Introduction
The sucrose yield in sugarcane depends on carbon assimilation and source–sink interactions, which are two key factors exhibiting significant seasonal variation (De Souza et al., Reference De Souza, Grandis, Arenque-Musa and Buckeridge2018). In subtropical conditions, photosynthesis is constrained by low temperatures and drought, limiting the availability of CO2 due to stomatal closure and also reducing the activity of photosynthetic enzymes (Marchiori et al., Reference Marchiori, Machado, Sales, Espinoza-Núñez, Magalhães Filho, Souza, Pires and Ribeiro2017; Silveira et al., Reference Silveira, Marcos, Frungillo, Moura, Seabra, Salgado, Machado, Hancock and Ribeiro2017; Cerqueira et al., Reference Cerqueira, Santos, Marchiori, Silveira, Machado and Ribeiro2019). Additionally, poor light distribution within plant canopy leads to decreased CO2 uptake by sugarcane (Marchiori et al., Reference Marchiori, Machado and Ribeiro2014). However, sugarcane response to these constraining conditions differs substantially among varieties. For instance, phenotypic plasticity due to water deficit varies among sugarcane genotypes as well as the photosynthetic flexibility under low light conditions (Marchiori et al., Reference Marchiori, Ribeiro, Da Silva, Machado, Machado and Scarpari2010, Reference Marchiori, Machado, Sales, Espinoza-Núñez, Magalhães Filho, Souza, Pires and Ribeiro2017; Sales et al., Reference Sales, Ribeiro, Hayashi, Marchiori, Silva, Martins, Silveira, Silveira and Machado2018).
The strength of the sink (stalks) and the accumulation of sugars in sources (leaves) have also a regulatory effect on sugarcane photosynthesis (McCormick et al., Reference McCormick, Cramer and Watt2006, Reference McCormick, Cramer and Watt2008a; Ribeiro et al., Reference Ribeiro, Machado, Magalhães Filho, Lobo, Martins, Silveira, Yin and Struik2017). Previous studies on sugarcane plants using cold-girdling and sugar feeding techniques have demonstrated that increased leaf sugar concentrations reduce chlorophyll content, Rubisco expression and activity, and overall photosynthetic rates (McCormick et al., Reference McCormick, Cramer and Watt2008b; Lobo et al., Reference Lobo, De Oliveira, Lima, Machado, Ribeiro and Silveira2015; Ribeiro et al., Reference Ribeiro, Machado, Magalhães Filho, Lobo, Martins, Silveira, Yin and Struik2017). On the other hand, enhanced sink demand leads to increased photosynthetic rates and reduced leaf sugar concentration in sugarcane plants (Ribeiro et al., Reference Ribeiro, Machado, Magalhães Filho, Lobo, Martins, Silveira, Yin and Struik2017; Verma et al., Reference Verma, Roopendra, Sharma, Chandra and Kamal2019). This suggests that increased stalk storage capacity and the decoupling of pathways mediating the feedback inhibition of sources by sinks are needed for enhancing sucrose yield in sugarcane (McCormick et al., Reference McCormick, Watt and Cramer2009). Unfortunately, most of studies have primarily focused on source tissues (Huang et al., Reference Huang, Qin, Gui, Zhao, Chen, Wang, Sun, Liao, Li and Lakshmanan2017).
Carbon partitioning varies between leaves and stalks, and this changes the ratio of insoluble to soluble compounds in each sugarcane organ (Mason et al., Reference Mason, Hoang, Botha, Furtado, Marquardt and Henry2022, Reference Mason, Hoang, Botha, Furtado, Marquardt and Henry2023). The carbon allocation is also influenced by the sugarcane phenology, mainly at maturation (Mason et al., Reference Mason, Hoang, Botha, Furtado, Marquardt and Henry2022; García et al., Reference García, Molina, Simister, Taibo, Setten, Erazzú, Gómez and Acevedo2023). While sucrose is rapidly degraded and used in respiration and other biosynthetic pathways when plants are young and in active growth, there is a significant build-up of sucrose concentration in sinks during the maturation (Mason et al., Reference Mason, Hoang, Botha, Furtado, Marquardt and Henry2022). This process is coordinated by several enzymes, including sucrose phosphate synthase (SPS), sucrose synthase (SuSy), and invertases, which are responsible for the breakdown and synthesis of sucrose (Wang et al., Reference Wang, Nayak, Koch and Ming2013), and play a crucial role in determining the final sugar yield (Anur et al., Reference Anur, Mufithah, Sawitri, Sakakibara and Sugiharto2020).
The invertases hydrolyse sucrose to glucose and fructose, playing an important role in the control of metabolic fluxes, phloem loading and unloading, sucrose partitioning, and plant development (Rossouw et al., Reference Rossouw, Kossmann, Botha and Groenewald2010). SPS, on the other hand, is a pivotal enzyme in leaves, controlling the flow of carbon into sucrose (Verma et al., Reference Verma, Upadhyay, Verma, Solomon and Singh2011). SuSy catalyses a reversible reaction, primarily cleaving sucrose in sink tissues (Mirajkar et al., Reference Mirajkar, Suprasanna and Vaidya2016). Additionally, the activity of these enzymes is dependent on environmental conditions (Shanthi et al., Reference Shanthi, Alarmelu, Mahadeva, Lakshmi, Verma, Rajput, Li, Song, Solomon and Rao2023). For instance, acid invertase activity is reduced under low air temperature and water stress (Du and Nose, Reference Du and Nose2002). Moreover, several studies on sugarcane have revealed high genotypic variation in the expression and activity of enzymes involved in carbohydrate metabolism (Verma et al., Reference Verma, Upadhyay, Verma, Solomon and Singh2011; Chandra et al., Reference Chandra, Verma, Islam, Grisham, Jain, Sharma, Roopendra, Singh, Singh, Verma and Solomon2015; Huang et al., Reference Huang, Qin, Gui, Zhao, Chen, Wang, Sun, Liao, Li and Lakshmanan2017; Mason et al., Reference Mason, Hoang, Botha, Furtado, Marquardt and Henry2022).
