the Clarence and Richmond River catchments were also highest in the 0–20 cm layer and generally declined with depth, but concentrations of around 1 mg g−1 and 3 mg kg−1 were still present in all layers below 40 cm in the Clarence and Richmond catchments, respectively. There was no decline in soil NH4+ concentration with depth in the Tweed catchment, with high concentrations of 20–25 mg kg−1 present in all soil layers (Figure 3).
Total mineral N to a depth of 1 m ranged from 18–51 N kg ha−1 in fields in the Clarence River catchment, 13–65 N kg ha−1 in the Richmond River catchment and 165–242 N kg ha−1 in the Tweed River catchment (Table 2). Ammonium was the dominant mineral N form throughout the soil profile at all sites with the exception of the Pimlico 3 field in the Richmond River catchment.
An average of 55% of root mass was in the top 20 cm of the profile across all fields, with a further 10–15% of root mass in each of the 20–40, 40–60 and 60–80 cm zones and only 5% in the 80–100 cm zone (Figure 4). However, these distributions differed between regions. The Clarence and Richmond River catchments had 50–60% of roots in the top 20 cm, with a decline in root mass with soil depth. However, the mean percentage of roots in the 0–20 cm layer in the Tweed River catchment was around 30%, with >15% of root mass present in each of the 40–60, 60–80 and 80–100 cm layers (Figure 4). Only five fields did not have roots present in the 80–100 cm zone (Palmers Channel 2, Pimlico 3, Empire Vale 2, South Ballina 1 and South Ballina 2; Supplementary Table S1).

3.3. Potentially Mineralisable N to a Depth of 40 cm and Total Available N

Soil PMN in the top 40 cm ranged from 15 kg N ha−1 at the Murwillumbah field site to 195 kg N ha−1 at the Tatham 1 field site (Table 2). With the exception of the Tweed River catchment sites and Teven 1 in the Richmond River catchment, the PMN to 40 cm was greater than the available mineral N in the soil profile.

