Physiological response as a tolerant mechanism to Yellow Sugarcane Aphids (YSA) (S. flava) herbivory on selected commercial sugarcane varieties (Saccharum officinarum)

doi:10.21203/rs.3.rs-4287978/v1

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Physiological response as a tolerant mechanism to Yellow Sugarcane Aphids (YSA) (S. flava) herbivory on selected commercial sugarcane varieties (Saccharum officinarum)

https://doi.org/10.21203/rs.3.rs-4287978/v1

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Physiological tolerance in response to Yellow Sugarcane Aphid feeding remains an unexplored area in the sugar industry of Zimbabwe and elsewhere. A 7 × 2 factorial in a complete randomized block design (CRBD) replicated four times was used, with seven sugarcane varieties (00-1165, ZN 3L, ZN 8, ZN 9, 96-1107, N14 and ZN 10) under two treatments of aphid infestation (un-infested (control) and infested). Results indicated that there were highly significant differences (p< 0.001) amongst the sugarcane varieties on chlorophyll content and percentage chlorophyll loss in control (sprayed) and infested (unsprayed) plots. Summer results recorded a great increase in percentage chlorophyll loss (21.4%) margin scored on ZN 10. Findings of the regression analysis displayed a highly significant (p< 0.001) strong positive correlation (r= 0.85) between chlorophyll loss and aphid number. Summer results showed highly significant differences (p< 0.001) on gas exchange responses in control and infested plots. Nevertheless, in YSA infested plots, 00-1165 recorded the highest compensatory photosynthetic rate (32.52), transpiration rate (4.32), stomata conductance (218.2) when compared to the least obtained from ZN 10 and N14 at day 28. Obtained results of the regression analysis presented highly significant positive correlation between; chlorophyll loss and photosynthesis (r= 0.44), photosynthesis and aphid number (r= 0.57). Ranking of sugarcane varieties was done according to YSA susceptibility; less susceptible (00-1165), moderate susceptible (ZN 3L, ZN 8 and ZN 9) and highly susceptible (96-1107, N14 and ZN 10). Sugarcane growers should add 00-1165 sugarcane variety amongst the existing sugarcane varieties as it is highly tolerant to YSA damage as proven by physiological compensatory and maintaining behavior.

compensatory

photosynthetic

transpiration

stomata conductance

tolerant

Sugarcane is vital crop that produces sugar, ethanol, and other byproducts including molasses, filter cake, and bagasse globally (Clowes and Breakwell 1998; Esterhuzein 2012; Shabani et al. 2020). Sugarcane has become a subject of damage to persistent incursion of Yellow Sugarcane Aphids (S. flava) in sugarcane fields. Feeding by yellow sugarcane aphids in sugarcane has been reported to have negative effects on stalk number and stalk height (Hall,2001), biomass yield (Hall 2001; Madiope et al. 2021) and yield (Miskimen 1970, Wilson 2019).

Yellow Sugarcane Aphids (YSA) (Sipha flava) are relatively small, less than 2 mm in diameter, light in color and have abundant short stiff hairs covering the head, thorax and abdomen (Wilson 2019). Winged (alate) forms are distinguished from wingless (apterous) forms by their bright yellow abdomen. Climatic conditions allow the YSA to exhibit two modes of reproduction; non-mating (parthenogenetically) in warmer climates (Halbert et al. 2013), this result in proliferation and mating, where live individuals are produced when females mate with males in cold winters as low temperatures induce ovivipary (Nuessly 2005; Way et al. 2014), The contribution of climate to YSA damage in winter and summer will be considered because heavy infestation and persistent feeding on young plant result in leaves turning from yellow to red followed by premature leaf senescence and death of the stems (Nuessly and Hentz 2002; Way et al. 2015; Wilson 2019). Breen and Teetes (1986) as cited in Dumont et al. (2023) highlighted that red to purple discoloration and premature leaf necrosis is as a result of injection of saliva by YSA resulting in leaf chlorosis. Moreover, Akbar et al. (2010) reported that leaf discoloration is associated with a chlorophyll content decline associated with a drastic decrease in the photosynthetic rate (White 1990).

Toxins injected in saliva contain toxic oxygen which may cause biochemical changes within the crop resulting in bleaching of leaves and increased metabolism (Macedo et al. 2003). The saliva contains a variety of effector proteins, hydrolytic enzymes, and toxic chemicals that cause plants to perceive the invasion of aphids and may potentially increase the buildup of reactive oxygen species (ROS) (Shankar and Yinghua 2021).

Photosynthesis stops when chlorophyll leaf pigments, mainly chlorophyll a and b, are reduced, which also affects the leaf's general health (Green and Durnford 1996; Jensen 2000; Taiz and Zeiger 2010, Craigie 2022). Nitrogen is required in the formation of chlorophyll while magnesium give structural integrity to the chlorophyll molecule. Gonzales et al. (2002) reported ultrastructural damage to chloroplasts in leaves of Johnson grass (Sorghum halepense L.) infested with S. flava, results of light microscopy showed increased chloroplast volume, loss of grana structures and aggregation of starch granules. However, high rates of fecundity of aphids were linked to plants with high nitrogen contents (Mahmoud's 2005; El-Rawy et al. 2007). This clearly shows nitrogen encourages the leaf to become soft and allow easier probing of the aphid stylet into phloem sieves. Additionally, Slman (1997) noted that the density of cereal aphids responded favorably and significantly to increased nitrogen rates and their interactions with P₂ O₅.

Yellow Sugarcane Aphid is being managed by chemical, biological, and cultural techniques. In Zimbabwe, Actara 25 Wettable Granules (WG) (25% thiamethoxam and ACETA (Allice) are registered for use. Way et al. (2015) observed the existence of biological agents such as earwigs (Dermaptera), hover flies (Syrphidae), ladybird species (Coccinellidae), spiders (Araneae), and ants (Formicidae) as natural enemies. Several authors (Nuessly 2005; Nuessly et al. 2010; Gallun et al. 1966; Roberts et al. 1979; Roberts and Foster 1983; Sosa 1990; White 1990; ZSAES unpublished) suggested that resistant sugarcane cultivars can be used for S. flava cultural management. Aphid resistance may develop as a result of ongoing usage of insecticides belonging to the neonicotinoid, pyrethroid, and organophosphate classes; however, such research in Zimbabwe's sugar industry is lacking. Studies examining physiological crop tolerance as a host plant defense mechanism against YSA are necessary in light of this and required for integration into existing comprehensively Integrated Pest Management (IPM) strategies. This strategy is environmentally friendly and is compatible with current management strategies. Physiological tolerance can be defined as the ability of plants to withstand insect injury through physiological and biochemical compensation (Painter 1951). Tolerant plants can maintain or increase physiological mechanisms (chlorophyll content and gas exchange) in response to insect herbivory as reported by various authors (Akbar 2009; Koch 2015; Mbulwe 2017; Paudyal, 2019; Paudyal et al. 2020).