Despite these findings, our understanding about the enzymatic balance underlying vegetative growth and sugar accumulation in sugarcane remains limited, as well as our knowledge about how varieties differ in sucrose metabolism along crop cycle (Shanthi et al., Reference Shanthi, Alarmelu, Mahadeva, Lakshmi, Verma, Rajput, Li, Song, Solomon and Rao2023). Here, we argue that the differential sucrose yield among sugarcane genotypes is attributed to differences in carbon metabolism and related processes. In-depth knowledge about the factors driving sucrose yield in sugarcane can greatly benefit breeding programmes in releasing varieties with superior performance (Wang et al., Reference Wang, Nayak, Koch and Ming2013; Misra et al., Reference Misra, Mall, Ansari and Ansari2022). In this context, we investigated the carbohydrate dynamics along the crop cycle of two sugarcane varieties differing in biomass production and considered the carbon metabolism in both leaves and stalks. Our aim was to reveal the physiological and biochemical processes underlying the differential sucrose yield and biomass production in sugarcane varieties.
Materials and methods
Plant material and field conditions
A field experiment was carried out with the sugarcane (Saccharum spp.) varieties IACSP95-5000 and IACSP94-2094, two commercial varieties with differential biomass production and yield (Silva et al., Reference Silva, Pires, Ribeiro, Machado, Blain and Ohashi2016). IACSP95-5000 presents high yield in non-restrictive environments (Cruz et al., Reference Cruz, Pacheco, Silva, Almeida, Miranda, Pissolato, Machado and Ribeiro2021), while IACSP94-2094 is considered rustic and drought-resistant and indicated for less favourable environments (Sales et al., Reference Sales, Ribeiro, Silveira, Machado, Martins and Lagôa2013).
The plants were grown on a dystrophic red latosol in Campinas SP, Brazil (22º52′S, 47º04′W, 665 m a.s.l.). The experimental design was in randomised blocks, with four replications and plots composed of six rows of 14 m spaced by 1.5 m. The bud density in planting was 14 ± 1 buds per linear meter. The NPK fertiliser (8:28:16) was applied just before (50%) and 70 days after (50%) planting. In total, plants received 210, 135, and 195 kg ha–1 of N, K, and P, following a previous soil chemical analysis. The field was irrigated with a subsurface drip system, with one drip line per planting row. The drip lines were installed before planting, 0.20 m depth in the furrows. The emitters had nominal flow rate of 1.6 L h–1 and were spaced by 0.50 m. For soil water monitoring, we used the Enviroscan (Sentek Sensor Technologies, Stepney, Australia) and Diviner 2000 capacitance probes (Sentek Sensor Technologies, Stepney, Australia). Measurements were taken every 0.1 m down to 1.0 m depth. Three access tubes per variety were used for Diviner 2000 readings, and one access tube per variety for the EnviroScan probe, each with a length of 1.0 m and internal diameter of 0.05 m. Irrigation was meticulously managed on a daily basis to maintain soil moisture levels close to the field capacity. The water volume applied was calculated daily based on data from the Enviroscan and Diviner 2000. Field capacity was determined following the method outlined by Moraes et al. (Reference Moraes, Libardi and Dourado Neto1993). At 410 days after planting (DAP), irrigation was stopped to induce sugarcane maturation.
Environmental conditions were monitored using an automatic meteorological station installed inside the field plot, with sensors of air temperature (model HMP45C, Campbell, North Logan UT, USA), rainfall (model CS700, Campbell, North Logan UT, USA), wind speed (model 014A, Campbell, North Logan UT, USA), and incident photosynthetic active radiation (PARIN, model LI-190R, LI-COR, Lincoln NE, USA). The above variables were measured every minute, and data stored every five minutes in a data logger model CR1000 (Campbell, North Logan UT, USA). The climatological water balance was calculated according to Thornthwaite and Mather using the practical method described by Rolim et al. (Reference Rolim, Sentelhas and Barbieri1998) and considering 98 mm of soil water storage capacity.
Sampling was performed at four phenological stages, varying in terms of crop growth rate and stalk maturation: (1) maximum growth (158–362 DAP); (2) early maturation (363–397 DAP); (3) medium maturation (398–460 DAP); and (4) late maturation (461–491 DAP).
Leaf gas exchange, photochemistry, water potential, chlorophyll, and nitrogen contents
Measurements of leaf gas exchange, total chlorophyll index (Chl), and leaf water potential (ψ) were taken every ∼30 days, starting at 150 DAP and ending at the final harvest (491 DAP). Leaf gas exchange and Chl were measured in the same region on the first (+1) and third (+3) fully expanded leaves with visible dewlap of four plants in each plot. As the preliminary statistical analyses revealed no differences between leaves +1 and +3, data from both leaves were pooled.