4. Discussion

To determine whether mineral N levels in soil following sugarcane harvest are sufficient to warrant incorporating into existing models that estimate N fertiliser requirements for the subsequent ratoon crops in subtropical Australia, we quantified soil mineral N reserves in 25 fields after the 2016 cane harvest to a depth of 1 m, as well as PMN in the top 40 cm. Cane yields across the fields for the 2016 harvest ranged from 50–90 t ha−1 for 1 year old cane and from 111–211 t ha−1 for 18-month or 2-yr-old cane. These yields are typical for 1- and 2-year cane crops in the region [20] and hence the residual N remaining in the soil after harvest is likely representative of typical seasons. While some time had elapsed between cane harvest and sampling for residual N (up to 65 days), studies from irrigated cane in the tropics indicate N uptake by sugarcane is minimal in the first two months after ratooning [21]. Thus, we presume only a limited amount of N would have been taken up from the soil by the ratoon cane crops in the early spring months following ratooning in the subtropics in our study, and suggest the data provide a reasonable estimate of typical residual mineral N levels in soils after sugarcane harvest in this region. Further, there was no significant correlation between days elapsed between cane harvest and soil sampling and total mineral N to 1 m depth (R2 = 0.001), suggesting the difference in sampling times was not the key driver of residual mineral N. In the Clarence and Richmond River catchments the residual mineral N was typically between 10–65 kg N ha−1 (Table 2). Most of this mineral N was in the form of NH4+ in the top 40 cm zone of soil, which would likely be available to ratoon crops if active roots were present, since sugarcane roots readily take up NH4+ as a N source [22,23]. Sugarcane roots can grow beyond 4 m deep [24,25,26], and while we only sampled to 1 m depth, we found roots were present in the 80–100 cm layer in all but six fields (all located in the Clarence and Richmond River catchments). The bulk of root mass was located in the top 20 cm of the profile, declining with depth such that only 5% of root mass was present in the 80–100 cm layer in the Richmond and Clarence River catchments (Figure 4), consistent with the sharp decline in root density with depth observed in other studies [27,28,29]. The lower density of roots at depth is attributed to the presence of vertical, penetrating roots termed ‘rope roots’ [26]. These roots do not have an abundance of lateral roots emerging from them at depth, but even a low density of vertical, penetrating roots at depths is thought to be sufficient for capture of water deep in the profile or uptake of nutrients, including NO3−, that move to roots via mass flow [30].
The extent to which these deep soil N reserves make a substantial contribution to ratoon cane growth, and therefore warrant inclusion in fertiliser N budgets, depends on whether cane roots quantified after harvest remain functional or whether they subsequently senesce to be replaced with new roots emerging from the regrowing stool. Despite the potential importance of existing roots to ratoon crops, the functionality of these old roots remains unclear [26]. However, while it is apparent that a proportion of rope roots die after harvest, Glover [31] showed that some rope roots at depths > 1 m were able to transfer 32P to new leaves 44 days after harvest, indicating that a proportion of old rope roots remain functional as then new ratoon crop grows. The presence of active biotrophic fungal root symbionts colonising ratoon crop sett root systems for up to 49 days also suggests live root activity for this duration [32]. Twelve of the fields were sampled within 15 days of harvest in our study (Table 1), and it is possible that in-tact roots washed out of soil cores may have been non-functional and in the process of senescing. However, the remaining 13 fields were sampled from 19–65 days after harvest, and we presume that in-tact roots washed from cores in these fields were still functional and could thus contribute to nutrient uptake.
In contrast to the modest residual mineral N levels in soils in the Clarence and Richmond River catchments, soil profiles in the Tweed River catchment had 165–242 kg N ha−1 to a depth of 1 m after harvest (Table 2), and a substantial proportion of roots was observed lower in the profile in fields in the Tweed River catchment, with only 30% of root mass present in the top 40 cm of the soil profile (Figure 4). However, it must be acknowledged that only three fields were sampled in this catchment, owing to the availability of participatory farmers. The most conspicuous feature of the residual mineral N in these soils was the high concentrations of NH4+ at depth (>20 mg N kg−1; Figure 2), coupled with low NO3− concentrations below 40 cm. It is possible that the high proportion of root mass at depth is a direct response to the high concentrations of NH4+ lower in the profile, where high concentrations of NH4+ trigger root proliferation [33], particularly lateral branching [34]. Conversely, it is possible that the presence of roots and root exudates at depth may stimulate N mineralisation [35] and therefore formation of NH4+. Questions still remain as to why this NH4+ is not taken up by crops, and thus reducing the concentrations, or microbially processed to NO3−. Whether the low pH in these soil layers is inhibiting nitrification, or whether other factors are contributing to the disconnect between NH4+ production and immobilisation/assimilation across the Tweed River catchment sites is not known, but provides an interesting avenue for future research.

5. Conclusions

Ratoon cane crops grown in the Australian subtropics generally accumulate up to 150 kg N ha−1 in aboveground plant material [10]. The 8–63 kg N ha−1 in the rooting zones of ratoon cane crops in the Clarence and Richmond River catchments could potentially contribute roughly 5–40% of the crop’s requirements, assuming minimal denitrification or leaching losses. When coupled with 14 d PMN in the top 40 cm, the total available N per ha of 44–250 kg N ha−1 in the Richmond and Clarence River catchments could contribute 30–100% of the crop’s N requirement, and the 194–346 kg N ha−1 in the Tweed River catchment could potentially provide all of the crop’s N requirement. Further field studies are required to determine the actual contributions that these N reserves can make to the N nutrition of ratoon cane crops, and the ramifications of those contributions to fertiliser recommendations.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/agronomy15102299/s1, Table S1: Root mass, root mass %, soil pH and mean soil total N, total C, NO3− and NH4+ concentrations in five soil layers in individual fields in the Clarence, Richmond and Tweed River catchments. SEM (n = 3) is presented for soil total N, total C, NO3− and NH4+ concentrations.