Peterson and Higley (2001) and Gordy et al. (2019) indicated that understanding the physiological changes resulting from aphid feeding is an important step in developing accurate economic damage levels for different crop cultivars. Much focus of physiological tolerance studies has been directed on sugarcane aphid (Melanaphis sacchari) (Singh et al. 2004, Akbar, 2009; Armstrong et al. 2015, 2017; Bowling et al. 2016a,b; Mbulwe et al. 2016; Mbulwe 2017; Paudyal 2019; Paudyal et al. 2020) in sorghum and YSA in sorghum (Akbar 2009). Existing conclusions were made that the aforementioned aphid induced tolerance compensatory mechanisms such as increased photosynthesis, transpiration and stomata conductance in tolerant varieties while in susceptible genotypes, gaseous exchange processes were drastically reduced by sugarcane aphid. This mechanism if captured and utilised enhances sugarcane productivity in the sugar industry. However, limited studies have reported on how sugarcane crop can exhibit tolerance to YSA feeding. Thus, this study will evaluate how sugarcane plant may physiologically tolerate YSA infestation. This will be an interesting and worthy research avenue in sugarcane in the presence of YSA feeding as suggested by Macedo et al. (2003, that phloem feeders such as aphids can dramatically decrease host plant photosynthesis, while sap feeders such as scale insects can cause an increase in leaf assimilation rate (Retuerto et al. 2004). An increase in photosynthesis after herbivory is interpreted as a plant strategy to compensate for the effect of the herbivore (Trumble et al. 1993). This hypothesis has driven the need for such a study to conclusively document the existence of such mechanism among sugarcane varieties in response to YSA feeding.

Studies have indicated that aphids have the potential to alter host plant physiological processes such as photosynthesis, stomata conductivity, respiration and chlorophyll content (Warrington et al. 1989; Welter 1989; Macedo et al. 2003; Peterson et al. 2004; Aldea et al. 2005; Delaney and Higley 2006; Shannag 2007; Paudyal 2019) and several authors (Capinera 1981; Ryan et al. 1987; Meyer and Whitlow 1992; Larson 1998; Haile et al. 1999; Macedo et al. 2003a; Diaz-Montano et al. 2007; Pierson et al. 2011; Paudyal 2019) emphasized that this change ultimately affects plant growth, development and yield. Overreliance of symptoms caused by YSA feeding has been the major talk by farmers hence side-lining the science behind contributing to the symptoms. Therefore, this research seeks to document physiological tolerance data to fill that gap in order to predict present and future damage posed by prolonged YSA feeding. Sugarcane productivity will be enhanced by identifying tolerant varieties resilience to YSA infestation.

Welter (1989) emphasized that increased rates of transpiration caused by leaf-eating insects result in disruption of leaf integrity. This proves a compensatory behaviour which gives plants a competitive advantage to carry out their physiological processes without being disadvantaged by insect pests. In support of this, Hoad et al. (1998) pointed out that this creates the potential for uncontrolled water loss through cut edges or abraded cuticles. However, it is the essence of this study to investigate such proposed processes so as to build concrete literature that pertains to sugarcane. Numerous authors (Peterson and Higley 1993; Macedo et al. 2003; Shannag, 2007) have declared that gas exchange is the primary process governing plant growth, development, and ultimately fitness adequate characterization of photosynthesis, respiration, and water vapor is required. This study seeks to understand how YSA injury affects these primary metabolic parameters so as to develop general models of plant response as Peterson (2001) suggested.

A number of studies (Miller et al. 1994; Deol et al. 2001; Diaz-Montano et al. 2007; Limaje et al. 2017) have reported the effects of aphid feeding on the physiological response of plants and found reduced chlorophyll content of plants. Chlorophyll is major driver for photosynthesis to occur in crop. Therefore physiological tolerance is a requirement for sugarcane varieties to compensate for YSA injury so as to maintain physiological integrity. In addition to this, Haile et al. (1999) and Frazen et al. (2007) described that aphids have the potential to directly reduce photosynthesis, either by preventing the normal flow of nutrients or by blocking the opening of the stomata and thus limiting CO₂ uptake due to honeydew accumulation on the leaves. In contrast, Nagaraj et al. (2002) suggested that the reduction in chlorophyll may not indicate a net reduction in the rate of photosynthesis hence the need for further studies to confirm this. Therefore, this study will also explore and fill such a gap. Moreover, Retuerto et al. (2004) and Gutsche et al. (2009) indicated that some plants have the ability to compensate the rate of photosynthesis in response to insect invasion. However, phloem feeders such as aphids demonstrate the ability to strongly and actively reconfigure leaf metabolism via effectors (Giron et al. 2018), which can defeat the plant's mitigation strategy.

The physiological mechanisms underlying indirect damage to leaf arthropods through transpiration, stomata conductance and other physiological processes are not well understood as mentioned in literature (Hunter 2001; Peterson et al. 2004). There is a vacuum on physiological tolerance of sugarcane varieties as more studies were directed other aphid species and sugarcane aphid in sorghum (Singh et al. 2004; Akbar 2009; Armstrong et al. 2015, 2017; Bowling et al. 2016a,b; Koch 2015; Mbulwe et al. 2016; Mbulwe 2017; Paudyal 2019; Paudyal et al. 2020) in other cereal crops. The above mentioned studies conclusively indicated that aphids reduced chlorophyll content and gaseous exchange process in susceptible varieties. On the other hand, tolerant varieties were able to compensate, maintain/ increase chlorophyll content and gas exchange responses in response to insect feeding.

Entomologists, plant physiologists, and plant breeders have long been intrigued by the interactions between plants. The mechanisms behind plant responses have not received much attention in research (Macedo et al. 2003). Moreover, a large body of research has concentrated on how plants react to insect herbivory, but it has not thoroughly examined the biochemical and physiological reactions of the insect (Peterson and Higley 2001). Approximating the quantity of the insect population and visualizing plant damage in order to compare it to a known susceptible genotype or cultivar are the most fundamental ways of screening for tolerance. Since insect populations are similar to those of susceptible plant kinds, populations can grow to extremely high levels, and visual damage assessments using traditional phenomic screening can introduce bias and limit precision. There is need for integrating the use of phenotypic approaches which are not labor intensive such as using SPAD and CIRAS-4 portable photosynthetic system devices to determine tolerance. The use of plant metabolite assays, insect population assays, hand-held spectrophotometry (SPAD meter) to measure chlorophyll content in leaves, enzyme-linked immunosorbent assays (ELISAs) to measure virus transmission, and the EPG technique to measure feeding behavior are some of the methods that have received support from a variety of authors (McLean and Kinsey 1964; Tjallingii 1988; Deol et al. 1997; Girma et al. 1998; Walker 2000; Chan et al. 2010; Chen et al. 2012; Ménard et al. 2013; Koch et al. 2016).