Leaf gas exchange was evaluated with an infrared gas analyser model LI-6400XT (LI-COR, Lincoln NE, USA) equipped with a fluorometer model 6400-40 LCF (LI-COR, Lincoln NE, USA). The measurements were performed under air CO2 concentration of 400 μmol mol−1 and natural variation of air temperature, relative humidity, and light intensity, with leaf exchange values being recorded after temporal stability and when the low total coefficient of variation was lower than 5%. Instantaneous measurements were taken every 2-hour intervals, predominantly on clear days, from 7h00 to 17h00. We assessed leaf CO2 assimilation (A), stomatal conductance (g s), intercellular CO2 concentration (C i), and the effective quantum efficiency of photosystem II (ФPSII). The instantaneous carboxylation efficiency (k) was calculated as A/C i. From the diurnal CO2 assimilation curves derived from these instantaneous measurements, the diurnal-integrated CO2 assimilation (A i, mol CO2 m–2 d–1) was estimated. Herein, g s, k, and ФPSII are shown at 13h00, when there was the maximum light intensity.
The Chl was measured using a portable chlorophyll meter (ClorofiLOG CFL1030, Falker, Porto Alegre RS, Brazil) and considered the sum of chlorophylls a and b. ψ was measured in leaf +3 between 12h00 and 14h00, with a pressure chamber (model 3005, SoilMoisture Equipment Corp., Santa Barbara CA, USA). The leaf nitrogen concentration (LNC) was quantified in lyophilised leaves after digestion with sulphuric acid by using the colorimetric method proposed by Baethgen and Alley (Reference Baethgen and Alley1989). Such quantification was done in leaves +3 sampled at the maximum growth, and at the early and late maturation stages.
Total soluble sugars, starch, and sucrose
Total soluble sugars and sucrose were extracted from samples (0.75 g) of leaves +1 and stalks, using 10 mL of a solution composed by methanol, chloroform, and water (12:5:3, v:v:v) (Bieleski and Turner, Reference Bieleski and Turner1966). Plant extracts were centrifuged at 600 g for 10 min, and the supernatants were homogenised with chloroform and water. After 24 h, the aqueous phase was used to determine the concentration of total soluble sugars and sucrose (Van Handel, Reference Van Handel1968). Starch was measured in leaves by the enzymatic method proposed by Amaral et al. (Reference Amaral, Gaspar, Costa, Aidar and Buckeridge2007), using 500 μL (120 U mL−1) of α-amylase (EC 3.2.1.1) of thermostable Bacillus licheniformis (code E-ANAAM, Megazyme, Bray, Ireland) and 500 μL (30 U mL−1) of amyloglucosidase (EC 3.2.1.3) from Aspergillus niger (code E-AMGPU, Megazyme, Bray, Ireland). The quantification of such carbohydrates was performed at the maximum growth and late maturation for leaves and at the early, medium, and late maturation for stalks. Stalks were segmented into three portions: top, medium, and bottom.
Enzymes of carbohydrate metabolism
The enzymatic analyses were performed in leaves and stalks (top, medium, and bottom portions) at the maximum growth and late maturation. The extraction of enzymes from stalks and leaves followed the method of Grof et al. (Reference Grof, Albertson, Bursle, Perroux, Bonnett and Manners2007), with modifications: 0.5 g of fresh samples were macerated in a mortar with 5% polyvinylpolypyrrolidone (PVPP) in liquid nitrogen and 3.5 mL of 50 mM Hepes buffer (pH 7.5), containing 10 mM MgCl2 and 1 mM EDTA. After centrifugation at 14 000 g for 20 min, 2.5 mL of the supernatant was collected and desalted on a Sephadex G25 column (PD-10, GE), previously saturated with the extraction buffer. The extract collected from the column after elution with the same extraction buffer was used to determine protein content and carry out enzymatic analyses. All extraction was carried out at low temperature (0–4°C), and the protein concentration in the enzymatic extract was determined following Bradford (Reference Bradford1976).
The activities of soluble acid (SAI, EC 3.2.1.26) and neutral (SNI, EC 3.2.1.26) invertases were measured according to Zhu et al. (Reference Zhu, Komor and Moore1997), using 240 mM sucrose at 37°C for 30 min. For SAI activity, the reaction was stopped by adding 2.5 M Tris and incubating at 100°C for 3 min. SNI assay reaction was stopped by incubation at 100°C for 3 min. For both SAI and SNI assays, the reducing sugars released were determined by the Somogyi–Nelson method (Nelson, Reference Nelson1944; Somogyi Reference Somogyi1945, Reference Somogyi1952). SuSy (EC 2.4.1.13) and SPS (EC 2.4.1.14) activities were evaluated according to Hubbard et al. (Reference Hubbard, Huber and Pharr1989), with modifications suggested by Zhu et al. (Reference Zhu, Komor and Moore1997). SuSy can either synthetise or hydrolyse sucrose, and its activity was measured towards sucrose synthesis. To determine the SuSy activity, the crude extracts were incubated with 50 mM of Tris-HCl (pH 7.5), containing 15 mM of MgCl2, 25 mM of fructose, and 50 mM of UDP-glucose. For the SPS activity, crude extracts were incubated in 200 mM of Tris-HCl (pH 7.5), containing 10 mM of MgCl2, 8 mM of fructose-6-phosphate, 40 mM of glucose-6-phosphate, 50 mM of UDP-glucose, and 2 mM of EDTA. SuSy and SPS activities were evaluated at 30°C for 0, 30, 60, and 90 minutes. Then, reactions were stopped by boiling (100 °C) for 3 min. Sucrose produced by both enzymes was assayed according to Van Handel (Reference Van Handel1968).