Author Contributions

Conceptualization, L.V.Z. and T.J.R.; methodology, L.V.Z. and M.T.R.; formal analysis, M.T.R.; investigation, J.R., L.V.Z. and T.J.R.; writing—original draft preparation, T.J.R.; writing—review and editing, L.V.Z., J.R. and M.T.R.; funding acquisition, L.V.Z. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the Australian Government Department of Agriculture and Water Resources as part of its Rural R&D for Profit programme. Application ID: AOTGR2-0005.

Data Availability Statement

All data are available from the corresponding author upon reasonable request.

Acknowledgments

We thank Scott Petty and Ken Lisha (NSW DPI) for technical assistance, and 25 NSW North Coast sugarcane growers for providing background information and access to their sugarcane fields. The authors have reviewed and edited the output and take full responsibility for the content of this publication.

Conflicts of Interest

The authors declare no conflicts of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript; or in the decision to publish the results.

References

  1. Keating, B.A.; Verburg, K.; Huth, N.I.; Robertson, M.J. Nitrogen management in intensive agriculture: Sugarcane in Australia. In Intensive Sugarcane Production: Meeting the Challenges Beyond 2000; Keating, B.A., Wilson, J.J., Eds.; CAB International: Wallingford, UK, 1997; pp. 221–242. [Google Scholar]
  2. Skocaj, D.M.; Everingham, Y.L.; Schroeder, B.L. Nitrogen management guidelines for sugarcane production in Australia: Can these be modified for wet tropical conditions using seasonal climate forecasting? Springer Sci. Rev. 2013, 1, 51–71. [Google Scholar] [CrossRef]
  3. Nachimuthu, G.; Bell, M.J.; Halpin, N. Nitrogen losses in terrestrial hydrological pathways in sugarcane cropping systems of Australia. J. Soil Wat. Conserv. 2017, 72, 32–35. [Google Scholar] [CrossRef]
  4. Bell, M.J. A Review of Nitrogen Use Efficiency in Sugar Cane; Sugar Research Australia Ltd.: Brisbane, Australia, 2014; 344p. [Google Scholar]
  5. Meier, E.A.; Thorburn, P.J. Long term sugarcane crop residue retention offers limited potential to reduce nitrogen fertilizer rates in Australian wet tropical environments. Front. Plant Sci. 2016, 7, 1017. [Google Scholar] [CrossRef] [PubMed]
  6. Wang, W.J.; Park, G.; Reeves, S.; Zahmel, M.; Heenan, M.; Salter, B. Nitrous oxide emission and fertiliser nitrogen efficiency in a tropical sugarcane cropping system applied with different formulations of urea. Soil Res. 2016, 54, 572–584. [Google Scholar] [CrossRef]
  7. Rose, T.J.; Morris, S.G.; Quin, P.; Kearney, L.; Kimber, S.; Van Zwieten, L. The nitrification inhibitor DMPP applied to subtropical rice has an inconsistent effect on nitrous oxide emissions. Soil Res. 2017, 55, 547–552. [Google Scholar] [CrossRef]
  8. Rose, T.J.; Quin, P.; Morris, S.G.; Kearney, L.J.; Kimber, S.; Rose, M.T.; Van Zwieten, L. No evidence for higher agronomic N use efficiency or lower nitrous oxide emissions from enhanced efficiency fertilisers in aerobic subtropical rice. Field Crops Res. 2018, 225, 47–54. [Google Scholar] [CrossRef]
