Growth & Development

6 Growth and Development Caroline S. Awmack1 and Simon R. Leather2 1Department of Entomology, University of Wisconsin-Madison, Madison, WI 53706, USA; 2Division of Biology, Imperial College London, Silwood Park Campus, Ascot, Berks, SL5 7PY, UK Introduction The growth and developmental rates of individual aphids have been studied extensively since the early investigations of Davis (1915) because they can be reliable indicators of future population growth rates (Leather and Dixon, 1984; Acreman and Dixon, 1989). In this chapter, we discuss the methods used to measure aphid growth and development, the relationships between these measures of aphid performance, and the reliability of using the results of such experiments to fpredict the performance of field populations of pest aphids. Individual aphids frequently have extremely high growth and developmental rates, allowing aphid populations to rapidly reach levels that are damaging to crop plants. Under optimal growth conditions, an individual aphid typically commences reproduction 7–10 days after it is born (Dixon, 1998). Such short development times are possible because newborn aphids contain the embryos of their first grand-daughters. This ‘telescoping of generations’ means that an individual aphid has already completed two-thirds of its development before it is born (Dixon, 1998). Growth and developmental rates can be used to predict future fecundity because somatic growth and reproductive development occur simultaneously in the developing nymph, ©CAB International 2007. Aphids as Crop Pests (eds H. van Emden and R. Harrington) and are thus simultaneously affected by any change in the rearing environment. Aphid growth and developmental rates are frequently measured in small-scale trials using single individuals, or small groups of similarly aged nymphs. In the first section of this chapter, the most commonly used measurements of aphid performance are described, with the relevant experimental techniques needed to investigate them. Factors that may affect the reliability of this approach are then discussed, with a particular emphasis on studies that have investigated the relationships between the various measures of individual performance, and between individual and population growth rates. Definitions Aphid growth and developmental rates have been used extensively to predict the performance of aphid populations on crop plants because they correlate well with potential fecundity, achieved fecundity, and the intrinsic rate of increase, rm (Leather and Dixon, 1984; Dixon, 1990). Many aphid species show strong positive relationships between growth rates and potential fecundity (Lyth, 1985; Fereres et al., 1989). Similarly, adult size is frequently correlated with both 135 136 potential fecundity (Dixon and Dharma, 1980; Kempton et al., 1980; Bintcliffe and Wratten, 1982; Llewellyn and Brown, 1985) and achieved fecundity (Leather and Dixon, 1984; Dixon, 1990; 1998). ‘Growth’ is defined here as an increase in aphid size; ‘development’ is used to imply increasing reproductive maturity; ‘potential fecundity’ is a measure of the reproductive potential of an individual aphid – for example, the number of mature embryos contained in an adult; while ‘achieved fecundity’ is the total number of progeny produced by an adult. Potential fecundity may be a useful measure of aphid performance, as potential and achieved fecundity frequently are strongly positively correlated (Dixon and Wratten, 1971; Dixon and Dharma, 1980; Frazer and Gill, 1981; Leather and Dixon, 1984). Developmental and reproductive rates can be combined in the intrinsic rate of increase, rm (Birch, 1948; Wyatt and White, 1977), an estimate of future population growth rates based on the performance of individual aphids. Uses of aphid growth and developmental rates The use of aphid relative growth rates to measure performance was initially suggested by van Emden (1969) as a means of measuring the direct effects of plant nutrition on aphid performance, without the confounding effects of maternal experience. Dixon (1990) and Leather and Dixon (1984) later demonstrated that relative growth rates and the intrinsic rate of increase, rm, (Wyatt and White, 1977) were strongly positively correlated (Fig. 6.1). As a consequence, many authors have used mean relative growth rate (MRGR) or relative growth rate (RGR), defined below, when quick estimates of aphid performance are required; for example, to assess the resistance of plants to aphid attack (van Emden, 1969; Bintcliffe and Wratten, 1982; Givovich et al., 1988; Leszczynski et al., 1989; Wojciechowicz-Zytko and van Emden, 1995; Farid et al., 1998; Telang et al., 1999), the quality of different growth stages of the host plant (Leather and Dixon, 1981), the effects Intrinsic rate of increase (rm) C.S. Awmack and S.R. Leather Mean relative growth rate or relative growth rate Fig. 6.1. Hypothetical relationship between aphid growth rates and the intrinsic rate of increase (rm). of temperature (Berg, 1984; Lamb et al., 1987), the effects of air pollution (Dohmen, 1985; Holopainen et al., 1997; Wu et al., 1997), and the contribution of bacterial symbionts to aphid nutritional ecology (Adams and Douglas, 1997). Similarly, the intrinsic rate of increase has been used to investigate the responses of individual aphids to changes in plant quality (Bintcliffe and Wratten, 1982; Lykouressis, 1984; Zuniga et al., 1985; Leszczynski et al., 1989), temperature (Landin and Wennergren, 1987; Liu and Yue, 2000), drought stress (Sumner et al., 1986), atmospheric pollutants (Warrington et al., 1987; Awmack et al., 1997), host plant virus infection (Fereres et al., 1989), and the sub-lethal effects of insecticide residues (Kerns and Gaylor, 1992). Measurement of Aphid Growth and Developmental Rates Growth rates Most investigations of aphid growth rates have used living aphids to measure gain in fresh weight over a defined time period, weighing the same individuals at the beginning and the end of the experiment (e.g. Banks and Macaulay, 1964; Tsitsipis and Mittler, 1976; Frazer and Gill, 1981; Kindlmann et al., 1992; Sunnucks et al., 1998; Manninen et al., 2000; Edwards, 2001), although some authors, such as Lamb (1992) have used dry 137 Growth and Development weight and have therefore used different individuals for each measurement. Other investigations of aphid growth rates have taken advantage of the allometric relationships between aphid skeletal structure and size (Reddy and Alfred, 1981), using hind tibia length (Campbell, 1983) or antennal length (Varty, 1964; Murdie, 1969a,b). However, since the ratios of the sizes of some aphid body parts may change as the developing aphid grows (Varty, 1964; Dixon, 1987), such allometric relationships should be used with caution. Aphid growth rates are usually measured using individual aphids or small groups of aphids, often the progeny of a single female. Young (typically newborn) aphids are removed from their host plants, weighed on a microbalance and returned to the host plants for the desired time period, after which they are removed and re-weighed. Aphid growth rates are a function of birth weight (Dixon et al., 1982; Carroll and Hoyt, 1986), as is weight loss when aphids are starved (e.g. Brough and Dixon, 1990). Since large aphids grow faster than small aphids (Dixon, 1998), measurements of aphid growth rates must correct for differences in initial weight. Both MRGR and RGR compensate for the increasing mass of the insect as it grows, since they are based on the logarithmic weight gain of the aphid. The formula used to determine MRGR is based on an equation originally used by plant scientists (Radford, 1967): MRGR ( mg / mg /day) = (log W 2 − log W 1 ) / t 2 − t 1 (6.1) where W1 = weight at the first weighing, W2 = weight at the next weighing, t2 − t1 = the time (in days) between first (t1) and second (t2) weighing. RGR is measured over the development time (D) of the aphid (i.e. from birth to the final moult but before the onset of reproduction), and therefore takes the effects of host-plant quality and maternal effects (such as ovariole number) into consideration: RGR ( mg / mg /day) = (log W 2 − log W 1 ) / D (6.2) Two major experimental drawbacks are associated with using MRGR or RGR to measure aphid performance. First, it is usually difficult to manipulate young aphids in the field. Second, a very accurate microbalance is required. Newborn aphids can weigh as little as 30 µg (Dixon, 1998), and hence small inaccuracies in the measurement of initial weight can have large effects on the final value of MRGR or RGR because of the logarithmic nature of insect growth. A simple solution