Biomass production and leaf area
Evaluations of biomass were made every ∼30 days (September to August) until the harvest. All plants from two linear metres of the central line of each plot were harvested for biomass quantification. Stalks and leaves were weighted to determine fresh biomass. For evaluating dry matter, fresh stalk and leaves were ground in a forage chopper and then a sample from each fraction was dried in a forced-air circulation oven at 60°C. The leaf area of all green leaves was measured with an electronic planimeter model LI-3000C (LI-COR, Lincoln NE, USA) coupled to the LI-3050C table accessory (LI-COR, Lincoln NE, USA). The leaf area index (LAI) was estimated considering all green leaves in 3 m2.
Data analyses
The experimental design was in complete randomised blocks, with four replications. Each block was composed by two sugarcane varieties. The causes of variation were sampling time and sugarcane varieties, and data were subjected to ANOVA followed by Scott–Knott post hoc test (P < 0.05). The statistical analyses were performed using the software Rbio (Rbio 143, Viçosa MG, Brazil), and all graphs were created using GraphPad Prism9 (GraphPad Software Inc., San Diego CA, USA).
Results
Environmental conditions and leaf water status
During the experimental period, the accumulated rainfall was 2,039 mm, and the air temperature ranged from 12 to 30°C, with an average air temperature of 21°C (Figure 1a). Plants faced water deficit at the beginning of crop cycle and during late maturation, when irrigation was stopped to induce sugarcane maturation. The maximum accumulated water deficit was 69 mm (Figure 1b).
Monthly rainfall, irrigation, average, maximum and minimum air temperature (a), and climatological water balance (b). The experimental period is delimited by dotted lines.
Along the crop cycle, IACSP95-5000 and IACSP94-2094 showed similar ψ, with the lowest values found at the beginning of maximum growth stage (about −1.25 MPa), with significant recovery of ψ noticed at the rainy season. ψ decreased again at the end of maturation for both varieties (Supplementary Material Figure S1).
Leaf gas exchange, photochemistry, chlorophyll, and nitrogen contents
Diurnal-integrated leaf CO2 assimilation (A i), stomatal conductance (g s), instantaneous carboxylation efficiency (k), and total chlorophyll index (Chl) exhibited a similar declining trend until the end of the maturation in both sugarcane varieties (Figure 2a–c, f). However, A i was higher in IACSP95-5000 compared to IACSP94-2094 in February (maximum growth) and in August (late maturation), as shown in Figure 2a. The g s followed a similar pattern of response as compared to A i, with the highest values found during the maximum growth and then a decreasing trend from April (early maturation) in both varieties (Figure 2b). However, IACSP95-5000 had higher g s than IACSP94-2094 during the maximum growth (October and January) period (Figure 2b). Both varieties exhibited decreases in k along the cycle, with the lowest values occurring during the maturation period (Figure 2c). Again, IACSP95-5000 exhibited higher k than IACSP94-2094 during the maturation (April, May, and July) (Figure 2c). While ФPSII in IACSP94-2094 showed a decreasing trend since the maximum growth, IACSP95-5000 presented a smaller variation of ФPSII along the cycle (Figure 2d). IACSP94-2094 displayed higher ФPSII than IACSP95-5000 in October and January, that is, during the maximum growth stage (Figure 2d). Both sugarcane varieties exhibited the lowest ФPSII values at late maturation. While IACSP95-5000 presented the highest LNC at the maximum growth and the lowest LNC at late maturation, non-significant variation of LNC was found in IACSP94-2094 along the crop cycle (Figure 2e). The total chlorophyll index decreased at the maturation for both varieties. However, IACSP94-2094 exhibited Chl values higher than IACSP95-5000 at the late maturation (Figure 2f).
Diurnal-integrated leaf CO2 assimilation (A i, in a), stomatal conductance (g s, in b), instantaneous carboxylation efficiency (k, in c), effective quantum yield of photosystem II (ΦPSII, in d), leaf nitrogen concentration (LNC, in e), and total chlorophyll index (f) of IACSP95-5000 and IACSP94-2094 during the maximum growth and early, medium, and late maturation periods. Symbols represents mean ± SE; LNC, n = 4, all others n = 8 (pooled data from leaves +1 and +3). *Statistical difference between varieties, and distinct lowercase letters indicate statistical difference among sampling times at P < 0.05.
Carbohydrate dynamics in leaves and stalks
IACSP95-5000 and IACSP94-2094 showed higher leaf concentrations of total soluble sugars at the late maturation when compared with the maximum growth period (Figure 3a). At the maximum growth, IACSP94-2094 showed higher (1.45-fold) total soluble sugars than IACSP95-5000; however, this pattern was inverted at late maturation (Figure 3a). Starch concentration in leaves of IACSP95-5000 was higher at late maturation than at the maximum growth, whereas IACSP94-2094 presented a significant decline at late maturation when compared to the maximum growth period (Figure 3b). Overall, IACSP94-2094 exhibited higher concentration of starch than IACSP95-5000 in both periods (Figure 3b). The sugarcane varieties exhibited similar leaf sucrose concentration during the maximum growth stage, but IACSP94-2094 presented higher sucrose concentration than IACSP95-5000 at the late maturation (Figure 3c). Both varieties exhibited higher leaf sucrose concentration at late maturation when compared to the maximum growth period (Figure 3c).