  9. Rose, T.J.; Kearney, L.J.; Zeng, J.; Van Zwieten, L.; Rose, M.T. DMPP-urea restricts nitrification in the first month without improving agronomic N use efficiency. Nutr. Cycl. Agroecosyst. 2023, 126, 115–125. [Google Scholar] [CrossRef]
  10. Rose, T.J.; Rust, J.; Van Zwieten, L.; Rose, M.T. Polymer coated urea does not improve nitrogen use efficiency above urea in ratoon sugarcane crops in the wet subtropics. Nutr. Cycl. Agroecosyst. 2024, 130, 255–267. [Google Scholar] [CrossRef]
  11. Thorburn, P.J.; Biggs, J.S.; Palmer, J.; Meier, E.A.; Verburg, K.; Skocaj, D.M. Prioritizing crop management to increase nitrogen use efficiency in Australian sugarcane crops. Front. Plant Sci. 2017, 8, 1504. [Google Scholar] [CrossRef]
  12. Thorburn, P.J.; Biggs, J.S.; Webster, A.J.; Biggs, I.M. An improved way to determine nitrogen fertiliser requirements of sugarcane crops to meet global environmental challenges. Plant Soil 2011, 339, 51–67. [Google Scholar] [CrossRef]
  13. Schroeder, B.L.; Wood, A.W.; Moody, P.W.; Bell, M.J.; Garside, A.L. Nitrogen fertiliser guidelines in perspective. Proc. Aust. Soc. Sugar Cane Technol. 2005, 27, 291–304. [Google Scholar]
  14. Bock, B.R.; Kelley, K.R.; Meisinger, J.J. Predicting N fertilizer needs in humid regions: Summary and future directions. In Predicting N Fertilizer Needs for Corn in Humid Regions; Bock, B.R., Kelley, K.R., Eds.; Bulletin Y-226, Muscle Shoals, A1; National Fertilizer and Environmental Research Center, Tennessee Valley Authority: Muscle Shoals, AL, USA, 1992; p. Ch. 9. [Google Scholar]
  15. Rayment, G.E.; Lyons, D.J. Soil Chemical Methods–Australasia; CSIRO Publishing: Clayton, Australia, 2011. [Google Scholar]
  16. Allen, D.E.; Bloesch, P.M.; Orton, T.G.; Schroeder, B.L.; Skokaj, D.M.; Wang, W.; Masters, B.; Moody, P.M. Nitrogen mineralisation in sugarcane soils in Queensland, Australia: I. evaluation of soil tests for predicting nitrogen mineralisation. Soil Res. 2019, 57, 738–754. [Google Scholar] [CrossRef]
  17. Hope, R.M. Rmisc: Rmisc: Ryan Miscellaneous. R package Version 1.5. 2013. Available online: https://CRAN.R-project.org/package=Rmisc (accessed on 22 September 2025).
  18. R Core Team. R: A Language and Environment for Statistical Computing; R Foundation for Statistical Computing: Vienna, Austria, 2020; Available online: https://www.R-project.org/ (accessed on 25 May 2024).
  19. Wickham, H. Ggplot2: Elegant Graphics for Data Analysis; Springer: New York, NY, USA, 2016. [Google Scholar]
  20. Australian Sugar Milling Council. Sugar Cane Statistics. 2025. Available online: https://sugarmanufacturers.org/annual-industry-statistics/ (accessed on 24 September 2025).
  21. Wood, A.W.; Muchow, R.C.; Robertson, M.J. Growth of sugarcane under high input conditions in tropical Australia. III. Accumulation, partitioning and use of nitrogen. Field Crops Res. 1996, 48, 223–233. [Google Scholar] [CrossRef]
  22. Robinson, N.; Brackin, R.; Vinall, K.; Soper, F.; Holst, H.; Gamage, H.; Paungfoo-Lonhienne, C.; Rennenberg, H.; Lakshmanan, P.; Schmidt, S. Nitrate paradigm does not hold up for sugarcane. PLoS ONE 2011, 6, e19045. [Google Scholar] [CrossRef]
