7.1 Weeds

Weeds are a problem in a sugarcane crop due to yield reduction caused by competition or allelopathy and interference with harvesting machinery which reduces product quality (McMahon et al. 2000). In Australia, in 2000, weeds were estimated to cost the sugarcane industry $70 million each year, in both control costs and lost production (McMahon et al. 2000). As well as controlling weeds within the crop, it is important to control weeds around the farm to reduce any high protein food, such as weed or grass seeds, which rats need to breed (McMahon et al. 2000). See section 7.2.1 for a discussion of rats as a pest of sugarcane.

There are a number of weeds that infest sugarcane plantations including grasses, broadleaf weeds, vines and sedges. Those weeds that are a major problem in Australia are discussed below.

The predominant grasses that occur in sugarcane growing districts include barnyard grass (Echinochloa crus-galli), awnless barnyard grass (Echinochloa colona), couch grass (Cynodon dactylon), wild sorghum (Sorghum spp.) and guinea grass (Panicum maximum). Pasture grasses in particular can be problematic when the land is subsequently used to grow sugarcane (McMahon et al. 2000). Hymenachne (Hymenachne amplexicaulis) is an aquatic grass to 2.5 m tall and has been declared a Weed of National Significance (Queensland Department of Primary Industries and Fisheries 2007b). It was planted extensively in northern QLD and the Northern Territory (NT) as a fodder crop and has since escaped from cultivation. It can block irrigation and drainage channels in sugarcane plantations and contaminate sugarcane crops. Spraying with herbicides every three months is used to control hymenachne (CRC for Weed Management 2003). Imperata cylindrica is a perennial species that commonly grows on degraded or burnt-off land in most Australian sugarcane-growing districts (Lazarides et al. 1997). It is a common weed in QLD, and although it occurs in all Australian states, it is not listed as a noxious weed in any jurisdiction (National Weeds Strategy Executive Committee 2009).

The most important and prevalent weed of sugarcane is sedge nut grass (Cyperus rotundus), although in wetter areas other sedges also occur (McMahon et al. 2000). C. rotundus was estimated in 1967 to reduce cane yields by 10 t ha-1 (Chapman as cited in McMahon et al. 1989). It spreads mainly by tubers, which are produced in very large numbers and are carried in soil and by flood waters. It also reproduces by seed, although apparently only rarely. It withstands cultivation extremely well, and this process rapidly spreads the tubers around and between fields (DPIW- Tas 2009).

Broadleaf weeds such as blue top/billygoat weed (Ageratum spp.) and purslane/pigweed (Portulaca oleracea) tend to be less of a problem and can be controlled relatively easily if targeted when the plants are young. Broadleaf weeds tend to be more regional and soil specific (McMahon et al. 2000). The giant sensitive plant, also known as tropical blackberry (Mimosa diplotricha or M. invisa), is a serious weed in northern QLD and has been identified as a weed that can invade sugarcane crops (Queensland Department of Primary Industries and Fisheries 2007c). It can be controlled using an introduced sap sucking insect (Heteropsylla spinulosa) as a biological control agent, or with slashing or herbicide use (McMahon et al. 2000; Queensland Department of Primary Industries and Fisheries 2007c).

Vines have become an increasing problem after the adoption of trash-blanketing, although a thick layer of trash has been shown to inhibit their growth (Fillols & Callow 2010). They have the potential to grow rapidly and if left uncontrolled can impede the harvesters (McMahon et al. 2000). The most problematic vines in sugarcane include bindweed (Convolvulus spp.), passionvine (Passiflora spp.) and morning glory (Ipomoea spp.) (McMahon et al. 2000).

There are a number of herbicides that can be used to control weeds in sugarcane. These include pre-emergent herbicides such as isoxaflutole, imazapic or a diuron/hexazinone mix (Fillols & Callow 2010). Herbicides such as 2,4-D amine can be used on broadleaf weeds. Paraquat, a non-selective herbicide, can be used on broadleaf, grassy and other weeds (McMahon et al. 2000). However, a study in north QLD which evaluated a range of herbicides for their effectiveness against major sugarcane weeds showed unacceptable damage to a number of sugarcane cultivars from post-emergence paraquat application (Makepeace & Williams 1986).

7.2 Pests and diseases

The cost of controlling the major pests and diseases of sugarcane to the sugarcane industry in Australia was estimated to be $111 million in 1996 (McLeod et al. 1999). This included $14 million in lost production and control costs for pests, and $97.4 million in loss and control for diseases (McLeod et al. 1999). The major pests and diseases that cause losses in sugarcane production include cane grubs, feral pigs, ratoon stunting disease (RSD), sugarcane rusts, chlorotic streak and soil-borne diseases (McLeod et al. 1999). More recently, sugarcane smut has become a serious threat to the industry. To reduce the impact and prevent new outbreaks of pests and diseases Australia maintains strict control and quarantine guidelines and develops a response strategy in the event of an incursion. To this end the sugarcane industry has worked with government agencies to develop the National Sugar Industry Biosecurity Plan (Plant Health Australia 2009). The major pests and pathogens relevant to the Australian sugarcane industry are discussed below.

7.2.1 Pests

Invertebrate pests
There are many insect pests of sugarcane and some insects such as plant hoppers (Perkinsiella saccharicida), are also known vectors of diseases (Croft et al. 2000; Allsopp et al. 2002). Appendix 1 gives an overview of these insect pests and the major insect pests are discussed below.

Cane grubs (melolonthine white grubs, larvae of the endemic melolonthine beetle) are major pests affecting the sugarcane industry. In 1996 they were estimated to cost the Australian sugarcane industry over $11 million a year in production loss and control costs (McLeod et al. 1999). They destroy the roots of the sugarcane plants, preventing water and nutrient uptake and causing lodging (Allsopp et al. 2000). There are 19 native species of cane grub which cause significant damage in cane fields in different regions, with the greyback canegrub (Dermolepida albohirtum) showing the most widespread damage (Robertson et al. 1995). This was estimated to cause a crop loss of 1 million t of cane in the 2000–2001 season (Chandler & Tucker 2010). The different species occur on different soil types and in different geographic regions and have either a one or two year lifecycle, yet in both cases the damage is caused by the third (final) larval stage (Allsopp et al. 2000). Several methods can be used for the control of these insects (Robertson et al. 1995). The application of the insecticide chlorpyrifos or the biological control agent Metarhizium anisopliae (a fungus that attacks the larvae) soon after planting, control the species for two to three years. Other insecticides such as granular cadusafos and liquid imidacloprid, applied after harvest or applied to the ratoon stubble, are also of use but require irrigation or rain to make them effective. Insecticide applications are complicated by factors such as stability in different soil types, long term nature and inaccessibility of the crop, difficulties associated with soil dwelling insects, and differences in life-cycle duration of the species (Robertson et al. 1998; Allsopp et al. 2002). Farming practices have also been shown to have an effect on the incidence of Childers canegrubs (Antitrogus parvulus Britton), with ratoon crops, fields that had been ploughed out and immediately replanted, and dryland crops having the largest infestations (Allsopp et al. 2003). A study on the greyback cane grub indicates that it preferentially oviposits on tall sugarcane plants, and that the taller blocks of cane show higher damage levels (Ward 2003).

Other insect pests of sugarcane include sugarcane and yellow soldier flies (Inopus rubriceps and Inopus flavus respectively), wireworms (Agrypnus variabilis, Heteroderes spp. and Conoderus spp.), armyworms including day and night feeding species, as well as loopers (Allsopp et al. 2000). Nematodes, including root-knot nematodes and lesion nematodes can be a serious pest. Estimates of yield losses have suggested that they may cause a 10% loss in plant crops and a 7% loss in ratoon crops (Blair & Stirling 2007). This has been estimated to cost the Australian sugarcane industry $82 million year-1 (Ogden-Brown et al. 2010).

Vertebrate Pests
There are numerous vertebrate pests of sugarcane including ground rats (Rattus sordidus), climbing rats (Melomys burtoni), wallabies, striped possums (Dactylopsila trivirgata), the eastern swamphen (Porphyrio porphyrio), cockatoos (Cacatua galerita), foxes and feral pigs. All these, except for the fox and feral pig, are native to Australia and are consequently protected. Permits for control of native animals in cane fields must be obtained from the relevant Cane Protection and Productivity Board. Rodents are the most serious pest to the sugarcane industry after the cane grub. During the 1999 and 2000 seasons they destroyed 825,000 t of sugarcane valued at $25 million (Dyer 2005). Ground rats cause more economic damage than climbing rats as they are more numerous, especially south of the Herbert River (Dyer 2005). They cause yield loss directly by gnawing the cane, but the damage also allows the cane to dry out and provides entry-points for bacterial and fungal attack (Dyer 2005). In addition, rats are known to be carriers of the bacterium Leptospira which can result in Leptospirosis disease in humans. The disease can be spread through soil, mud and water that have been contaminated with urine from infected animals (NSW Health 2007). Integrated pest management is now widely employed to discourage and control rats (Smith et al. 2002). Strategies such as controlling crop weeds have been shown to reduce juvenile rat numbers by 50% and reduce crop damage by 60% (Dyer 2005).

7.2.2 Pathogens

Various biological agents including bacteria, fungi and viruses cause diseases of sugarcane. Important diseases of sugarcane that have been identified in Australia are listed in Appendix 2 and the major pathogens are discussed below.

Disease control in sugarcane is based on an integration of legislative control, resistant cultivars and other management procedures. Short term spraying options are available, but their economic viability may not be sustained. Hygiene is important to disease management strategies, particularly for diseases transmitted through cuttings such as ratoon stunting disease (RSD) and leaf scald. Cutting one infected stalk may lead to significant infection to the next 100 cuttings which are subsequently cut by the same blade (Croft et al. 2000). Machine harvesters can also transmit disease.

Many sugarcane diseases are also managed through the use of disease-free planting material supplied through Cane Protection and Productivity Boards. Hot-water treatments are used to disinfect planting material. Long hot-water treatment (three hours at 50C) is used to control RSD. Soaking in ambient temperature running water for ~40 hours followed by three hours at 50C is used to control leaf scald bacteria. Short hot-water treatment (50C for 30 minutes) is used to control chlorotic streak and some insect pests (Croft et al. 2000).

Bacterial diseases
RSD is probably the most important disease of sugarcane. It is a highly infectious disease caused by Leifsonia xyli, (formerly named Clavibacter xyli subsp xyli) which infects vascular tissues of sugarcane. It was first reported in QLD in 1944–45 and has been identified in most countries that grow sugarcane (Bailey 2004). It was estimated in the early 1990’s that it affected approximately 30% of farms in NSW (Roach et al. 1992; McLeod et al. 1999) and the estimated loss from this disease Australia-wide was $6.3 million in 1996 (McLeod et al. 1999). The symptoms are poor growth and stunted shoots, which might not be obvious if most plants in the field are infected. The visual symptoms of red-orange dots in the vascular tissues can be seen only when the stalks are cut and sliced (Croft et al. 2000). The disease is transmitted through healthy plants coming in contact with diseased plant material or contaminated cutting implements. Yield loss is higher in dry weather and often becomes more severe in subsequent ratoon crops (Frison & Putter 1993). The incidence of the disease in parts of NSW has been reduced due to an extension campaign promoting clean seed cane (McGuire et al. 2009)

Leaf scald is caused by the bacterium Xanthomonas albilineans which infects the vascular tissues of sugarcane. It is found in most sugarcane districts in QLD (Croft et al. 2000), although it is hard to identify and the disease often has a latent period after infection (Bailey 2004). Leaf scald is characterised by a long white to cream streak on the leaves. Severely infected leaves appear scalded and roll inwards, with the top of the shoots becoming chlorotic. Yield loss occurs through the death of infected cane stalks and poor ratooning (BSES Ltd 2005b). Leaf scald can spread by wind-blown rain, plant material and contaminated cutting equipment such as planters and harvesters (Croft et al. 2000). Leaf scald can infect many other grasses which are alternate hosts and act as a reservoir for the disease. Extremes of moisture and temperature favour disease transmission. Resistant cultivars are used to curb the spread of the disease and susceptible plants are not used in breeding programs (BSES Ltd 2005b).

Fungal and Oomycete diseases
The two major rusts in sugarcane are orange and common sugarcane rusts (Braithwaite et al. 2009). Orange rust is caused by Puccinia kuehnii and is not as economically important as the common rust, caused by P. melanocephala. These are both obligate parasitic fungi spread by windblown spores. The disease symptoms of the two rusts are distinct. Pustules of the orange rust are orange and tend to be grouped in clusters, while those of sugarcane rust are reddish brown and are distributed evenly on leaves. Pustules rupture the leaves and allow water to escape from the plant, leading to moisture stress. (Croft et al. 2000). Both diseases are most severe in humid environments with temperatures below 25C (Walker 1987).

In 1999–2000, sugarcane crops in Australia were affected by an outbreak of orange rust, which severely damaged the most widely grown commercial cultivar, Q124 (Croft et al. 2000). Highly susceptible parents are no longer used in any breeding programs. More recently, cultivars such as Q173 and Q182 have also been found to be affected by the disease (NSW Sugar 2005).

Yield loss from sugarcane rust depends on environmental conditions and was estimated to cause an economic loss of $3.5 million in 1996 (McLeod et al. 1999).

Sugarcane smut, caused by Ustilago scitaminea, is a serious disease of sugarcane that can reduce yields by 30–100% (Watson 2007). Infection occurs through the sugarcane buds from wind-blown spores (Walker 1987). The disease causes severe stunting and multiple thin stalks. It is characterised by black, whip-like structures that form at the growing points of sugarcane plants (Croft et al. 2000) (Figure 7). These whips replace the spindle leaves and are formed in the shoots developing from infected cane cuttings (Frison & Putter 1993). The whips break open to release the mature spores which are spread by wind (BSES Ltd 2006). There was an outbreak of smut in Australia in July 1998 in the Ord River area of WA. This outbreak was controlled and the disease was not detected in eastern Australia until 2006, when it first appeared in Childers, QLD. Sugarcane smut is now seen as being widely spread and established (Croft et al. 2008). Estimated losses in susceptible cultivars are up to 62% in the Herbert region (Magarey et al. 2010). The spread and occurrence of the disease is being controlled through planting of resistant cultivars, using uninfected seed canes and removing infected crops (BSES Ltd 2009b). However, there has not been a complete conversion to resistant cultivars as the resistant cultivars can have a lower yield than the non-resistant cultivars. Yield of resistant cultivars is also dependant on soil type in the region (Watson 2007).

In order to reduce the spread of sugarcane smut, the movement of sugarcane and sugarcane machinery is restricted in QLD by the Plant Protection Regulation 2002. Provisions under the Plant Protection Act 1989 (QLD) allow for inspectors to order the destruction of diseased cane and practical guidelines have been developed to control the spread of the disease (BSES Ltd 2005a; BSES Ltd 2007a; BSES Ltd 2007b; Queensland Government 2009).

Figure 7. Smut on Saccharum spp. hybrid in Bundaberg (May 2010). Photo taken by H. Mitchell, OGTR

Pachymetra root rot caused by Pachymetra chaunorhiza, an Oomycete, is a disease only found in Australia in the QLD sugarcane districts (Magarey & Bull 2003). It was first identified as northern poor root syndrome in the 1970s, before the disease-causing organism was identified (Magarey 1994). The disease seems to favour high rainfall areas, and spores can survive up to five years in the soil. In northern QLD, surveys indicate that almost every field is infected with the pathogen. The disease is characterised by a soft rot of the primary and some secondary roots, leading to poor root development. Yield loss caused by Pachymetra root rot was estimated to be up to 40% in highly susceptible cultivars (Croft et al. 2000). Fungicides have not been effective at an economical rate and control is based on planting resistant cultivars (Magarey 1996).

Other fungal diseases of sugarcane are minor (see Appendix 2) and cause less impact on yield.

Viral diseases
Sugarcane can be affected by a number of viral diseases (see Appendix 2).

Chlorotic streak is thought to be caused by a virus. The disease occurs in all eastern sugarcane districts, especially in wet and poorly drained fields. Lower incidence of the disease is generally found in drier regions (Croft et al. 2000). The symptoms are yellow to white streaks on the leaf, midrib and leaf sheath. Older streaks change to yellow and are more visible than younger streaks. This is followed by the appearance of chlorosis in the middle of the leaves. Internal vascular bundle tissues may be reddish in colour. (Croft et al. 2000). The disease is transmitted by soil water and diseased seed cane. Yield losses may be up to 40%, with waterlogging compounding the losses. Ratooning may also be poor (BSES Ltd 2009a).

Fiji leaf gall (previously called Fiji leaf disease) is caused by Fiji disease virus (FDV) and can lead to stunting and death of infected plants (Ridley et al. 2006). The initial symptoms are whitish galls raised on the underside of the leaf blade and midrib. Galls are produced due to the disorder of cell proliferation in the phloem and xylem. Galls can vary from white to green and the surface is usually smooth. When the gall is old, the epidermis may be ruptured and appear brown. At an advanced stage of infection, stem development slows down. Successive leaves become smaller and stiffer with the whole top part of the stem developing a fan-like appearance (Croft et al. 2000). Fiji disease can be transmitted by infected cuttings and plant hoppers (Perkinsiella saccharicidae) are a known vector for the disease. Significant yield loss was recorded in 1970s in QLD (Croft et al. 2000) but due to the intensive management program put in place, there have been no reports of disease incidence since the 1980s. However, FDV is present in southern cane growing areas and plant hoppers are present in all canegrowing areas of QLD and NSW (Ridley et al. 2006).

Worldwide, sugarcane mosaic is caused by a number of potyviruses such as the Sugarcane mosaic virus (SCMV). Currently in Australia, only the Sugarcane mosaic virus strain A is present, which is a mild form of the virus (BSES Ltd 2008b). The mosaic symptom pattern appears in young growing leaves. Once the leaves are older, infected leaves may appear relatively normal as the mosaic becomes green. Yield loss caused by sugarcane mosaic was 40% in some fields in Australia (Croft et al. 2000). Aphids transmit the disease, as can seed produced by infected cane.

7.2.3 Other biotic interactions

Sugarcane may have symbiotic relationships with a number of bacteria that fix nitrogen.

In Brazil, sugarcane is grown with low nitrogen inputs (50 kg ha-1) compared to other countries who use >200 kg ha-1 (Boddey et al. 1991). The cane is commonly burnt before harvesting in Brazil so little nitrogen is returned to the field. This low nitrogen requirement has led to the suggestion that some cultivars of sugarcane can obtain nitrogen via biological nitrogen fixation (BNF). The occurence of BNF has been suggested in several pot studies where some cultivars of sugarcane have thrived for several generations without the addition of nitrogen (Boddey et al. 1991; Urquiaga et al. 1992). Differences were seen between plant genotypes, but it was estimated that BNF could account for 25–60% of the nitrogen assimilated in one study (Boddey et al. 2001) and up to 70% in another study (Urquiaga et al. 1992). The organisms responsible for this have not been unequivocally determined. Studies have focussed on endophytic bacteria such as Gluconacetobacter diazotrophicus (previously called Acetobacter diazotrophicus), however these bacteria were not shown to be producing nitrogenase in planta (James et al. 2001). Despite this, a study using G. diazotrophicus - innoculated plants found large increases in nitrogen fixation under nitrogen deficient conditions. This nitrogen fixation did not occur after inoculation with a mutated nitrogenase deficient form of the bacterium (Sevilla et al. 2001).

G. diazotrophicus may also play a role in defence against sugarcane pathogens. It inhibited in vitro growth of Colletotrichum falcatum (red-rot) (Muthukumarasamy et al. 2000) and Xanthomonas albilinean (leaf scald) (Pin et al. 2002; Blanco et al. 2005). Additionally G. diazotrophicus-inoculated sugarcane stems were resistant to infection by X. albilineans (Arencibia et al. 2006). There is also some evidence that it may promote sugarcane growth by production of a growth promoting factor (Sevilla et al. 2001), such as auxin (IAA; indole-3-acetic acid) or by solubilisation of mineral nutrients (as reviewed in Saravanan et al. 2008).

Other bacterial species have been isolated from sugarcane that may play a role in nitrogen fixation including Agrobacterium diazotrophicus (Xing et al. 2006), Herbaspirillum spp (Reis et al. 2007) and Azospirillum spp. (Baldani et al. 1997). Experiments have shown that co-inoculation of G. diazotrophicus and Herbaspirillum spp gave enhanced sugarcane biomass compared to inoculation with either the single species, or to uninoculated controls (Muthukumarasamy et al. 2006). A field-based experiment and surveys of sugarcane fields in NSW and QLD showed no evidence of biological nitrogen fixation as a source of nitrogen (Biggs et al. 2000).

Vesicular-arbuscular mycorrhizal fungi (VAM) have been found in north QLD sugarcane fields in association with sugarcane roots. These fungi are known to colonise plant roots and may supply the plant with mineral nutrients, especially phosphorous. Pot experiments, using soil and mycorrhial spores from cane fields, showed that the addition of VAM increased the yield of soybean and maize plants. However, no effects have been seen on sugarcane growth from addition of the VAM Glomus clarum at various phosphorus levels in pot experiments (Kelly et al. 2001; Kelly et al. 2005). Similar experiments in wheat have shown that although there is no increased yield following root colonisation with VAM, 50% of the phosphorus in the plants had been absorbed via VAM (Li et al. 2006).

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