Concentration of total soluble sugars (a), starch (b), and sucrose (c) in leaves of IACSP95-5000 and IACSP94-2094 during the maximum growth and late maturation periods. Symbols represents mean ± SE; n = 4. *Statistical difference between varieties, and distinct uppercase letters indicate statistical differences among sampling times at P < 0.05. DW is dry weight.
Concerning the total soluble sugars along the stalk, the top portion of both varieties showed the lowest values at early maturation, with IACSP94-2094 exhibiting higher values than IACSP95-5000 (Figure 4a). At the middle of maturation period, there was non-significant change (P > 0.05) in total soluble sugars along the stalk of IACSP94-2094, while IACSP95-5000 showed higher concentration in medium stalk portions (Figure 4a). IACSP94-2094 showed decreases in the total soluble sugars in medium and bottom portions at the late maturation. At that time, IACSP94-2094 had higher concentrations of total soluble sugars than IACSP95-5000 in top (1.3-fold) and medium (1.25-fold) stalk portions (Figure 4a). Overall, sucrose concentration increased gradually from top to bottom portions at the early maturation (Figure 4b). During the medium maturation, the highest concentration of sucrose occurred in the medium stalk portion of IACSP95-5000 and in the bottom stalk portion of IACSP94-2094 (Figure 4b). Despite the stalk portion and maturation time, IACSP94-2094 exhibited higher sucrose concentration than IACSP95-5000 (Figure 4b).
Concentration of total soluble sugars (a) and sucrose (b) in stalks portions (top, medium, and bottom) of IACSP95-5000 and IACSP94-2094 during the early, medium, and late maturation periods. Values are mean ± SE; n = 4. *Significant difference between varieties, distinct uppercase letters indicate statistical difference between sampling times, and distinct lowercase letters among stalk positions (top, medium, and bottom) at P < 0.05. DW is dry weight.
Carbohydrate metabolism: leaf enzymatic activity
The SPS activity in leaves of IACSP95-5000 increased 1.9-fold at the late maturation when compared with the maximum growth stage, while an inverse pattern was found for IACSP94-2094 (Figure 5a). Conversely, SuSy activity did not vary (P > 0.05) between the maximum growth and the late maturation for both varieties (Figure 5b). Leaves of IACSP95-5000 presented higher SuSy activity (1.7-fold) than ones of IACSP94-2094 at the maximum growth (Figure 5b). Leaf SAI activity was similar between periods for IACSP95-5000, which presented higher values than IACSP94-2094. This latter presented a significant decrease of leaf SAI activity at the late maturation (Figure 5c). The leaf SNI activity decreased at the late maturation of both varieties, with IACSP94-2094 exhibiting higher SNI activity than IACSP95-5000 (Figure 5d).
Activity of sucrose phosphate synthase (SPS, in a), sucrose synthase (SuSy, in b), soluble acid invertase (SAI, in c), and soluble neutral invertase (SNI, in d) in leaves of IACSP95-5000 and IACSP94-2094 during the maximum growth and late maturation periods. Symbols represents mean ± SE; n = 4. *Statistical difference between varieties, and distinct uppercase letters indicate statistical difference among sampling times at P < 0.05. suc is sucrose.
Carbohydrate metabolism: stalk enzymatic activity
At the maximum growth stage, stalk SPS activity of IACSP94-2094 decreased from top to bottom. When compared to IACSP95-5000, stalk SPS activity of IACSP94-2094 was higher in top (ca. 1.6-fold) and medium (ca. 1.8-fold) portions (Figure 6a). At the late maturation, IACSP94-2094 had higher SPS activity than IACSP95-5000 in top and bottom stalk portions (Figure 6a). We measured the highest stalk SuSy activity in the bottom portion of IACSP95-5000 at the maximum growth, which presented higher SuSy activity than IACSP94-2094 (Figure 6b). At the late maturation, the highest SuSy activity was found in top portion of IACSP95-5000 and medium portion of IACSP94-2094 (Figure 6b). When comparing stalk portions at the maximum growth, the highest SAI activities were found in top portions of both varieties (Figure 6c). Stalk SAI activity in top and bottom portions was higher in IACSP94-2094 than in IACSP95-5000 (Figure 6c). IACSP95-5000 showed an increasing trend of SNI activity from top to bottom portions at the maximum growth. In bottom portion, SNI activity was 8.5-fold higher in IACSP95-5000 than in IACSP94-2094 (Figure 6d). Finally, the activities of SAI and SNI decreased at the late maturation in both varieties when compared to the maximum growth (Figure 6c, d).
Activity of sucrose phosphate synthase (SPS, in a), sucrose synthase (SuSy, in b), soluble acid invertase (SAI, in c), and soluble neutral invertase (SNI, in d) in stalks portions (top, medium, and bottom) of IACSP95-5000 and IACSP94-2094 during the maximum growth and late maturation periods. Values are mean ± SE; n = 4. *Statistical difference between varieties, distinct uppercase letters indicate statistical differences between sampling times, and distinct lowercase letters among stalk portions (top, medium, and bottom) at P < 0.05. suc is sucrose.