  23. Boschiero, B.N.; Mariano, E.; Trivelin, P.C.O. “Preferential” ammonium uptake by sugarcane does not increase 15N recovery of fertilizer sources. Plant Soil 2018, 429, 253–269. [Google Scholar] [CrossRef]
  24. Battie Laclau, P.; Laclau, J.P. Growth of the whole root system for a plant crop of sugarcane under rainfed and irrigated environments in Brazil. Field Crops Res. 2009, 114, 351–360. [Google Scholar] [CrossRef]
  25. Chopart, J.L.; Azevedo, M.C.B.; Le Mezo, L.; Marion, D. Sugarcane root system depth in three different countries. Proc. Int. Soc. Sugar Cane Technol. 2010, 27, 1–8. [Google Scholar]
  26. Smith, D.M.; Inman-Bamber, N.G.; Thorburn, P.J. Growth and function of the sugarcane root system. Field Crops Res. 2005, 92, 169–183. [Google Scholar] [CrossRef]
  27. Chopart, J.L.; Rodrigues, S.; de Azevedo, M.C.; Medina, C.D. Estimating sugarcane root length density through root mapping and orientation modelling. Plant Soil 2008, 313, 101–112. [Google Scholar] [CrossRef]
  28. Chumphu, S.; Jongrungklang, N.; Songsri, P. Association of physiological responses and root distribution patterns of ratooning ability and yield of the second ratoon cane in sugarcane elite clones. Agronomy 2019, 9, 200. [Google Scholar] [CrossRef]
  29. Pierre, J.S.; Giblot-Ducray, D.; McKay, A.C.; Hartley, D.M.; Perroux, J.M.; Rae, A.L. DNA based diagnostic for the quantification of sugarcane root DNA in the field. Sci. Rep. 2018, 8, 16720. [Google Scholar] [CrossRef] [PubMed]
  30. Barber, S.A. Nutrient uptake by plant roots growing in soil. In Soil Nutrient Bioavailability: A Mechanistic Approach; John Wiley & Sons Inc.: New York, NY, USA, 1995; pp. 85–109. [Google Scholar]
  31. Glover, J.D. The behaviour of the root-system of sugarcane at and after harvest. Proc. S. Afr. Sug. Technol. Ass. 1968, 42, 133–135. [Google Scholar]
  32. Claassens, A.; Nock, C.J.; Rose, M.T.; Van Zwieten, L.; Rose, T.J. Colonisation dynamics of arbuscular mycorrhizal fungi and dark septate endophytes in the sugarcane crop cycle. Rhizosphere 2018, 7, 18–26. [Google Scholar] [CrossRef]
  33. Drew, M.C. Comparison of the effects of a localized supply of phosphate, nitrate, ammonium and potassium on the growth of the seminal root system, and the shoot, in barley. New Phytol. 1975, 75, 479–490. [Google Scholar] [CrossRef]
  34. Lima, J.E.; Kojima, S.; Takahashi, H.; von Wirén, N. Ammonium Triggers Lateral Root Branching in Arabidopsis in an AMMONIUM TRANSPORTER1;3-Dependent Manner. Plant Cell 2010, 22, 3621–3633. [Google Scholar] [CrossRef]
  35. Pries, C.E.H.; Sulman, B.N.; West, C.; O’Neill, C.; Poppleton, E.; Porras, R.C.; Castanha, C.; Zhu, B.; Wiedemeier, D.B.; Torn, M.S. Root litter decomposition slows with soil depth. Soil Biol. Biochem. 2018, 125, 103–114. [Google Scholar] [CrossRef]
Agronomy 15 02299 g001
Figure 1. Mean soil pH in the 0–20 cm, 20–40 cm, 40–60 cm, 60–80 cm and 80–100 cm soil layers in each catchment. Error bars represent SEM (n = 8 for Clarence River catchment, n = 14 for Richmond River catchment, n = 3 for Tweed River catchment).