is to measure groups of aphids and use the average initial and final weights to determine growth rates. MRGR and RGR are, however, very simple ways to investigate treatment effects on aphid performance since they may be measured over as little as two days (van Emden and Bashford, 1969; Adams and van Emden, 1972). A second benefit of both MRGR and RGR is that they involve relatively little disturbance of the aphids (Adams and van Emden, 1972). The newly moulted adults produced after RGR is measured can be returned easily to the host plant to measure achieved fecundity and rm, or dissected to investigate treatment effects on potential fecundity. Developmental rates Developmental rates are determined by recording the time period between particular events (for example, from birth to adult) and reporting the results as the reciprocal of the data (Berg, 1984; Carroll and Hoyt, 1986; Lamb et al., 1987; Lamb, 1992; Cabrera et al., 1995). Most aphids pass through four nymphal instars (Dixon, 1973) before moulting to the adult stage (although some species, such as Rhopalosiphum nymphaeae, may have five instars, depending on rearing temperature (Rohita and Penman, 1983; Ballou et al., 1986). Instar duration may also be a useful measure of development, particularly in studies investigating treatment effects on the vulnerability of aphids to natural enemies that preferentially attack specific instars (Ives et al., 1999; Chau and Mackauer, 2001). Measurements of development are particularly useful in studies investigating 138 C.S. Awmack and S.R. Leather treatment effects on aphids reared in field situations, as the aphids do not need to be removed from the host plant to be weighed. Developmental times or rates are particularly useful when predictions about treatment effects on future population growth rates are required, since they are an integral component of the intrinsic rate of increase, rm. The intrinsic rate of increase, rm The intrinsic rate of increase, rm (Wyatt and White, 1977), relates the fecundity of an individual aphid to its development time: rm = (ln Md × c ) / D (6.3) where Md is the number of nymphs produced by the adult in the first D days of reproduction after the adult moult. The constant, c, has a value of 0.738 and is an approximation of the proportion of the total fecundity produced by a female in the first D days of reproduction. It is obvious from this equation that a small change in development time will have a greater effect on rm than an increase in fecundity of a similar magnitude. Although rm has limitations (Awmack et al., 1997), it is a favoured way of estimating population growth as it is much easier than counting the thousands of aphids that are likely to occur in a real population. Experimental Techniques Aphid cages Although the growth and development of individual aphids has been recorded under unrestricted field conditions (Cannon, 1984), most studies have used individual aphids reared in the laboratory or in controlled environments (van Emden, 1972; Dixon, 1998). Since newborn aphids are so small, it has become standard protocol to cage either an individual aphid or a group of aphids on the host plants. The most commonly used aphid cage is the clip cage, originally designed by MacGillivray and Anderson (1957). Clip cages may range in size from those that cover a few cm2 of the leaf to those that enclose whole leaves. As not all leaves used by aphids are flat, tubular cages based on gelatin capsules can also be used. These cages slip snugly over the entire leaf or stem, but can be opened easily without disturbing the occupants (Fisher, 1987). Alternatives to clip cages include gauze sleeves that can be used to confine aphids to specific parts of the plant (for example, stems). Whole plant cages, usually constructed of PVC tubes, open at the bottom and covered with gauze at the top, can be used to monitor colonies or aggregations of aphids that need free access to all or part of the plant (Markkula and Rautapaa, 1963). Two clip cages, suitable for use on many crop species, are shown in Fig. 6.2. The Type I cage is the typical clip cage (MacGillivray and Anderson, 1957). The second cage (Type II) uses soft foam (Plastozote®, Watkins and Doncaster, UK) to minimize damage to the leaf surface and can also be supported with a standard plant stake, reducing strain