Plant growth: leaf area and above-ground biomass
IACSP95-5000 exhibited higher LAI than IACSP94-2094 during the maximum growth period, with IACSP95-5000 and IACSP94-2094 reaching the highest LAI in December and January, respectively (Figure 7a). Accordingly, IACSP95-5000 produced more biomass than IACSP94-2094 at the end of crop cycle, that is, about +31% and +28% on fresh and dry bases, respectively (Figures 7b and S2).
Leaf area index (LAI, in a) and total above-ground dry biomass (b) of IACSP95-5000 and IACSP94-2094 during the maximum growth and early, medium, and late maturation periods. Values are mean ± SE; n = 4. *Statistical difference between varieties, and distinct lowercase letters indicate statistical difference among sampling times at P < 0.05.
Discussion
Here, we compared two sugarcane varieties with differential biomass production under field conditions. While IACSP95-5000 exhibited higher biomass production (Figures 7b and S2), IACSP94-2094 accumulated more sucrose per unit of stalk mass (Figure 4b). These differences can be explained by the carbohydrate metabolism of stalks and leaves (Figure 8). Notably and despite those contrasting strategies, both varieties displayed similar final sugar yield and then low stalk sucrose content in IACSP95-5000 was compensated by increased biomass production. Such compensatory mechanism was based on several metabolic adjustments for optimising resource allocation and promote high sugar yield.
Scheme of sucrose accumulation and synthesis in leaves and stalks of IACSP94-2094 (a) and IACSP95-5000 (b) during the maximum growth and maturation periods. Arrow thickness denotes the intensity of responses when comparing varieties, with red arrows indicating sucrose degradation and green arrows indicating sucrose synthesis. Grey circles represent sucrose; Suc: sucrose; Hex: hexose; SPS: sucrose phosphate synthase; SuSy: sucrose synthase; SAI: soluble acid invertase; SNI: soluble neutral invertase.
Underlying mechanisms leading to low photosynthesis at maturation: a common response to ageing
Photosynthetic rates decreased while stalk sucrose levels increased (Figures 2a and 4b). Decline in photosynthesis of maturing sugarcane plants occurs with increasing leaf sugar concentrations (McCormick et al., Reference McCormick, Cramer and Watt2008a), with such build-up of leaf sugar being a possible consequence of impaired phloem loading and then reduced transport to sinks (stalks). Based on the source–sink relationship, one would expect stimulation of photosynthetic activity by increasing sink demand (Ribeiro et al., Reference Ribeiro, Machado, Magalhães Filho, Lobo, Martins, Silveira, Yin and Struik2017), that is, sucrose accumulation in stalks. However, this did not happen here and there is evidence that other exogenous (environment) or endogenous factors are limiting sugarcane photosynthesis at the end of plant cycle (De Souza et al., Reference De Souza, Grandis, Arenque-Musa and Buckeridge2018). Taking into account the environment, plants were clearly facing water deficit and low temperature at the maturation stage (Figure 1b) and such limiting condition is known to reduce stomatal conductance and photosynthesis (Sales et al., Reference Sales, Marchiori, Machado, Fontenele, Machado, Silveira and Ribeiro2015; Cerqueira et al., Reference Cerqueira, Santos, Marchiori, Silveira, Machado and Ribeiro2019).
Here, our data suggest that photosynthesis is reduced during maturation due to stomatal and non-stomatal factors (Figure 2b, c). Considering stomatal aperture, previous studies revealed that sucrose accumulation in the guard cells affects the stomatal dynamics and then photosynthesis (Kelly et al., Reference Kelly, Moshelion, David-Schwartz, Halperin, Wallach, Attia, Belausov and Granot2013; Daloso et al., Reference Daloso, Antunes, Pinheiro, Waquim, Araújo, Loureiro, Fernie and Williams2015). In C4 plants, most of nitrogen is invested in the photosynthetic enzymes Rubisco and PEPC (Tazoe et al., Reference Tazoe, Noguchi and Terashima2006), and then low LNC would justify decreases in k (Tofanello et al., Reference Tofanello, Andrade, Flores-Borges, Kiyota, Mayer, Creste, Machado, Yin, Struik and Ribeiro2021) and also in total chlorophyll index with ageing, but only in IACSP95-5000 (Figure 2e, f). Even with non-significant change in LNC of IACSP94-2094 as maturation progressed, we noticed a decrease in CO2 assimilation (Figure 2a, e). As a possible explanation, N allocation likely changed during the plant cycle and more N was driven to the synthesis of proteins not related to photosynthesis at latter phenological stages of IACSP94-2094. In addition to the water deficit and low temperature, high leaf sucrose content would be an additional factor downregulating photosynthesis (Ribeiro et al., Reference Ribeiro, Machado, Magalhães Filho, Lobo, Martins, Silveira, Yin and Struik2017).
We would argue that low Chl (Figure 2f) led to low ΦPSII as maturation advanced, mainly for IACSP94-2094 (Figure 2d). Interestingly, reduction in Chl during the early and medium maturation of IACSP95-5000 did not result in low ΦPSII (Figure 2d, f). In fact, photochemical activity is not defined solely by Chl (Silveira et al., Reference Silveira, Seabra, Marcos, Pelegrino, Machado and Ribeiro2019) and decreases in photochemical activity were reported with advancing maturation, when sugarcane plants were facing water deficit (De Souza et al., Reference De Souza, Grandis, Arenque-Musa and Buckeridge2018). Here, we found evidence that plants were under water deficit only at the late maturation period, when irrigation was stopped to induce sugarcane maturation (Figure 1b). However, our data on leaf water potential did not reveal this (Figure S1). For instance, the highest and lowest diurnal-integrated CO2 assimilation were noticed with ψ around −1.2 MPa (Figures 2a and S1). Then, low temperature seems to have an important role during the maturation period (Figure 1a), limiting photosynthesis in field-grown plants under subtropical conditions. Previously, we found sugarcane plants are sensitive to low temperature, with reduced photosynthetic rates due to stomatal, photochemical, and biochemical (low Rubisco activity) limitations (Cerqueira et al., Reference Cerqueira, Santos, Marchiori, Silveira, Machado and Ribeiro2019).
Taken together, these findings suggest that the reduction in photosynthetic rates as sugarcane plants mature is due to ageing-related changes as an endogenous factor and also due to low temperature, both inducing decreases in carboxylation, photochemical activity, and stomatal conductance (Figure 2). Both varieties exhibited similar photosynthetic dynamics throughout the experimental period (Figure 2a), but IACSP95-5000 presented higher CO2 assimilation (Figure 2a) and higher LAI (Figure 7a) than IACSP94-2094. As consequence, IACSP95-5000 produced more biomass than IACSP94-2094 (Figures 7b and S2). On the other hand, IACSP94-2094 accumulated more sucrose in stalks as compared to IACSP95-5000.
Stalk sucrose accumulation: an interplay between leaves and stalks
When considering biomass production (Figures 7b and S2) and sucrose accumulation (Figure 4b), differences between IACSP95-5000 and IACSP94-2094 can be explained by the carbohydrate metabolism of stalks and leaves (Figure 8). During the period of maximum growth, IACSP94-2094 showed high SPS activity (Figures 5a and 8) and low SAI activity in leaves (Figures 5c and 8). In contrast, leaves of IACSP95-5000 exhibited high SAI and SuSy activities (Figures 5b, c and 8). While increased leaf SPS activity promotes sucrose synthesis (Chandra et al., Reference Chandra, Verma, Islam, Grisham, Jain, Sharma, Roopendra, Singh, Singh, Verma and Solomon2015), low leaf SAI activity reduces sucrose degradation (Anur et al., Reference Anur, Mufithah, Sawitri, Sakakibara and Sugiharto2020). In IACSP95-5000, high leaf activities of SAI and SuSy likely accelerated sucrose hydrolysis (McCormick et al., Reference McCormick, Cramer and Watt2008a; Anur et al., Reference Anur, Mufithah, Sawitri, Sakakibara and Sugiharto2020; Mason et al., Reference Mason, Hoang, Botha, Furtado, Marquardt and Henry2023), a way to supply hexoses for growth respiration and then support cell division and expansion. Accordingly, leaf area was higher in IACSP95-5000 than in IACSP94-2094 (Figure 7a). Increased sucrose degradation is also an alternative to avoid the downregulation of photosynthesis induced by high leaf sucrose concentration (Ribeiro et al., Reference Ribeiro, Machado, Magalhães Filho, Lobo, Martins, Silveira, Yin and Struik2017; Anur et al., Reference Anur, Mufithah, Sawitri, Sakakibara and Sugiharto2020). Again, photosynthetic activity in IACSP95-5000 was significantly higher than that in IACSP94-2094 (Figures 2 and 5c).
In sink organs such as stalks, IACSP95-5000 had high activity of SuSy and SNI (Figures 6b, d and 8), two key enzymes playing an important role in controlling the rate of sucrose hydrolysis, mainly in immature tissues (Rossouw et al., Reference Rossouw, Bosch, Kossmann, Botha and Groenewald2007, Reference Rossouw, Kossmann, Botha and Groenewald2010). SuSy operates primarily towards the sucrose degradation, which is cleaved into fructose and UDP-Glu and then used in respiration and polymer (starch or cell wall constituents) biosynthesis (Wang et al., Reference Wang, Nayak, Koch and Ming2013). For instance, low SNI activity reduces sucrose mobilisation and impairs growth in sugarcane plants due to low availability of hexoses for respiration (Rossouw et al., Reference Rossouw, Bosch, Kossmann, Botha and Groenewald2007). As compared with IACSP95-5000, stalks of IACSP94-2094 showed lower SNI activity, but higher SPS and SAI activities (Figures 6a, c, d and 8). According to You-Qiang et al. (Reference You-Qiang, Hai-Ling and Yang-Rui2009), sugarcane varieties with high sucrose accumulation exhibit higher SPS activity throughout the development stages. As stalk SAI activity provides substrates for supporting the growth of immature and expanding tissues in sugarcane (Rossouw et al., Reference Rossouw, Bosch, Kossmann, Botha and Groenewald2007), a greater capacity for sucrose accumulation at maturation is expected in varieties presenting high SAI activity, as IACSP94-2094. In addition, increased SAI activity may contribute to the remobilisation of sucrose stored in vacuoles (Liu et al., Reference Liu, Zhang, Gong, Li, Pan, Davis, Jing, Wu and Godwin2021).
Low amounts of soluble sugars in leaf tissues may indicate high sucrose transport rates to sinks rather than low photosynthesis (Mason et al., Reference Mason, Furtado, Marquardt, Hodgson-Kratky, Hoang, Botha, Papa, Mortimer, Simmons and Henry2020). At the maturation period, IACSP94-2094 presented high SPS activity and low SAI activity in stalks (Figures 6a, c and 8). In principle, these metabolic adjustments would support a greater accumulation of sucrose. In contrast, IACSP95-5000 showed an increased activity of SNI in stalks (Figures 6d and 8). Overall, the activity of SNI decreased significantly at late maturation in both varieties (Figures 6d and 8), but no impact is expected as glycolytic flux is significantly reduced at this time (Rossouw et al., Reference Rossouw, Bosch, Kossmann, Botha and Groenewald2007). Furthermore, decreased SNI activity in stalks is compensated by increased SuSy activity (Rossouw et al., Reference Rossouw, Kossmann, Botha and Groenewald2010).
Our data revealed the underlying physiological mechanisms responsible for differential biomass production between the sugarcane varieties studied (Figure 8). The high-yielding variety – IACSP95-5000 – used energy and carbon skeletons to produce more biomass, which was linked to increased enzymatic degradation of sucrose in leaves and stalks. On the other hand, IACSP94-2094 demonstrated a more efficient strategy for sucrose accumulation in stalks (Figure 8). Such higher efficiency was attributed to a higher activity of SPS in leaves and stalks during the maximum growth and maturation, as well as low activity of SAI in leaves during the maximum growth and in stalks during the maturation (Figure 8). This pattern would maximise the sucrose flux to stalks and its consequent accumulation in IACSP94-2094. As mentioned before, both varieties displayed similar total sugar yield, varying between ∼34 Mg ha–1 in IACSP95-5000 and ∼29 Mg ha–1 for IACSP94-2094 with such difference being non-significant (t test; P > 0.05). Our findings indicate a compensatory mechanism between biomass production and sucrose accumulation in sugarcane, involving several metabolic adjustments for optimising resource allocation and promote high sugar yield. From a broad perspective, a potential failure in such compensatory mechanism between biomass production and sucrose accumulation or even a change in resource (carbon and nitrogen) allocation could explain the declining trend of cane and sucrose yield ratoon after ratoon, with important consequences for sugarcane management and industry.
Agronomic and industrial perspectives: maximising biomass vs. sucrose yield
Here, IACSP94-2094 showed higher and faster sucrose accumulation in stalks than IACSP95-5000, as shown here (Figures 4, 8) and evidenced by soluble solids (oBrix, data not shown). Such characteristics would favour early harvesting of IACSP94-2094 (Singh et al., Reference Singh, Pathak, Singh, Mishra and Sharma2017). Additionally, the need for ripeners would be reduced in IACSP94-2094 as compared to IACSP95-5000, an interesting benefit for managing sugarcane fields. Besides the environmental impact (De Almeida et al., Reference De Almeida, Véliz, Sartori and Santos2022), the use of ripeners can reduce ratoon sprouting, affecting sugarcane regrowth and longevity (Mehareb et al., Reference Mehareb, Abou-Elwafa and Galal2016).
Considering an industrial perspective, the choice of varieties with high biomass production or high sugar yield depends on the purpose. For conventional first-generation ethanol production, one would prefer varieties with high sucrose concentration in stalks (De Almeida and Colombo, Reference De Almeida and Colombo2023), while the best options for the second-generation ethanol would be sugarcane varieties or other species with high biomass production, such as energy cane (Dias et al., Reference Dias, Junqueira, Jesus, Rossell, Maciel Filho and Bonomi2012; Cruz et al., Reference Cruz, Pacheco, Silva, Almeida, Miranda, Pissolato, Machado and Ribeiro2021). As lower sucrose concentration is compensated by higher biomass production, IACSP95-5000 would be a better option than IACSP94-2094 when the aim is the first (from sucrose) and second (from bagasse) generation ethanol. Industrial capacity is an important aspect for such double-purpose varieties, a discussion beyond the scope of this paper. On the other hand, IACSP94-2094 is rustic and less sensitive to limiting conditions, such as water deficit (Ribeiro et al., 2003; Contiliani et al., Reference Contiliani, Nebó, Ribeiro, Landell, Pereira, Ming, Figureira and Creste2023). As biomass production is limited not only by endogenous factors but also by environmental ones, IACSP94-2094 would be an interesting sugarcane variety to supply industry when growing areas are rainfed and present a large seasonal variation of water availability.
Supplementary material
The supplementary material for this article can be found at https://doi.org/10.1017/S0014479724000061
Acknowledgements
This research was supported by the São Paulo Research Foundation, Program for Research on Bioenergy (BIOEN/FAPESP, Brazil, Grant #2008/57495-3), which also provided scholarships to TSM and DFSM (Grants #2022/09154-0 and #2014/12969-9). PERM, ECM, and RVR are fellows of the National Council for Scientific and Technological Development (CNPq, Brazil, Grants #312663/2021-8, #311345/2019-0 and #304295/2022-1). PERM and JRMF received scholarships from CNPq (Grants #141675/2014-4 and #140296/2011-5). LPC is a fellow of the Brazilian Agave Development Program (BRAVE, Brazil, Research Agreements #5902 and #5903).
Competing interests
The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.