Figure 1. Mean soil pH in the 0–20 cm, 20–40 cm, 40–60 cm, 60–80 cm and 80–100 cm soil layers in each catchment. Error bars represent SEM (n = 8 for Clarence River catchment, n = 14 for Richmond River catchment, n = 3 for Tweed River catchment).
Agronomy 15 02299 g001
Agronomy 15 02299 g002
Figure 2. Mean soil total carbon (TC) and total nitrogen (TN) in the 0–20 cm, 20–40 cm, 40–60 cm, 60–80 cm and 80–100 cm soil layers in each catchment. Error bars represent SEM (n = 8 for Clarence River catchment, n = 14 for Richmond River catchment, n = 3 for Tweed River catchment).
Figure 2. Mean soil total carbon (TC) and total nitrogen (TN) in the 0–20 cm, 20–40 cm, 40–60 cm, 60–80 cm and 80–100 cm soil layers in each catchment. Error bars represent SEM (n = 8 for Clarence River catchment, n = 14 for Richmond River catchment, n = 3 for Tweed River catchment).
Agronomy 15 02299 g002
Agronomy 15 02299 g003
Figure 3. Mean soil ammonium (NH4+) and nitrate (NO3−) concentrations in the 0–20 cm, 20–40 cm, 40–60 cm, 60–80 cm and 80–100 cm soil layers in each catchment. Error bars represent SEM (n = 8 for Clarence River catchment, n = 14 for Richmond River catchment, n = 3 for Tweed River catchment).
Figure 3. Mean soil ammonium (NH4+) and nitrate (NO3−) concentrations in the 0–20 cm, 20–40 cm, 40–60 cm, 60–80 cm and 80–100 cm soil layers in each catchment. Error bars represent SEM (n = 8 for Clarence River catchment, n = 14 for Richmond River catchment, n = 3 for Tweed River catchment).
Agronomy 15 02299 g003
Agronomy 15 02299 g004
Figure 4. Mean percentage of total roots in the 0–20 cm, 20–40 cm, 40–60 cm, 60–80 cm and 80–100 cm soil layers in each catchment. Error bars represent SEM (n = 8 for Clarence River catchment, n = 14 for Richmond River catchment, n = 3 for Tweed River catchment).
Figure 4. Mean percentage of total roots in the 0–20 cm, 20–40 cm, 40–60 cm, 60–80 cm and 80–100 cm soil layers in each catchment. Error bars represent SEM (n = 8 for Clarence River catchment, n = 14 for Richmond River catchment, n = 3 for Tweed River catchment).
Agronomy 15 02299 g004
Table 1. Catchments and time of sugarcane harvest and soil sampling in 25 subtropical sugarcane fields.
Table 1. Catchments and time of sugarcane harvest and soil sampling in 25 subtropical sugarcane fields.
FieldRiver CatchmentSugarcane HarvestSoil CoringDays Between Harvest and Coring
Maclean 1Clarence27 September 201618 October 201621
Maclean 2Clarence22 September 201618 October 201626
Palmers Island 1Clarence21 August 201624 October 201664
Palmers Island 2Clarence2 October 201624 October 201622
Harwood 1Clarence5 October 201624 October 201619
Harwood 2Clarence5 October 201624 October 201619
Palmers Channel 1Clarence9 October 201624 October 201615
Palmers Channel 2Clarence1 September 201624 October 201615
Pimlico 1Richmond20 September 201628 September 20168
Pimlico 2Richmond25 July 201628 September 201665
Pimlico 3Richmond20 September 201628 September 20168
Coraki Richmond25 September 201610 October 201615
Empire Vale 1Richmond16 November 201617 November 20161
Empire Vale 2Richmond14 November 201617 November 20163
South Ballina 1Richmond25 October 20168 November 201614
South Ballina 2Richmond25 October 20168 November 201614
Tatham 1Richmond30 September 201613 October 201613