on the leaf petiole, and can be opened without disturbing the aphids inside (Awmack, 1997; D. Huggett, personal communication). Although some aphid species appear not to be adversely affected by regular removal from their host plants (Newton and Dixon, 1990b), disturbance should be kept to a minimum when measuring growth rates, because aphids may take several hours to locate a suitable phloem element and commence feeding (Tjallingii, 1995). Frequent disturbance may therefore lead to low estimates of aphid performance. The settling and feeding behaviour of aphids may also be affected by the treatment under investigation. Aphid settling and feeding behaviour may be affected by components of host-plant quality such as concentrations of plant defensive metabolites (Zehnder et al., 2001), nutrient availability, and water stress (Ponder et al., 2001), and by environmental factors such as elevated CO2 atmospheres (Awmack et al., 1996). Feeding and settling behaviour may also vary according to the prior experience of the aphids used (Ramirez and Niemeyer, 2000) and even between 139 Growth and Development Type 1 Type 2 Fig. 6.2. Clip cages suitable for measuring aphid growth and development. individuals within a species (Bernays and Funk, 2000). Disadvantages of aphid cages Many clip cage designs have been criticized because they interfere with leaf gas exchange and may damage the leaf surface, changing leaf quality in comparison with uncaged leaves (Crafts and Chu, 1999). The effects of larger cages on aphid performance have been documented from field experiments, as sunlight can be concentrated on to the plants leading to temperatures within the cage that are considerably higher than ambient external temperatures (Woodford, 1973). In this case, large, muslin-covered frames may be placed over the plants to minimize fluctuations in microclimate. An equally serious concern associated with the use of cages to measure aphid growth and development is that aphid performance within cages may not reflect the performance of uncaged aphids. Figure 6.3 shows the adult weight, development time, and 7-day fecundity of Rhopalosiphum padi (bird cherry–oat aphid) reared without cages (control), in clip cages, and in PVC tubes on oat seedlings (Avena sativa cv. ‘Aster’). The data clearly show that clip cages and PVC tubes had an adverse effect on the adult weight of the aphids, and a small but significant effect on development time. However, cages had no significant effects on 7-day fecundity (S.R. Leather, unpublished results). Adult weight and 7-day fecundity were positively correlated when the aphids were reared in clip cages (r2 = 0.218, P < 0.05) and PVC tubes (r2 = 0.417, P < 0.01), but not when they were reared on uncaged control plants (r2 = 0.005). Cages therefore affected not only the performance of R. padi, but also the relationships between adult weight and potential or achieved fecundity. This example demonstrates clearly the problems inherent in comparing data collected using insect cages to data collected when the aphids are reared in more natural environments and able to select feeding sites. 140 (a) C.S. Awmack and S.R. Leather 1.0 mg 0.8 0.6 0.4 0.2 0.0 Control (b) Cage PVC 8 Days 6 4 2 0 Control Cage PVC (c) 60 N 40 20 0 Control Cage PVC Fig. 6.3. Performance of the aphid Rhopalosiphum padi reared without cages, in clip cages, and in PVC tubes on oat seedlings (Avena sativa cv. ‘Aster’). (a) Adult weight (mg) (F = 8.36, df = 2/45, P < 0.001). (b) Development time (days) (F = 6.10, df = 2/45, P < 0.001). (c) 7-day fecundity (F = 0.86, df = 2/45, P > 0.05). All data are presented as the means of 16 replicates and are shown with ± the standard error of the mean. Factors Affecting Aphid Growth and Development Aphid growth and developmental rates are affected by a wide range of both intrinsic and extrinsic factors such as diet quality (e.g. Watt and Dixon, 1981; Gruber and Dixon, 1988; Tsai and Wang, 2001; Vacanneyt, 2001), plant growth stage (Zhou and Carter, 1992), the abiotic environment (Kenten, 1955; Dean, 1974; Walgenbach et al., 1988; McVean and Dixon, 2001), maternal experience (Johnson, 1965; Dixon and Glen, 1971; Chambers, 1982; Kidd and Tozer, 1984), and maternal morph (Leather, 1989). In this section, some of the factors affecting the reliability of aphid growth rates as predictors of the performance of aphid populations are outlined, and some of the most common factors affecting the differences between measures of aphid growth and developmental rates and the performance of natural aphid populations are discussed. A key consideration in these types of studies is that aphids used to measure growth and developmental rates in greenhouse-based investigations tend to have been raised at low densities at constant temperatures while natural aphid populations interact with a variable biotic and abiotic environment. Genetic variability may also contribute to the variability inherent in natural populations: in many studies, the aphids are derived from parthenogenetic lineages, produced by a single parthenogenetic female. Natural populations are rarely as uniform (except in the case of pests of greenhouse crops, which may have very low levels of genetic diversity (Rochat et al., 1999). When apterous Sitobion avenae (grain aphid) were reared on oats (A. sativa), pink individuals developed more quickly than green individuals, highlighting the need to use multiple aphid genotypes (Araya et al., 1996). Similarly, many aphid species such as Myzus persicae (peach–potato aphid), Aphis craccivora (cowpea aphid) (Edwards, 2001), Sitobion miscanthi and Sitobion near fragariae (Sunnucks et al., 1998), Acyrthosiphon pisum (pea aphid) (Sandström, 1994), and Phorodon humuli (damson–hop aphid) (Lorriman and Llewellyn, 1983) show genetic variation in their growth and developmental rates, even when reared on hosts of similar quality. Aphids used in laboratory-based studies frequently are confined to a specific part of the host plant (for example, in clip cages) 141 Growth and Development and reared at very low densities in the absence of other pests and diseases. Many aphid species prefer specific host-plant parts, and experiments that cage aphids on the ‘wrong’ part of a plant do not give an accurate representation of the performance of natural aphid populations. Hopkins et al. (1998) showed that even though M. persicae prefers senescing leaves at the base of its host plants (three Brassica cultivars, differing in their glucosinolate content) while Brevicoryne brassicae (cabbage aphid) prefers young leaves, regardless of the cultivar of the host plant, aphid performance was unaffected by plant defensive chemistry. Similarly, Williams (1995) investigated the impacts of host plants of different ages, with and without Beet yellows virus infection, on M. persicae, and showed that performance was better on younger leaves than on old leaves, and that virus infection increased aphid performance. Table 6.1 shows the fecundity and development time of S. avenae when reared on different parts of oat and wheat plants (from Watt, 1979). Factors Affecting the Reliability of Size ¥ Fecundity Relationships While aphid growth and developmental rates can often be used to predict future fecundity (and hence population growth rates), some treatments affect the reliability of these relationships (Leather, 1988; Awmack and Leather, 2002). Aphid size is determined predominantly by the quality of Table 6.1. Effects of rearing on different host plant parts of winter wheat (cv. ‘Maris Huntsman’) on the fecundity and development of the aphid Sitobion avenae. Ears Young leaves Lower leaves Flag leaves Senescent leaves 7-day fecundity Time to adult moult (days) 41.7 26.5 13.3 17.6 14.5 8.5 8.4 10.4 9.8 9.0 the larval host plant (Banks and Macaulay, 1964; van Emden and Bashford, 1969; Leather and Dixon, 1982; Acreman and Dixon, 1989; Gange and Pryse, 1990; Caillaud et al., 1994), although the quality of the adult’s host plant is also important (Markkula and Rouka, 1970; Watt, 1979; Leather and Dixon, 1981; McLeod et al., 1991) since aphids continue to mature offspring after the final adult moult. Some aphid species vary allocation of their resources between reproductive and somatic tissues, e.g. Myzocallis boerneri (Turkey oak aphid) (Sequeira and Dixon, 1996), Drepanosiphum platanoidis (sycamore aphid) (Douglas, 2000), Megoura viciae (vetch aphid) (Brough and Dixon, 1990), and S. avenae (Helden and Dixon, 1998). Thus, insect size may not necessarily be a reliable predictor of future fecundity (Leather, 1988). Maternal effects (i.e. the host plant on which the parent of the reproducing aphid was reared) may also affect aphid size × fecundity relationships (Leather, 1989; Messina, 1993). While generally there are strong and positive correlations between aphid growth and developmental rates and the intrinsic rate of increase, rm, it has been shown that this relationship varies with both the species of aphid under investigation and the growth stage of the host plant. Guldemond et al. (1998) demonstrated that the relationship between MRGR and rm varied with both the species of aphid investigated (Aphis gossypii – cotton or melon aphid, or M. persicae) and the growth stage of the host plant (chrysanthemum, Dendranthema x grandiflorum). If experimental treatments affect such relationships (e.g. Kerns and Gaylor, 1992; Sarao and Singh, 1998), individual growth rates need not reflect population growth rates. While individuals with high MRGRs also tend to have high RGRs, the two growth rates need not be the same for any individual aphid/treatment combination, and may not be the same throughout the entire nymphal development time. Some aphid species, such as S. avenae, have high nymphal growth rates during early instars (Newton and Dixon, 1990b), while growth rates often decrease in later instars as the 142 aphid switches resources to embryo maturation (Kindlmann and Dixon, 1989, 1992; Newton and Dixon, 1990b). Instar duration frequently varies, with the fourth instar being significantly longer than the first three (Kieckhefer et al., 1989; Araya et al., 1996; Dixon, 1998), particularly in the case of individuals destined to be alate (Newton and Dixon, 1990a). Treatments may therefore not affect all aphid instars equally, and early aphid instars may also be more sensitive to changes in the quality of their host plants than later instars. When Schizaphis graminum (greenbug) was reared on maize (Zea mays) or sorghum (Sorghum bicolor) cultivars, the host plant significantly affected the development time of the first and second aphid instars, but not the third or fourth (McCauley et al., 1990). Similarly, the development of first-instar A. pisum was slower on aphid-resistant red clover (Trifolium pratense) cultivars than on susceptible cultivars, but since later instars took less time to develop on resistant cultivars, the total developmental time was unaffected (Zeng et al., 1993). Not all aphids show this variation in larval growth rates between instars; growth rates of M. persicae remained constant throughout larval development (van Emden, 1969). Since many other insect groups show similar declines in growth rates as they develop (Scriber and Slansky, 1981), M. persicae may be an exception, rather than the rule. Changes in the reproductive rates of individual aphids may also affect the reliability of assumptions about the relationships between growth rates and adult size and fecundity, as not all aphids produce nymphs at a constant rate throughout their adult life. Zeng et al. (1993) showed that A. pisum produced more nymphs during the daytime than at night, demonstrating that measurements of fecundity must take place over at least 24 h. Many aphid species produce a rapid ‘burst’ of reproduction shortly after the final moult, and then show a reduced rate of reproduction for the remainder of their adult life (Dixon, 1998). Aphid reproductive strategies may also vary according to predictable changes in plant quality (Leather, 1987) or unpredictable C.S. Awmack and S.R. Leather environmental conditions such as starvation (Leather et al., 1983; Ward et al., 1983; Brough and Dixon, 1990; Kouame and Mackauer, 1992; Gruber and Dixon, 1988). The first-born nymphs may not be representative of the entire progeny (Dixon et al., 1993) because birth order may affect subsequent performance. The first-born nymphs of A. pisum are smaller than those born on subsequent days (Murdie, 1969b), but after about 7 days of reproduction, nymphal weight begins to fall and by 14 days the nymphs born are smaller than those born in the first 2 days. Similarly, first-born M. persicae nymphs show greater cold hardiness than later born nymphs (Clough et al., 1990). While rm includes both developmental and reproductive rates, and is often a more reliable measure of aphid performance than MRGR or RGR, it has several disadvantages. Measurements of rm are very labour intensive, and the risk of losing replicates (and hence statistical power) increases with the duration of the experiment. rm is also a less sensitive measure of small, but biologically significant, changes in aphid performance (Lykouressis, 1984). A further disadvantage of rm is apparent when treatments affect adult longevity since it is inaccurate when a treatment affects longevity but not development time (Sumner et al., 1986). The fundamental assumption underlying the rm equation is that a reproducing female will produce 95% of her progeny in the first D days of reproduction (Wojciechowicz-Zytko and van Emden, 1995). If this assumption is not met (for example, if a female dies after only a few days of reproduction), values of rm overestimate the contribution of individual aphids to the growth of the population. Population growth rates of the pea aphid A. pisum reared on Vicia faba (broad bean) and exposed to a neem-based insecticide depended on the age at which the aphids were exposed to the insecticide. When individual A. pisum were exposed to this insecticide from birth, the population growth rates were negative. However, when A. pisum were exposed as adults, there was no effect of this insecticide on rm, highlighting the need to determine treatment effects on all 143 Growth and Development aphid life stages (Stark and Wennergren, 1995). Difference between Nymphs Destined to be Apterous and Alate Measurements of aphid growth and developmental rates should also take into account differences between nymphs destined to be apterous adults and nymphs destined to be alate adults. Although alate aphids have a longer development time than apterae (Tsumuki et al., 1990; Araya et al., 1996), in some species (e.g. R. padi), alatae have greater longevity than apterae (Foster et al., 1988) and may therefore make a greater contribution to population growth rates. However, nymphs that develop into alate adults allocate a smaller proportion of their resources to reproduction (Newton and Dixon, 1990b) since they must produce wings and the flight muscles needed to power them (Dixon, 1998). As a result, alate aphids typically have lower fecundity than apterae (Elliott et al., 1988; Newton and Dixon, 1990b; Collins and Leather, 2001). Alate aphids may also have a lower ovariole number (Dixon and Dharma, 1980; Leather, 1987) than apterae, differ in their amino acid and carbohydrate metabolism (Tsumuki et al., 1990), and have greater lipid reserves (Febvay et al., 1992). These differences may therefore mean that alatae respond differently to experimental treatments such as starvation or environmental factors; e.g. Garsed et al. (1987) showed that the fecundity of alate Aphis fabae (black bean aphid) reared on V. faba increased as light levels increased, while that of apterae did not. Alata production is stimulated by crowding (Watt and Dixon, 1981; Bergeson and Messina, 1997; Dixon, 1998; Williams and Dixon, Chapter 3 this volume), although the proportion of the population developing into alatae may vary among populations of the same aphid species on different host plant species (Bommarco and Ekbom, 1996). Alata production may also be stimulated on virus-infected host plants (Blua and Perring, 1992), and may either be stimulated (Liu and Wu, 1994) or suppressed (Parish and Bale, 1990) by low temperatures, depending on the species of aphid involved. As apterous progeny of alatae may also have lower ovariole numbers than the progeny of apterae (Leather, 1987), the effects of experimental treatments on population growth rates may persist for more than one generation. Treatments that affect reproductive rates but not development times or growth rates also affect the reliability of rm. S. avenae reared on winter wheat (Triticum aestivum) at double-ambient CO2 had greater fecundity than at ambient CO2, but their development times were unaffected (Awmack et al., 1996). In a similar experiment, when Aulacorthum solani (glasshouse and potato aphid) was reared on either broad bean or tansy (Tanacetum vulgare) at elevated CO2, development time decreased and rm increased on tansy but not on bean, while fecundity increased on bean but not tansy (Awmack et al., 1997). In these examples, rm is unlikely to be a reliable predictor of aphid population growth rates as aphid fitness parameters appear to become uncoupled on exposure to this novel environment. The relationships between adult size and rm may also become uncoupled if the aphid is exposed to starvation and resorbs embryos to release essential nutrients (Ward and Dixon, 1982; Brough and Dixon, 1990). Thus, a fundamental assumption underlying the rm equation is that experimental conditions remain constant throughout the life of the aphid. Temperature Aphid responses to temperature are similar to those of other insects. Most aphid species show a strong linear relationship between temperature and growth or development within a range of approximately 7 and 25°C (Campbell et al., 1974; Frazer and Gill, 1981), followed by a decline at increasing temperatures. The exact shape and range of the curve depends on both the aphid species and the geographic origin of genotypes within the species (Auclair and Aroga, 1987; Lamb and Mackay, 1988; Akey and 144 C.S. Awmack and S.R. Leather Butler, 1989), and may also be genotypespecific within a population (Lamb et al., 1987). Aphids reared at high temperature may also grow into small adults containing fewer embryos (Leather and Dixon, 1982; Collins and Leather, 2001), or have a poor ability to maintain embryo maturation after the adult moult (Carroll and Hoyt, 1986). High temperatures may also affect the slope of the relationship between adult weight and embryo number, making predictions of fecundity from adult size unreliable (Carroll and Hoyt, 1986). Plant quality may also modify the impacts of temperature on aphid growth and development (Leather and Dixon, 1982; Acreman and Dixon, 1989), as may abiotic factors such as wind (Walters and Dixon, 1984). Other authors (Liu and Perng, 1987; Kieckhefer et al., 1989; Xia et al., 1999) have also demonstrated that aphid performance decreases when aphids are reared at fluctuating temperatures rather than the equivalent constant temperature. In contrast, Zhang et al. (1991) showed that the population growth rate of R. padi was greater at fluctuating temperatures than at constant temperatures. Hodgson and Godfray, 1999; Awmack and Harrington, 2000; Bosque and Schtozko, 2000; Awmack et al., 2004), although in some cases, aphid density has no effect on population growth rates (e.g. Messina, 1993). Some plant species may also exhibit a hypersensitive response, which is only apparent above threshold aphid densities (Lyth, 1985; Belefant-Miller et al., 1994). In contrast, many aphid species show enhanced performance when they are reared in groups, rather than singly (Way and Cammell, 1970; Dixon and Wratten, 1971) because they are able to divert nutrients from other plant tissues more effectively (Sandström et al., 2000). Interspecific competition between aphid species exploiting the same host plant may also reduce population growth rates (Fisher, 1987; Moran and Whitham, 1990; Thirakhupt and Araya, 1992; Gianoli, 2000), as may the presence of other herbivores, either directly (Masters, 1995) or via apparent competition involving shared natural enemies (Muller and Godfray, 1997) and plant diseases (Coleman and Jones, 1988; Blua and Perring, 1992; Castle and Berger, 1993). Population-scale factors Conclusions Although a detailed discussion of population-scale effects on aphid performance is beyond the scope of this chapter, population density also has significant effects on aphid performance. At high population densities, aphids frequently produce alatae and leave the host plant (e.g. Watt and Dixon, 1981). Crowding (and intraspecific competition) may also reduce the per capita rate of reproduction: as host-plant resources become limiting, the population growth rate decreases (e.g. Farid et al., 1998; Although it is tempting, and perhaps technically easier, to use the performance of individual aphids as an indicator of aphid performance, perhaps the most reliable way to predict aphid performance is to investigate population size. Comparisons of rm with population size (a much simpler and more reliable measure of aphid population growth rates) also take crowding and sink induction into account (Larson and Whitham, 1991; Sandström et al., 2000) and are much easier to use (Lykouressis, 1984). References Acreman, S.J. and Dixon, A.F.G. (1989) The effects of temperature and host quality on the rate of increase of the grain aphid (Sitobion avenae) on wheat. Annals of Applied Biology 115, 3–9. Adams, D. and Douglas, A.E. 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