To comprehend piercing sucking insects like YSA, a thorough approach is necessary. Such studies are a prerequisite in yield prediction due to YSA damage at early stages of growth (≤ 3months) (Wilson 2019). To date, hardly any study has reported an association between YSA herbivory, chlorophyll content loss and gas exchange response in response to YSA incursion. This knowledge of how YSA affects sugar cane physiology, particularly in resistant and susceptible lines, may help explain the physiological mechanisms underlying tolerance. The purpose of these experiments is to gain insight into the physiological responses of selected resistant and susceptible sugarcane genotypes to YSA feeding so as to predict yield losses at early stages of growth. The specific objective is to describe the physiological effects of YSA feeding on chlorophyll content and gas exchange responses (photosynthetic, stomata conductance and transpiration rates) under natural infestation conditions.

2.1 Study Site

The research was carried out in 2023/2024 season at the Zimbabwe Sugar Association Experiment Station, Masvingo Province, in the southeast Lowveld of Zimbabwe. The Zimbabwe Sugar Association Experiment Station (ZSAES) is found in agro-ecological region V which is characterized by very low and erratic rainfall of less than 500 mm per annum. ZSAES is situated on a 99km peg along the Ngundu-Tanganda road. It is located 430 m above sea level at latitude of 20^o01’ S and longitude 28^o 38’ E. The temperatures are very high especially during the summer season, particularly in the months of September, October and November with an average temperature of 34 ^oC. Low temperatures are recorded between May to July with the daily mean temperature of 25 ^oC.

2.2 Selection of varietal entries

A total of seven sugarcane varieties were selected and grown according to the Zimbabwe Sugar Production Manual (Clowes and Breakwell 1998). Selection of experimental material (varieties) was done by considering moderate level of resistance, susceptibility, previous cane yield and (Estimated Recoverable Crystal (ERC) %) based on recommendations from the Plant Breeding and Entomology Departments. Two preleased varieties namely; 00-1165 (low susceptible) and 96-1107 (highly susceptible) were selected. Moderate resistant varieties based on visual assessments; ZN 8, ZN 9 and ZN 3L, hence ZN 8 and ZN 9 have the ability to recover from damage inflicted by YSA. Lastly, ZN 10 and N14 were chosen despite being susceptible because of their high biomass and high estimated recoverable crystal (ERC %), hence they are leading in terms of area under production in both out-grower farmers and estates. Plant cane and ratoon cane will be assessed for chlorophyll content, physiological tolerance and aphid number. Cut back of sugarcane was done on the 22^th of September 2023 in order to mark the start of the new season in summer. Figure 1 shows field planting, established and cut back sugarcane crop.

2.3 Experimental design

A 7 × 2 factorial design in a complete randomized block design (CRBD) was used, replicated four times. Genotype was the first factor under seven levels (00-1165, 96-1107, ZN 10, ZN 8, ZN 9, ZN 3L and N14) and Aphid treatment was the second factor under two levels (un-infested (control) and infested).

2.4 Data collection

Data was collected on aphid number, chlorophyll content and gas exchange responses (photosynthesis, transpiration and stomata conductance).

2.4.1 Aphid number

Aphid assessments were done weekly on five selected primary tillers spaced 1 meter from each other on a row length of 15 meters. Tillers were randomly marked with a string on control (un-infested/sprayed) and aphid infested plots. These tillers were used for taking measurements throughout the experiment period. 40 days after natural YSA infestation, aphids were physically counted per plant/tiller on all leaves and summed so as to determine aphid number for winter and summer seasons.

2.4.2 Measurement of photosynthetic parameters

Photosynthetic parameter under investigation was chlorophyll content. However, gas exchange responses such photosynthesis, transpiration and stomata conductance were also determined. Taking of readings was done 60 days after planting on marked tillers for both control and YSA infested plots as discussed in detail.

2.4.2.1 Chlorophyll concentration

A chlorophyll meter (Model SPAD-502, Minolta Camera Co., Osaka, Japan) was used to measure the chlorophyll content in the sugarcane leaves (Figure 2). Handheld spectrophotometry (SPAD meter) is a portable device that absorbs light at wavelengths between 430 and 750 nm when passed over a leaf and estimates the chlorophyll content (Wood et al. 1992). Three readings from each leaf were taken for YSA un-infested (control) and infested (unsprayed) plots. These readings were averaged and a SPAD chlorophyll index (% chlorophyll loss) was calculated using the mean SPAD based on the formula: (C-I)/C modified from the one proposed by Deol et al. (1997); Akbar (2009) and Paudyal (2019), where C is the SPAD measurement from the YSA un-infested and I is the SPAD measurement from YSA infested sugarcane leaves. Golawska et al. (2010) indicated that SPAD measurements are reliable, quick and non-destructive for studying chlorophyll build up rates in different parts of the same leaf and also in determining of insect-plant interactions.

2.4.2.2 Gas exchange

Gas exchange responses were measured from YSA infested and un-infested (control) sugarcane leaves for all seven genotypes using a portable photosynthetic system (CIRAS-3 DC CO /H O Gas Analyzer) after every two weeks (Figure 3). The CIRAS photosystem was equipped with a LED chamber (1.75 cm²) so as to cover the leaf lamina excluding the midrib. The photosynthetic parameters were measured 60 days after natural infestation in both experimental plots. The collected data also included net photosynthesis rate (A. 𝜇molm^-2 s^-1), transpiration rate (E. mmolm^-2 s^-1), and stomatal conductance (gs, mmolm^-2 s^-1). The A equals the rate of photosynthetic (CO2 fixation minus the rate of CO2 loss) during respiration. Stomatal conductance is the rate at which CO₂ enters or water vapor escapes through the stomata, and internal CO2(Ci. Pascals) is the concentration of carbon dioxide inside the leaf (Meyer and Whitlow 1992; Paudyal 2019). Desiccants (soda lime, molecular sieve and drierite) were changed when 50 % of the colour changes. Every time, when going into the field to take measurements, the CO2canister was changed. Measurements were taken on the uppermost fully expanded total visible dew lap (TVD) leaf of aphid-infested and control plants between 11:00 and 14:00 h (CST) on days with full sunlight at a light intensity of 1200 umol photons m^-2 s^-1 and a reference CO₂ of ±400 ppm generated from a 12g CO2 cylinder connected to the meter. The TVD leaf was selected because it is the photosynthetic active leaf which utilises all the nutrients in the plant. The machine was switched on and allowed to warm up to a temperature of 55 ^oC before taking measurements. Prior to taking measurements, aphids were removed from the leaf by a fine brush so that they don’t interfere with results. Sugarcane leaves of control (un-infested) plants were also brushed to avoid undesirable effect resulting from the use of brush on the leaf tissues of aphid-infested plants. Measurements were taken in winter and summer of 2023/2024 season in both plant and first ratoon sugarcane (≤ 3 months).

2.5 Data analysis

For each measurement, GenStat 18^th version was used for analysis to examine differences in chlorophyll content, net photosynthesis rate, stomata conductance and water use efficiency. Parameters were analysed by a 7 (sugarcane genotypes) × 2 (YSA infested and control (sprayed) replicated four times. Means were separated using Fisher's Protected Least Significance at 5%. The relationship between: chlorophyll loss and aphid number, photosynthesis and chlorophyll loss, photosynthesis and aphid number, photosynthesis and SPAD values were determined by using regression analysis. The regression analysis was performed in order to integrate variable contribution to physiological tolerance among the sugarcane varieties. If the aphid number negatively affects chlorophyll content and percentage chlorophyll loss it reflects that it has an impact on photosynthesis. The loss in chlorophyll content is an indicator of feeding by aphids which interfere with nutrition components such as nitrogen and magnesium required in chlorophyll making and stability respectively. Moreover, aphid number and gas exchange responses correlation will have a significant effect on crop physiology. Furthermore, the regression analysis was valid in that it seeks to prove if there is a strong or weak correlation so as to effectively conclude that physiological tolerance in sugarcane is influenced by multiple gene expression as induced by YSA feeding on chlorophyll content, photosynthesis, transpiration and stomata conductance. This reflects that no independent variable can work alone in contributing to physiological tolerance.

2.6.1 Aphid number per plant 40 days after YSA natural infestation in winter and summer seasons

Results indicated that there were highly significant differences (p< 0.001) on number of aphids per plant in control and infested treatments. In winter, ZN 10 scored the highest aphid number (104) while 00-1165 recorded the least (31) in the control treatment (Figure 4). However, ZN 9 and ZN 3L were not significantly different (p> 0.05) from each other likewise ZN 8 and 96-1107. In aphid infested treatments, ZN 10 scored the highest aphid number (295) while 00-1165 scored the least. However, ZN 3L and ZN 9 were not significantly different from each other so as ZN 8 and 96-1107. In summer, in the control treatment, ZN 10 sugarcane variety recorded the highest (219) aphid number when compared to the least (74) realised by 00-1165 (Figure 1). In aphid infested plots, ZN 10 sugarcane variety scored the highest (962) aphid number while 00-1165 (237) recorded the least. Table 1 shows the temperature regimes and dates at which aphid numbers were recorded.

2.6.1 Effects of YSA on Chlorophyll content and percentage chlorophyll loss in selected sugarcane varieties

Results indicate that there were highly significant differences (p< 0.001) amongst the sugarcane varieties on chlorophyll content and percentage chlorophyll loss. In winter, in all the days of SPAD reading, ZN 3L scored the highest SPAD units (48.31) whilst N14 scored the least (43.34) at day 70 the control treatment (un-infested) and infested treatments (44.32, 38.85) respectively (Figure 5). Summer results showed that ZN 3L scored the highest chlorophyll content (48.34) while ZN 9 scored the least (43.37) at day 42 of SPAD reading. Furthermore, on aphid infested plots, ZN 3L recorded the highest (46.82) while ZN 10 scored the least (39.45) (Figure 6). In terms of chlorophyll loss, all the varieties scored < 10% chlorophyll loss (Figure 7). Furthermore, 14 days after the initial SPAD reading, ZN 10 scored the highest chlorophyll loss (7.3 %) which was not significantly different (p> 0.05) from 96-1107 and N14 sugarcane varieties. However, 00-1165 scored the least chlorophyll loss (4.3 %) which was not significantly different (p> 0.05) from ZN 3L. The same trend of results was realised in summer except that there was a great increase in percentage chlorophyll loss margin whereby ZN 10 recorded a 21.4% chlorophyll loss zero days of initial SPAD reading. However, chlorophyll loss of ZN 8 and ZN 9 was not significantly different (p> 0.05); the same trend was also reported between N14 and 96-1107.

2.6.2 Relationship between SPAD values and YSA aphid number on infested sugarcane varieties

Results obtained showed highly significant (p<0.001) positive correlation between SPAD values and Aphid number (Figure 8, Y = -39.5X+2209, P < 0.001, R= 0.69).

2.6.3 Relationship between chlorophyll loss and YSA aphid number on infested sugarcane varieties

Findings of the regression analysis showed a highly significant (p< 0.001) strong positive correlation between chlorophyll loss and aphid number after natural infestation. (Figure 9, Y = 0.028X-6.58, P < 0.001, R= 0.85).

2.7 Effect of YSA on gas exchange responses of selected commercial sugarcane varieties

2.7.1 Photosynthesis

Results exhibited that YSA had a significant effect (p< 0.05) on photosynthetic rate amongst the sugarcane varieties. In winter, 70 days after aphid infestation, ZN 3L scored the highest photosynthesis (31.81) as compared to N14 which scored the least (18.43) in the control treatment (Figure 10). In the aphid infested treatment, 00-1165 sugarcane variety scored the highest photosynthesis as from day zero up to day 48 while N14 scored the lowest (15.92) (Figure 10). Summer results displayed highly significant differences (p< 0.001) on photosynthetic rate. The highest photosynthesis was obtained at day 28 from ZN 3L which recorded (30.75) when compared to the least (18.96) realised on N14 in in the control treatment which was not significantly different (p> 0.05) from 96-1107 (Figure 8). However, in YSA infested plots, 00-1165 recorded the highest compensatory photosynthetic rate (32.52) when compared to the least (12.34) obtained from ZN 10 at day 28 (Figure 11).

2.7.2 Relationship between photosynthesis and chlorophyll loss on infested YSA sugarcane varieties

Results of the regression analysis reviewed highly significant (p= 0.019) positive correlation between chlorophyll loss and aphid number. (Figure 12, Y = -0.40X + 27.03, P = 0.019,R= 0.44).

2.7.3 Relationship between photosynthesis and aphid number on infested YSA sugarcane varieties

Obtained results of the regression analysis showed highly significant (p= 0.002) positive correlation between photosynthesis and aphid number. (Figure 13, Y = -0.01X + 33.25, P = 0.002, R= 0.57).

2.7.4 Relationship between photosynthesis and SPAD values on infested YSA sugarcane varieties

Results of regression analysis showed highly significant (p=0.004) positive correlation between photosynthesis and SPAD values (Figure 14, Y = 0.96X-16.6, P = 0.004, R= 0.52).

2.7.5 Transpiration

Results on transpiration revealed that YSA had a significant effect (p< 0.05) on transpiration rate amongst the sugarcane varieties. In winter, ZN 3L scored the highest transpiration rate (3.5) on day 28 which was significantly different (p< 0.05) from the other control treatments. The lowest transpiration rate (2.40) was recorded on 96-1107 which was not significantly different (p> 0.05) from N14 and ZN 10 sugarcane varieties (Figure 15). On aphid infested plots, 00-1165 sugarcane variety scored the highest transpiration rate (2.96) while N14 recorded the lowest (1.32) (Figure 15). As from day 48 up to day 70, ZN 3L sugarcane variety resumed its high transpiration rate whilst N14 continued to record the least when compared to other sugarcane varieties as YSA are less active. During summer, the highest transpiration rate in the control treatment, was recorded on ZN 3L (4.78) while N14 recorded the least (3.60) which was not significantly different from 96-1107 and ZN 10 (Figure 16). However on YSA infested plots, 00-1165 scored the highest transpiration rate (4.32) when compared to N14 which recorded the lowest (2.12) (Figure 16).

2.7.6 Stomata conductance

Findings of this studyexposed that YSA had a significant effect (p< 0.05) on stomata conductance among the sugarcane varieties. In winter, there were highly significant differences (p< 0.001) among the sugarcane varieties in control plots; 00-1165 scored the highest stomata conductance (308.8) compared to N14 which recorded (165.7) at day 14 (Figure 17). In YSA infested plots, at day 56, 00-1165 variety scored the highest stomata conductance (237.5) in comparison to the lowest (123.6) reported on sugarcane variety 96-1107, which was not significantly different (p> 0.05) from N14 and ZN 8 (Figure 14). Summer results displayed highly significant differences (p< 0.001) on stomata conductance. In aphid control (un-infested) plots, at day zero, 00-1165 variety scored the highest (286.9) while N14 recorded the least (149) (Figure 18). In aphid infested plots, at day 28, 00-1165 sugarcane variety recorded the highest (218.2) stomata conductance when compared to the least (152.9) obtained in N14 (Figure 15). However, 96-1107 and ZN 9 sugarcane varieties were not significantly different (p> 0.05) from each other. Our results highlight that; tolerant sugarcane varieties have the ability to increase or maintain gas exchange in response to YSA infestation.

2.8.1 Aphid number

The study's findings demonstrated that the Lowveld's summer and winter seasons have an impact on the YSA number. This could be the problem of temperature-dependent polymorphism in reproduction (ovivipary and parthenogenesis). Compared to asexual reproduction, which results in greater fertility, sexual method diminishes fecundity. Some authors also reported similar results, indicating that YSA can reproduce in two ways dependent on climate: mating (which produces live individuals and permits females to mate with males in cold winters when low temperatures induce ovivipary (Halbert et al. 2013) and non-mating (parthenogenetically) in warmer climates (Nuessly 2005; Way et al. 2015).

The greatest abiotic factor influencing insect development and reproduction, sex ratio, and longevity, according to a number of authors, is temperature (Harrison et al. 1985; Aalbersberg et al. 1987; Bleicher and Parra 1990; Davis et al. 2006; Keena 2006; Ozder and Saglam 2013; Souza and Davis 2018). According to Souza and Davis (2018), aphids exhibit a significant degree of phenotypic plasticity in response to the environmental circumstances that are present. Poikilothermic species like insects are sensitive to temperature changes since their body temperatures are dependent on the surrounding air temperature (Rosenzweig et al. 2001; Bale et al. 2002; Menendez 2007). Physiological performance of insects is progressively increased by body temperature up to a maximum value, after which it decreases (Briere et al. 1999).

According to Sharpe and DeMichele's (1977) theory, temperature thresholds, both lower and higher, regulate the growth and procreation of certain insect species. The finding that insect development and reproduction rates decline at temperatures above the optimum and finally approach an upper threshold by Briere et al. (1999) provided support for this theory. Aphids are ectothermic species, and temperature has a major impact on their bionomics.

Temperature variations have an impact on metabolic processes such as respiration rates, oxygen consumption, carbon excretion, and neurological and endocrine systems (Neven 1998, 2000). Insect growth lengthens at lower temperatures and shortens with decreasing size at higher temperatures (Angilletta et al. 2004; Pigliucci 2005; Sibly et al. 2007). The physiology and palatability of the host plant are also impacted by temperature (Acreman and Dixon 1989). Various writers have examined how various temperature regimes affect the life cycle, size, fecundity, polymorphism, mating and migration of aphids (Leather and Dixon 1982; Liu 1994; Collins and Leather 2001; Müller et al. 2001; Souza and Davis 2018). Temperature affects many aspects of life, including walking speed, spatial orientation, mortality, fecundity, feeding behavior, and rate of development (Langer et al. 2004; Harrington et al. 2007; Hassal et al. 2007; Colinet and Hance 2009; Zheng et al. 2015; Auad et al. 2015; Schlemmer 2018).

Hinson (2017) investigated how temperature affected the sugarcane aphid (Melanaphis sacchari) and found that 16–29 ⁰C is the ideal temperature range for its life cycle. After examining the effects of varying temperatures on S. flava development, survival, reproduction, life expectancy, and fertility tables while feeding on elephant grass (Pennisetum purpureum), Oliveira et al. (2009) came to the conclusion that the optimal temperature range for S. flava development and reproduction was between 20 ⁰C and 24 ⁰C. Rising temperatures have been proposed by many writers (Dixon 1977, 1987; Kuo et al. 2006; Oliveira et al. 2009; Auad et al. 2009; Auad et al. 2012; Auad et al. 2015) to have a direct impact on aphid survival and abundance.

Research conducted by Flynn et al. (2006) and Auad et al. (2012) highlighted that rising temperatures and CO₂ levels typically have an indirect impact on insectivores through changes in host plant physiology and phytochemistry. Furthermore, Souza et al. (2018) and (2019) noted that aphids' development rate slows down as a result of physiological stress brought on by high temperatures.

2.8.2 Chlorophyll content

The results clearly demonstrate that YSA infestation can have a negative effect on the chlorophyll content of sugarcane leaves (>10% of loss), however resistant plants (00-1165, ZN 9, ZN 8 and ZN 3L) had less chlorophyll loss (< 10% of loss) for both winter and summer seasons. This might be a contribution of genes that codes for more production of nitrogen required for chlorophyll formation. Similar results were reported by Wilson et al. (2011) who reported increased nitrogen in aphid tolerant infested plants due to increased nitrogen reductase. Susceptible genotypes (96-1107, N14 and ZN 10) might not have the same gene expression as in tolerant varieties resulting in increased percentage chlorophyll loss. Reports of increase in nitrogen content in the leaves attacked by the apple green aphid supports the sink theory, as results from other systems have shown (Syvertsen et al. 2003; Urban et al. 2004; Pincebourde and Ngao 2021). In support of this study, similar ranges of results were reported in sorghum against sugarcane aphids by Paudyal (2019) between tolerant and susceptible tested varieties. As revealed by this study, measurement of chlorophyll content has been used as an indication of tolerance for M. sacchari and S. flava (Deol et al.1997; Diaz-Montano et al. 2007b; Akbar 2009; Paudyal 2009) in sorghum. Results of reduced chlorophyll content in susceptible sugarcane genotypes (96-1107, N14 and ZN 10) might have been caused by degradation of chlorophyll or increased production of secondary metabolites. Similar results were reported by Janave (1997) who postulated that oxidative bleaching pathway is one of the two pathways of natural degradation of chlorophyll a which might have been a similar case in our study. In support of this, Ni et al. (2002) showed that feedingby D. noxia caused significant loss of chlorophyll a and b in the damaged regions in wheat. Regression analysis showed a positive strong correlation (r = 0.85) between chlorophyll loss and aphid number. Similar results were reported by Golawska et al. (2010) between Fabacea infested with aphids and SPAD values.

Results corroborates to findings by Haile et al. (1999) and Goławska et al. (2010) who pointed out that reduced levels of chlorophyll may be linked to increased production of many protective secondary metabolites, including saponins. Furthermore, plants infested with aphids may suffer significant harm as exhibited by ZN 10 from highly bioactive effector chemicals found in their salivary gland secretions resulting in yellowing and purplish leaves. This trend is supported by some authors (Cooper et al. 2010; Cooper et al. 2011; Nicholson et al. 2012; Rao et al. 2013; Stiytykiewicz et al. 2013). Furthermore, Gonzales et al. (2002) found ultrastructural damage to chloroplasts in leaves of Johnson grass (Sorghum halepense L.) infested with S. flava, light microscopy indicated increased chloroplast volume, loss of grana structures and aggregation of starch granules. Increased chlorophyllase activity has been seen by certain researchers in plants that have been colonized by aphids (Ni et al. 2002; Ciepiela et al. 2005; Sytykiewicz 2007). According to Sytykiewicz et al. (2013), chloroplast membrane biocatalyst chlorophyllase (also known as chlorophyll chlorophyllidohydrolase; EC 3.1.1.14) is in charge of hydrolyzing free chlorophyll substrates into their corresponding chlorophyllide forms.

Additionally, White (1990) highlighted that S. flava feeding results in leaf discoloration with possible photosynthetic decline, a similar case realized in susceptible sugarcane varieties used in this study. Akbar (2009) revealed that SPAD measurement in different sorghum cultivars ranged from 17-30 % loss of chlorophyll in response to M. sacchari feeding which fits in the category of ZN 10 susceptible sugarcane variety. Also, reports by Akbar (2009) state that chlorophyll loss from S. flava feeding in different sorghum ranged from 27-44 %. However, chlorophyll loss from this study falls short in susceptible sugarcane varieties when compared to the above mentioned range. This might be the effects of YSA host interaction on reproduction of S. flava and intrinsic population dynamics since their studies were in sorghum hence proving the contribution of aphid host interaction. In support of this, a study by Hentz and Nuessly (2004) observed that one to five nymphs per day are developed on sorghum, 8-15 days, they later develop into adults. In other host plants such as sugarcane, development from nymphs to adults takes about 18-22 days; this reveals that reproduction cycle has an impact on YSA prolificacy which influence host chlorophyll loss.

In support of our findings, Lage et al. (2003) pointed out that a variety that maintains relatively high chlorophyll content despite infestation is considered a good indicator of plant tolerance to herbivores as proven by 00-1165. However, ZN 9, ZN 8 and ZN 3L exhibited moderate physiological tolerance although they were not significantly different from each other. Chlorophyll ranges in our study confirms possible yield prediction that feeding by YSA can reduce sugarcane yield by 6% within the first two to three months of growth due to chlorosis losses of up to 19% when more than six leaves are chlorotic (Reagan 1994, Hentz et al. 2004, Nuessly 2005; Nuessly et al. 2010 and Wilson 2019). According to our findings, resistant sugarcane genotypes (00-1165, ZN 9, ZN 8 and ZN 3L) may have high levels of hydrogen peroxide (H₂ O₂₎ buildup and robust antioxidant gene overexpression which may have aided in the development of host plant tolerance. Similar confirmation of such results was reported in resistant sorghum genotypes in response to sugarcane aphid damage (Shankar and Yinghua 2021).

2.8.2 Gas exchange responses

Results of the study showed that gas exchange responses of commercial sugarcane varieties contributes significantly to physiological tolerance against YSA. In aphid infested plots, 00-1165 was more tolerant when compared to other varieties as it recorded high photosynthetic rate, transpiration rate and stomata conductance. Moderate level of physiological tolerance was exhibited in ZN 9, ZN 8 and ZN 3L sugarcane varieties. The reduction of conductance in ZN 10, N14 AND 96-1107 showed there are not tolerant, this further suggests that stomatal interference contributes to decreased photosynthetic rates in susceptible varieties (Meyer and Whitlow 1992).

In this study, infestation of YSA reduced photosynthetic rate. This might be reduction in chlorophyll content which is a pre-requisite for photosynthesis to occur. Phloem cells can be injected with saliva when there is an aphid infestation. Saliva is known to contain a range of effector proteins, hydrolytic enzymes, and toxic compounds that trigger plant perception of aphid invasion and may exacerbate the accumulation of reactive oxygen species (ROS), a precursor to oxidative stress (Shankar and Yinghua 2021). Oxidative stress, defined as high levels of reactive oxygen species (ROS) production within a cell, results in oxidative damage to membranes (lipid peroxidation), pigments, proteins, and nucleic acids, ultimately leading to cell death (Mittler 2002; Gechev 2006). Resistant sugarcane genotype (00-1165) might have been evolved in an antioxidant mechanism to counteract the harmful effects of ROS and shield the plant from oxidative damage by eliminating excess ROS from the cell as highlighted in a study by Shankar and Yinghua (2021). The antioxidant system consists of free transient metals (e.g., Fe2+), antioxidants (e.g., ascorbate, glutathione, and nicotinamide adenine dinucleotide phosphate (NADPH)), and ROS detoxifying proteins (e.g., superoxide dismutase (SOD), catalase (CAT), ascorbate peroxidase (APX), glutathione peroxidases (GPX, glutathione S-transferase (GST), peroxiredoxin (PRX), and ascorbate peroxidases (APX, GPX, and glutathione S-transferase (GST), ascorbate peroxidases (APX, GPX, and GST), ascorbate, and nicotinamide adenine dinucleotide phosphate (NADPH) (Mittler et al. 2004; Apel and Hirt; 2004; Pekker et al. 2002; Foyer and Noctor 2005; Gelhaye et al. 2005; Noctor and Foyer 1998; Pei et al. 2000; Dat et al. 2000; Grant and Loake 2000; Shankar and Huang 2021). Reactive oxygen species (ROS) are recognized for their role as signaling molecules in the control of cellular growth, stomatal closure (Apel and Hirt 2004; Foyer and Noctor 2005; Pei et al. 2000; Dat et al. 2000; Grant and Loake 2000; Shankar and Huang 2021), programmed cell death (Gechev et al. 2006), and response to biotic and abiotic stresses (Gechev et al. 2006; Suzuki et al. 2012). Moreover, increased ROS accumulation can also trigger host plant defense response mediated by systemic acquired immune responses SARS (Wu et al. 1997; Cao et al. 1998; Zhang et al. 1999; Asada et al. 2006). In support of this study, certain theories suggest that plants that experience ROS suppression become more susceptible to aphid attacks, while ROS build up may lead to aphid resistance (Shoala et al. 2018).

Similar results on chlorophyll loss were confirmed by Ni et al. (2002) who showed that feedingby D. noxia caused significant loss of chlorophyll a and b in the damaged regions in susceptible wheat varieties. However, in this study, tolerant (00-1165) and other moderate tolerant (ZN 9, ZN 8 and ZN 3L) varieties were able to maintain high chlorophyll content despite infestation. Results concurs to findings by Lage et al. (2003) who revealed that a variety that maintains relatively high chlorophyll content despite infestation is considered a good indicator of plant tolerance to herbivores.

According to Macedo et al. (2003), transient analysis of chlorophyll a fluorescence revealed two potential mechanisms underlying the plant responses: (i) end-product inhibition of photosynthesis brought on by feeding-induced source-sink manipulation (possibly with concurrent disruption of CO₂ assimilation on the dark reaction); or (ii) photo-inhibition on the light reactions resulting from an increase in activated oxygen species within the chloroplast due to the generation of triplet-chlorophyll in the pigment-protein complex concomitant with disruption of the thylakoid membrane. By measuring the concentrations of the decreasing glucose sugar, biochemical analysis of the non-structural carbohydrate pools in wounded leaves reveals considerable increases in carbohydrate pools following injury. Their results suggested that an accumulation of photosynthetic end-products, which might then set off a sequence of inhibitory reactions, was the main factor affecting plant photosynthesis. This might be the case associated with susceptibility in tested sugarcane varieties (96-1107, N14 and ZN 10).

Results indicating lower stomata conductance and transpiration in sensitive sugarcane cultivars (96-1107, N14 and ZN 10) may be due to YSA's stimulation of the abscisic acid signaling pathway, which in turn caused a reduction in stomata aperture size. Similar findings were published by Sun et al. (2015), who observed that aphid infestation on Medicago truncatula stimulated the abscisic acid (ABA) signaling pathway to reduce the plant's stomatal apertures, hence reducing leaf transpiration. This may have improved the phloem feeding time in aphid-infested plots by increasing xylem feeding time and decreasing hemolymph osmolarity. Moreover, YSA may have caused the decreased stomata conductance by upregulating ABA carbonic anhydrase in stressed YSA infested susceptible sugarcane varieties. The results corroborate those of Guo et al. (2016), who found that probing by aphids increased the expression of carbonic anhydrase, an ABA-independent enzyme.

The stress from YSA increased the amount of ABA produced which might have increased expression of mitogen-activated protein kinase (MPK). The expression of mitogen-activated protein kinases-4 (MPK4), which raises plant susceptibility to YSA and activates anion in the guard cells to start stomata closure, may have been stimulated by the ABA signaling pathway. The findings are corroborated by Jakobson et al. (2016), who observed that MPK4 inhibits HIGH LEAF TEMPERATURE1(HT1, a crucial CO₂-specific regulator of stomatal movement, in order to activate the anion channel SLAC1 in the guard cells and cause stomatal closure). Guo et al. (2017) also came to the conclusion that MPK4 has two effects: it suppresses the jasmonic acid (JA) signaling pathway, which might have caused reduced transpiration and stomata conductance insusceptible sugarcane varieties (96-1107, N14 and ZN 10). According to Guo et al. (2017) decreasing stomatal aperture in an environment with higher CO₂ enhanced aphid feeding efficiency and consequently aphid fitness. This might have been a similar case exhibited by sugarcane varieties under study.

Furthermore, the regression analysis indicated a positive correlation (r = 0.57) between photosynthesis and aphid number. This shows that aphid density influences the photosynthesis outcome as they inflict stress and injury to susceptible sugarcane genotypes. Similar results were reported by Haile et al. (1999) that there is an inverse relationship between the number of Russian wheat aphids (Diuraphis noxia (Mordvilko) and the rate of photosynthesis on wheat. This study reveals that aphids interfere negatively with physiological processes in plants in susceptible sugarcane varieties. Similar discoveries were reported by Nagaraj et al. (2002) who found out that photosynthesis was highly significantly positive correlated with chlorophyll content in green bug damaged sorghum. Furthermore, the same abovementioned author cited that with a small drop in chlorophyll content, a drastic decrease in photosynthesis was noticed (r = 0.36) which falls within the correlation of 0.44 we obtained. In line with this, a significant decline in chlorophyll concentration was reported by Burd and Eliot (1996) in infested leaf tissue of D. noxia-susceptible wheat and barley. The same decrease was also noticed in 96-1107, N14 and ZN 10 sugarcane varieties indicating that they cannot copy with high aphid infestations on tested physiological mechanisms. Genotypes such ZN 9, ZN 8 and ZN 3L were able to exhibit high infestation and still retain chlorophyll content and remain with high photosynthesis. Thus a genotype with a greater correlation or more sensitive relationship between drop in chlorophyll content and photosynthesis is undesirable since productivity will be reduced. In addition to this, Macedo et al. (2003a) reported similar results for wheat under continuous light, but under continuous darkness for 72 hours there was no change in photosynthesis induced by Russian wheat aphids. Furthermore, Frazen et al. (2007) showed that tolerant wheat had similar rates of photosynthesis compared to the control and an antibiotic-resistant cultivar delayed photosynthetic senescence. This similar trend was shown on the tolerant variety (00-1165) and moderate tolerant varieties (ZN 9, ZN 8 and ZN 3L) that these varieties can be used in environments where YSA is a menace because they expressed physiological tolerance. Contrary to our finding, Gomez et al. (2006) found no change in the rate of photosynthesis in cotton after a 9-day infestation with cotton aphids (Aphis gossypii G.).

Reduction in gas exchange responses has been observed in infested hosts in sucking insect pests such as Nilaparvata lugens in rice (Watanabe and Kitagawa, 2000) and Bemisia argentifolli in cotton (Lin et al., 1999). However, in contrary, Macedo et al.(2009) reported that Diuraphis noxia does not affect transpiration rate in wheat plants. Furthermore, Hawkins et al. (2006) found that aphid-infested plants had more net CO₂ exchange rates than their respective controls. Moreover, Heng- Moss et al. (2003) speculated feeding mainly on phloem tissue by the aphids result in pH change either on the luminal side of the thylakoid membrane, preventing the formation of zeaxanthin and the regeneration of violaxanthin on the stromal side where the process occurs. Additionally, Burd and Elliott (1996) showed that aphid feeding has the possibility of reducing protein synthesis causing photoinhibition to be irreversible and blocking electron transport on the photosystem II reaction center. This might result in over-reduction in the system reducing chlorophyll formation and photosynthesis in infested tested species. Also, Haile et al. (1999) and Heng-Moss et al. (2003) found a significant decline in the photosynthetic rate in aphid-infested leaves. Their study speculated that this might have resulted from increased biochemical resistance in response to insect herbivory hence a possible contribution to this study. The decline in chlorophyll concentration as exhibited by yellowish and purplish discoloration in our study might also be due to the increased production of defensive compounds. This calls for further studies to accurately conclude. Among the studied sugarcane species, the number of aphids was lowest on 00-1165 plants, indicating that they are less attractive to YSA.

Results of this study confirms those found by various authors (Haile et al. 1999, Haile and Higley 2003; Macedo et al., 2003a,b; Frazen et al. 2007; Gutsche et al. 2009a; Pierson et al. 2010a; Pierson et al. 2011; Prochaska 2015) who reported reductions and compensatory mechanisms specifically in crops with demonstrated tolerance to a specific aphid species. These studies indicate that compensatory mechanisms found in tolerant sugarcane varieties may be connected to the ability to maintain or increase Ribulose Bi-phosphate (RuBP) and rubisco activity. In support of this hypothesis, Haile et al. (1999) predicted that physiological responses such as gas exchange can contribute to Russian wheat injury resulting in wheat seedlings recording lower light saturation points in infested seedlings.

Most often, the loss of photosynthetic tissue after feeding by defoliating insects induces an increase in the photosynthetic rate per unit area in the remaining leaf tissues, allowing the plant to partially compensate for the herbivory (Welter 1989). In other cases, herbivory induces a decrease in assimilation rate in the remaining leaf tissues (Zangerl et al. 2002). Large reductions in photosynthesis have also been measured on leaves infested by mesophyll eaters such as spider mites (Welter 1989; Haile and Higley 2003) and stink bugs (Velikova et al. 2010). For example, aphid and spider mite effectors have been shown to suppress plant defense signals and responses, thereby enhancing herbivore performance (Atamian et al. 2013; Naessens et al. 2015; Schimmel et al. 2017).

The effects of insect feeding on leaf stomata conductivity and transpiration rate also vary widely among the sugarcane genotypes as indicated by our study. Similar results were confirmed by Shannag (2007) metabolically active substances secreted with aphid saliva into the phloem that increased transpiration and stomatal conductance rates. These in turn interfere with the regulation process of water vapour from the host plant. Yellowing and reddening of leaves inflicted by YSA in our study reduced leaf area for physiological processes. In support of this, Aldea et al. (2005) confirm that insect damage increased soybean water loss around damaged tissue. Additionally, the same authors further reported that both net photosynthesis and stomatal conductance of the remaining leaf tissue were unaffected by the leaf deciduous beetle (Popillia japonica) and caterpillar (Helicoverpa zea) (Aldea et al. 2005). In contrast, Tang et al. (2006) found that both water stress caused by increased water loss near the injured tissue in Arabidopsis thaliana and reduced stomatal conductivity in tissues distant from the injury by the Lepidoptera Trichoplusia ni inhibited photosynthesis. Pincebourde and Ngao (2021) concluded that assimilation and transpiration rates are affected simultaneously; water loss increases while photosynthesis decreases. In the first case, leaf efficiency (water use efficiency) is at best kept constant as the plant compensates for tissue loss due to herbivory. However, although full compensation is rare as reported by Peterson (2000), mitigation is possible and may contribute to plant tolerance to herbivores (Pincebourde et al. 2006). The observed variability in leaf gas exchange responses to insect damage remains difficult to explain as mentioned by Pincebourde and Ngao (2021).

Variations in the photosynthetic rate show different physiological mechanisms for resistance (Pierson et al. 2011; Paudyal 2019). Furthermore, YSA injury significantly reduced the photosynthetic rate of 96-1107, N14 and ZN 10. Significantly higher photosynthetic rates were observed in resistant entries, 00-1165, ZN 9, ZN 8 and ZN 3L on YSA infested plots. This is supported by the hypothesis by Pierson et al. (2011) in sorghum that plants increase their photosynthetic rate to compensate for the feeding injury, a similar case with YSA in our study.

Findings of this research demonstrates that YSA sugarcane resistant plants appear to be able to compensate for injury caused by YSA feeding through increased chlorophyll content, maintaining or increasing; photosynthetic integrity, transpiration and stomata conductance when compared to susceptible genotypes. These findings support previous research with other aphid species (Retuerto et al. 2004; Heng-Moss et al. 2006; Frazen et al. 2007; Gutsche et al. 2009; Akbar et al. 2009; Paudyal 2019; Paudyal et al. 2020). However other variables like; leaf age, nutritional composition and micro-environment parameters may also influence photosynthesis as reported by Nagaraj et al. (2002). Knowledge of the physiological alterations occurring in sugarcane leaves infested by YSA may provide information on resistance and defense responses. This will be leveraged to develop new YSA resistant sugarcane cultivars.

Sugarcane varieties under investigation displayed significant differences on chlorophyll content and gas exchange responses (photosynthesis, transpiration and stomata conductance). Furthermore, 00-1165 exhibited high physiological tolerance while moderate tolerance was exhibited on ZN 9, ZN 8 and ZN 3L as they were not significantly different from each other. These varieties were able to compensate/maintain chlorophyll content and gas exchange in response to YSA feeding. However, 96-1107, N14 and ZN 10 were the most susceptible to physiological damage posed by YSA.

2.10 Recommendations

Sugarcane growers should incorporate the pre-released sugarcane variety 00-1165 amongst the existing sugarcane varieties as it less susceptible to YSA damage as proven by physiological compensatory/maintaining behaviour. There is need to integrate physiological tolerance into existing IPM strategies in the management of YSA. Molecular breeding is required in order to incorporate the gene expression of physiological tolerance among susceptible sugarcane varieties thereby enhancing sugarcane productivity in the presence of YSA.

2.11 Future studies

To gain a better understanding of the molecular mechanisms of ROS-mediated aphid resistance in sugarcane, additional experiments involving additional enzymes and remaining members of the enzyme families are required. Our study has been restricted to supporting discussions on specific genes associated with the ROS scavenging pathways that may be involved in sugarcane resistance to YSA. To close the information gap regarding the genetic and metabolic basis of aphid resistance in sugarcane, detailed elucidation of H₂ O₂ interactions with other plant defense pathways, such as plant hormones and transcription factors, is also necessary.

Author contributions

Sakadzo, Nyasha: conceptualization; experimental design; data collection, analysis and interpretation; writing – original draft; writing and editing. Dr Mubvuma, Michael: supervision– review of content and editing. Ms Mukanga, Concilia: supervision; funding acquisition; writing - review and editing: Dr Mabveni, Audrey.R.S: conceptualization; funding acquisition; proof reading. Prof Musundire, Robert: supervision - review and editing; revising critically for intellectual content: final approval of the version to be published.

Acknowledgement

We would like to appreciate the Zimbabwe Sugar Association Experiment Station and sugar industry for providing resources during the experiment.

Disclosure statement

Authors declare no potential conflict of interest.

Supplementary material

This will be made available by the authors without any reservation.

Funding

No funding was received for this work.

Data availability statement

The data that support the findings of this study are in the manuscript and readily available on request from the corresponding author.

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Physiological response as a tolerant mechanism to Yellow Sugarcane Aphids (YSA) (S. flava) herbivory on selected commercial sugarcane varieties (Saccharum officinarum)

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