Tatham 2Richmond28 September 201613 October 201615
Tatham 3Richmond28 September 201613 October 201615
Teven 1Richmond13 October 20168 November 201636
Teven 2Richmond10 October 20168 November 201639
WoodburnRichmond29 September 20167 October 20168
Stott’s CreekTweed28 November 201614 December 201616
TyalgahTweed27 November 201621 December 201624
MurwillumbahTweed21 November 201621 December 201623
Table 2. Mineral N to 1 m depth in 25 subtropical sugarcane fields and availability based on rooting depth; and 0–40 cm depth layer potentially mineralisable N (14 d). Errors are SEM (n = 3).
Table 2. Mineral N to 1 m depth in 25 subtropical sugarcane fields and availability based on rooting depth; and 0–40 cm depth layer potentially mineralisable N (14 d). Errors are SEM (n = 3).
Field NH4+-N (kg ha−1)NO3−-N (kg ha−1)Total Mineral-N (kg ha−1)Available Mineral N (kg ha−1)14 d PMN to 40 cm (kg ha−1)Total Available N
(kg ha−1)
Maclean 134 ± 2.97.8 ± 0.742 ± 3.04264106
Maclean 227 ± 4.424 ± 2.651 ± 2.151111166
Palmers Island 119 ± 1.25.7 ± 1.525 ± 2.125104129
Palmers Island 214 ± 0.711 ± 3.625 ± 3.225108133
Harwood 115 ± 0.514 ± 2.429 ± 2.4296594
Harwood 219 ± 1.98.9 ± 2.128 ± 3.92896124
Palmers Channel 123 ± 6.17.3 ± 2.230 ± 5.4305989
Palmers Channel 211 ± 1.27.0 ± 1.918 ± 2.0167692
Pimlico 148 ± 6.23.9 ± 0.1 52 ± 6.252107159
Pimlico 227 ± 1.66.2 ± 1.733 ± 3.1335184
Pimlico 313 ± 2.353 ± 16.565 ± 176378141
Coraki 32 ± 3.314 ± 1.647 ± 3.247193240
Empire Vale 121 ± 4.12.1 ± 0.024 ± 4.1244367
Empire Vale 211 ± 0.11.8 ± 0.113 ± 0.28.13644
South Ballina 127 ± 6.72.9 ± 0.429 ± 6.8207797
South Ballina 214 ± 0.23.9 ± 0.618 ± 0.7135770
Tatham 147 ± 2.77.2 ± 1.954 ± 2.154195249
Tatham 236 ± 1.35.4 ± 0.742 ± 1.04278120
Tatham 329 ± 0.317 ± 4.147 ± 3.847153200
Teven 139 ± 205.3 ± 0.644 ± 20444185
Teven 222 ± 1.25.6 ± 2.127 ± 2.1276087
Woodburn42 ± 3.03.2 ± 0.745 ± 2.945194139
Stott’s Creek163 ± 132.3 ± 0.5165 ± 1316529194
Tyalgah236 ± 566.8 ± 0.6242 ± 56242106346
Murwillumbah166 ± 5.74.0 ± 0.1170 ± 5.617025195
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Rose, T.J.; Rust, J.; Rose, M.T.; Van Zwieten, L. Potential Contributions of Residual Soil Nitrogen to Subsequent Ratoon Sugarcane Crops in the Wet Subtropics. Agronomy 2025, 15, 2299. https://doi.org/10.3390/agronomy15102299

AMA Style

Rose TJ, Rust J, Rose MT, Van Zwieten L. Potential Contributions of Residual Soil Nitrogen to Subsequent Ratoon Sugarcane Crops in the Wet Subtropics. Agronomy. 2025; 15(10):2299. https://doi.org/10.3390/agronomy15102299

Chicago/Turabian Style

Rose, Terry James, Joshua Rust, Michael Timothy Rose, and Lukas Van Zwieten. 2025. "Potential Contributions of Residual Soil Nitrogen to Subsequent Ratoon Sugarcane Crops in the Wet Subtropics" Agronomy 15, no. 10: 2299. https://doi.org/10.3390/agronomy15102299

APA Style

Rose, T. J., Rust, J., Rose, M. T., & Van Zwieten, L. (2025). Potential Contributions of Residual Soil Nitrogen to Subsequent Ratoon Sugarcane Crops in the Wet Subtropics. Agronomy, 15(10), 2299. https://doi.org/10.3390/agronomy15102299

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop