Annu. Rev. Entomol. 1994. 39:155-78 Copyright © 1994 by Annual Reviews Inc. All rights reserved

BIONOMICS AND

MANAGEMENT OF

ANASTREPHA

M. Aluja Instituto de Ecologfa, A.C., Apartado Postal 63, 91000, Xalapa, Veracruz, Mexico

KEY WORDS: Diptera, Tephritidae, biosystematics, zoogeography, biology, ecology, life strategies, behavior, pest management

PERSPECTIVES AND OVERVIEW

Flies of the genus Anastrepha (Diptera: Tephritidae) are among the world's most devastating agricultural pests. At the same time, they display remark­able ecological and behavioral characteristics, which have served as models in the development of general theories on insect mating systems and the physiology of host marking. Anastrepha species are endemic to the New World and restricted to tropical and sUbtropical environments. The genus's range extends from the southern US to northern Argentina and includes most of the Caribbean Islands. There are 184 described Anastrepha species. The great majority of these are poorly known biologically, and knowledge is basically restricted to seven economically important species: Jraterculus, grandis, ludens, obliqua, serpentina, striata, and suspensa. Much of what we know today about Anastrepha is based on very thorough studies of its basic biology carried out between 1900 and 1944 (13, 53, 85, 115, 188; see also 21, 45, 167). More recently, work has concentrated on phylogeny and taxonomy, behavioral ecology, chemical ecology, demography, and species surveys. Anastrepha control has in many ways been conceptually and technically stagnant for the past 35 years: poisoned bait sprays and McPhail traps are still standard procedure even though such methods have been proven ineffective and environmentally questionable.

Here, rather than providing an all-encompassing review of the voluminous literature on Anastrepha, I attempt to expose the reader to a balanced

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selection of poorly advertised studies from Latin America as well as work published in mainstream journals. I place special emphasis on recent ad­vances in phylogeny, taxonomy, distribution, host-plant relationships, de-' mography, population genetics, behavioral ecology, and some new control methods and management approaches.

BIOSYSTEMATICS AND ZOOGEOGRAPHY

Taxonomy and Phylogeny

Anastrepha belongs to the family Tephritidae (true fruit flies) and was recently placed in the tribe Toxotrypanini of the subfamily Trypetinae (67). Many species of Anastrepha were first described in other genera such as Dacus, Pseudodacus, Lucumaphila, Phobema, and Trypeta (132, 185). To date, the most thorough revision of the genus is Stone's 1942 monograph (188), which was partially updated by Steyskal (184), Zucchi (196), Foote (66), Norrbom (132), and Hernandez-Ortiz (83). One should note that the taxonomy of Anastrepha is based almost exclusively on the adult female. Only recently have efforts been directed toward identifying males (132). The immature stages of Anastrepha are poorly known. So far, descriptions of third-stage larvae have appeared for only the following species: A. bistrigata, A. consobrina, A. distincta, A. Jraterculus, A. grandis, A. interrupta, A. limae, A. ludens, A. nunezae, A. obliqua, A. pal/ens, A. sagittata, A. serpentina, A. striata, and A. suspensa (13, 23, 42, 49, 64, 75, 82,132, 142, 181-184, 194). Eggs have been described for the following species: A. atrox, A. cordata, A. leptozona, A. iudens, A. nigrijascia, A. obliqua, A. pal/ens, A. pittieri, A. serpentina, A. striata, and A. suspensa (57, 98, 132).

Cladistic analysis and immunological data support the placement of Anastrepha in a monophyletic group with the genus Toxotrypana (67, 93, 132). Anastrepha is most probably of South American origin (83). Morgante et al (119) suggest that many species of Anastrepha are relatively young and that speciation has been rapid and recent. The first serious attempts at studying the phylogenetic relationships of Anastrepha were recently pub­lished by Norrbom (132-134) and Norrbom & Kim (136). These authors divided the genus into 18 preliminary groups mainly based on morphological characters: benjamini, chiclayae, cryptostrepha, dacijormis, dentata, doryphoros, !raterculus, grandis, leptozona, mucronota, pseudoparaUela, punctata, schausi, serpentina, spatulata, striata, ramosa, and robusta. Given the size of the group, the rate at which new species are being described and the presence of many cryptic species, future infrageneric classifications will have to be based, in my view, on a combination of criteria such as

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ANASTREPHA BIONOMICS AND MANAGEMENT 157

morphology, cytogenetics, isozyme studies, molecular (DNA) analyses, and further knowledge of the ecology and behavior of the group.

Zoogeography

Anastrepha is endemic to the New World and is restricted to tropical and subtropical environments (83). Its range covers part of North America (southern Florida, the Rio Grande Valley in Texas, and almost all of

Mexico), Central and South America (except Chile and southern Argentina), and most of the Caribbean Islands (84). A thorough zoogeographic analysis

at this stage is limited by the fact that the exact distribution of most species of this genus has not yet been established. Nevertheless, according to

Norrbom & Foote (135) the distribution at the species group level in Anastrepha lacks a discernible pattern, and most of the subgroups are

widespread. Certain species (e.g. A. alveata. A. acris. A. bicolor. A. tripunctata) seem to be associated with tropical deciduous forests with long dry seasons, while others (e.g. A. cordata, A. crebra. A. bahiensis, A. tumida) are associated with tropical rain forests (83). Regional studies in Mexico (13, 83), Costa Rica (92), Venezuela (41), Peru (95, 96), Brazil (40, 126, 196, 197), and Argentina (25) reveal that the range of species such as A. jraterculus. A. obliqua, and A. striata is remarkably large. Overall, of the 184 Anastrepha species described so far, 43% are only found in South America, 15% are restricted to Central America (including Panama), 4% are found only in North America (Mexico and the US), and 1 % are found only in the Lesser and Greater Antilles (84).

BASIC BIOLOGY

Life Cycle and Life Strategies

The basic life cycle is very similar among all Anastrepha species for which the biology is known. In the majority of species, the females deposit their eggs in the epi- or mesocarp region of ripening host fruit. Some species have an extremely slender and long ovipositor (e.g. A. sagittata), which may allow them to oviposit into seeds instead of the pulp (13, 132). Depending on the species, eggs are laid singly or in clutches. For example, A. obliqua lays one egg per oviposition (43) whereas A. grandis can lay clutches of up to 110 eggs (52). Larvae go through three instars before

leaving the fruit and burrowing into the ground to pupate (on occasion larvae pupate inside the host fruit). The larvae of most species feed exclusively on the fruit pulp, although A. manihoti feed on stems and buds (140), and at least some species in the A. serpentina. A. leptozona, A. daciformis, and A. cryptostrepha groups feed on pulp and seeds or only on

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seeds (13, 114, 132, 137, 188). The rate of immature-stage development is determined to a great degree by environmental factors such as temperature (14, 99, 165) and rearing medium (43, 100), and is sexually dimorphic (173). Pupation depth is greatly influenced by soil pH and type (e.g. sand, clay) and the degree of soil compaction and humidity (28, 53, 115). Exit of larvae from fruit is determined by fruit characteristics (e.g. pH, internal temperature, degree of rotting) and by physical signals experienced when ripe fruit drops to the ground or is exposed to rain (53, 115). After emergence, adults crawl to a sheltered spot nearby until their wings unfold and dry. Before becoming sexually active, adults go through a maturation period during which they need to feed regularly on carbohydrates and water to survive and on protein sources to allow for gonad maturation (45).

Anastrepha species typically inhabit highly variable environments (i.e. seasonal, unpredictable, or ephemeral in time, and patchy or isolated in space) where they live in close association with their host plants. Zw6lfer (198) has indicated that they are opportunistic, multivoltine, and highly mobile; exhibit high reproductive potential; and are usually long-lived. These

attributes are broadly applicable to the genus as a whole, but we simply know too little about Anastrepha field ecology to utilize hard-and-fast categories at this stage. For example, strictly monophagous species such as A. crebra, in which adult emergence is closely timed with host-plant fruiting phenology, are probably univoItine and undergo obligatory diapause(V. Hernandez, personal communication). In contrast, polyphagous species, such as A. ludens or A. obliqua, may be able to cope with resource patchiness and ephemerality by going through periods of estivation and by tracking seasonal hosts scattered over large areas by moving effectively from habitat to habitat (5).

Host-Plant Relationships

Norrbom (132) indicates that Anastrepha is clearly not old enough to have strictly coevolved with its hosts but suggests that host-plant associations of Anastrepha appear to be at least partially correlated with phylogenetic relationships within the genus. The fact that the cryptostrepha and daciformis groups (primitive Anastrepha) are strongly associated with sapotaceous hosts suggests that the Sapotaceae may have been ancestral hosts for Anastrepha (132). An interesting feature of Anastrepha host-plant relationships is the close association between certain species and particular plant taxa: for example, the chiclayae and pseudoparallela subgroups are associated almost exclusively with the genus Passijlora (Passifloraceae). The serpentina, zeteki, daciformis, dentata, robusta, and leptozona subgroups are mainly associated with the family Sapotaceae (132).

Host-plant records are only known for approximately 39% of all reported

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ANASTREPHA BIONOMICS AND MANAGEMENT 159

Anastrepha species (137). Furthermore, seven economically important spe­cies account for -70% of all reported Anastrepha host records (132). Unfortunately, a considerable number of these records are incidental or the result of misidentifications, were obtained under laboratory conditions, or are related to cultivated and, in many cases, exotic hosts (8, 137). Almost no information is available on wild host plants in unperturbed environments. Based on host-plant lists, Anastrepha species have been classified as mo­nophagous, stenophagous, oligophagous, and polyphagous (2). Such clas­sification, though useful in general terms, is confounded by the fact that many host-plant associations are a local phenomenon (i.e. polyphagous species behave as monophagous or stenophagous species in certain areas).

The metabolic pathways used by larvae to detoxify many deleterious compounds found in food plants are poorly understood. For example, larvae of A. acris feed on Hippomane mancinella fruit (Euphorbiaceae), which contains alkaloids close to the virulent toxin strignin (M Aluja, G Siller &

R Rosiles, unpublished data). Matioli et al (112, 113) have found alcohol dehydrogenase (AD H)-specific activity in larvae of several Anastrepha species and suggest that selective pressures for enhanced ethanol metabolism have occurred in the evolutionary history of Anastrepha species.

Nutrition

As is the case with other tephritids, immature and adult Anastrepha require amino acids, vitamins, minerals, carbohydrates, and water for development, reproduction, and survival (76). Such nutrients are obtained in nature from liquids oozing from overripe or damaged fruit, bird feces, leaf and fruit surfaces, and rain drops (5, 13). Adult flies can survive for long periods of time on carbohydrates alone, but optimal male salivary gland development (important for sexual pheromone production) and female ovary development depend on extrinsic sources of protein (62, 68, 76, 116, 162). Laboratory studies with A. ob/iqua have shown that the optimal protein/carbohydrate ratio for ovarian development is 3:5 (63). Furthermore, a mixture of all amino acids at a concentration of 15% mixed with vitamins and sucrose yields the greatest number of eggs and the highest egg eelosion percentages (27). When the larvae feed on seeds, as in A. cordata, female and male gonad development is completed shortly after emergence and apparently without the need for protein intake (M Aluja & V Hernandez, unpublished data).

Associations with Bacteria

The· role of bacteria in the nutrition and survival of tephritid larvae and adults is best known in Bactrocera and Rhagoletis species (65). As is the case with other tephritid genera, Anastrepha have anatomical adaptations

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for housing microorganisms, such as gastric ceca in larvae and an esophageal diverticulum (bulb) in adults (175). Bacteria housed in the gut of Anastrepha and other fruit fly larvae and adults are known as fruit fly type bacteria

(Enterobacteriaceae) (65). Murillo et al (123) and Solferini (175) have isolated bacteria such as Acinetobacter lwojfi, Enterobacter agglomerans, Enterobacter cloacae, Enterobacter gergoviae, Klebsiella oxytoca, Klebsiella pneumoniae, Klebsiella ozanae, Proteus mirabilis, Proteus vul· garis, Proteus ratgeri, Pseudomonas jluorescens, and Pseudomonas cepaceae from the gut and eggs of A. fraterculus, A. obliqua, A. bistrigata, and A. pickeli. In a previous study on A. ludens and A. suspensa, the bacteria Micrococcus sp., Erwinia sp., Pseudomonas sp. and Pseudomonas ovalis were also identified, but such findings could have been the result of artificial laboratory conditions (26, 156). What role bacteria play in the biology of adult flies is still poorly understood. Anastrepha adults regurgitate food on leaf surfaces shortly after feeding on fruit juices (6, 10). Fletcher, interpreting a similar behavior in Bactrocera spp., suggested that flies inoculate the surface of fruit and utilize the resulting bacterial colonies as protein sources (65 and references therein). Bacterial syinbiotes may also play an important role in synthesizing essential amino acids, in the break· down and digestion of fruit tissues, in detoxifying plant.defense chemicals, and in suppressing pathogenic fruiHot microorganisms (65).

POPULATION ECOLOGY

Population Genetics

Both the cytogenetics of individual species and population genetics of Anastrepha have been studied. Karyotype analyses have been performed on A. barnesi, A. bistrigata, A. fraterculus, A. obliqua, A. pickeli, A. pseudoparallela, A. serpentina, A. ludens and A. striata (36, 176). These studies have revealed that the typical chromosome number in Anastrepha is 2n = 12. Variations in the diploid number have been detected in a few species, particularly those having an XX:XO or X1X,X:zX2:X,X2 Y mecha· nism of sex determination (177).

Population genetics studies in Anastrepha have been restricted to one species: A. jraterculus. This species is polyphagous, widely distributed, and exhibits greater morphological variation than related species. According to several authors, it represents an unresolved complex of cryptic species (13, 118, 119, 180, 188). Four distinguishable karyotypes in Brazilian popula· tions have been described (176), but no host·dependent genetic differentiation is evident ( lOS, 106, 118). Recently, Steck (180) carried out an isozyme analysis that revealed sharp genetic discontinuities among A. jraterculus

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ANASTREPHA BIONOMICS AND MANAGEMENT 161

populations. For example, populations from northeastern Brazil, coastal Venezuela, Costa Rica, and Mexico were all very similar. In contrast, populations from southern Brazil, Andean Venezuela, and Peru were ge­netically distinct from the first group and possibly from each other as well. Steck (180) concludes that there is strong evidence that a complex of cryptic species is included in the nominal species A. Jraterculus.

Demography and Population Dynamics

Basic demographic parameters (e.g. preadult 'survival and development rates,

adult survival, gross and net fecundity) have been determined for A. ludens, A. obliqua, A. serpentina, and A. suspensa under laboratory conditions (43). Longevity and survivorship curves have also been described for A. sororcula and A. bistrigata (29). Adult survivorship curves of several species studied so far are close to type III curves. Age at first reproduction is strongly influenced by temperature and ranges from 8 to 20 days depending on the species (13, 43, 101). Food type, availability of water, fly size, and fly density influence life expectation and gross fecundity rates (101, 115). For example, in large vs small (mean pupal weight of 24 vs 18 mg, respectively) A. ludens, mean gross fecundity was 1597 vs 1450, respectively (101). Two points stand out from these studies: (a) gross fecundity rates and daily egg production under ideal laboratory conditions can be extremely high (e.g. 1000 eggs/female) and (b) adults can live for prolonged periods of time. For example, female and male A. ludens individuals can live as long

as 11 and 16 months respectively, under laboratory conditions (13, 53) and 12 months under field conditions (169).

Few studies in the vast Anastrepha literature examine the causes of mortality. Scattered efforts by early workers are incomplete and were carried out under laboratory conditions (13, 53, 115). Biotic mortality factors have been repeatedly identified, but the true impact on the population dynamics at the field level has never been determined. All immature stages (eggs, larvae, pupae) of Anastrepha are attacked by a series of indigenous parasitoids. Surveys in Mexico (9), Guatemala (60), Costa Rica (91, 193), Brazil (12), and Argentina (61, 124) indicate that the following indigenous hymenopterous parasitoids commonly attack Anastrepha spp.: Bracana­strepha anastrephae, Doryctobracon aerolatus, D. brasiliensis, D. craw­Jordi, D. zeteki, Opius hirtus, O. vierecki, O. tucumanus, O. argentinus (all Braconidae); Ganaspis carvalhoi, Eucoila pelleranoi (all Cynipidae); Odontosema anastrephae, Cothonaspis sp. (Eucoilidae); and Psylus sp. (Diaprididae). Some exotic parasitoids are also quite commonly encountered, including Diachasmimorpha longicaudata and Biosteres arisanus (Braconidae), Aceratoneuromyia indica (Eulophidae), and Pachycrepoideus vindemiae (Pteromalidae). Note that D. aerolatus and D. brasiliensis have

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been recovered from wild, native host plants in unperturbed environments (12, 144). Other biotic mortality factors are staphylinid beetles (e.g. Xenophygus analis, Belonuchus rufipennis) and ants (e. g. Solenopsis geminata) (2, 13) that attack both larvae and pupae. Pathogens have also been described. For example, Stigmatomyces aciurae fungi (ascomycetes) have been reported on A. striata adults collected in the field (79).

The most important abiotic mortality factors regulating population dy­namics are water and temperature. Too much or too little water causes both immatures and adults to die. Pupal desiccation in dried soil appears to be a major mortality factor. The first days after puparium formation are critical in this respect (115). Excessively dry conditions may reduce female fecundity and affect survival rate of newly emerged adults. Fruits exposed to rain decay faster and cause fIrst- and second-instar larvae to die. Excess water also hastens emergence from the fruit and concomitant puparium formation, and may reduce survivorship (13).

Adult Population Fluctuations

Based on studies in the US (131), Mexico (2, 70, 108), Belize (88), Costa Rica (90, 178), Colombia (138), Venezuela ( l09), and Brazil (104, 126), a picture is emerging indicating that adult populations in commercial orchards exhibit strong fluctuations from year to year and that these fluctuations are correlated with two factors: availability of host plants and climatic factors (especially rainfall). In monocrop orchards, population numbers typically reach a peak shortly after host fruit has ripened and crash when no hosts are available. In mixed orchards, fluctuations are probably dampened by the presence of alternative host plants. Trapping studies in orchards clearly show that even though up to 15 Anastrepha species are commonly captured, one or two are dominant. The degree of species dominance is influenced by ecological background (i. e. host plant species richness and diversity) and by altitudinal gradients. For example in Mexico, in mixed mango orchards at 1100 m above sea level, of 14 species captured, 30% of all individuals were A. ludens. At lower elevations (680 m), of 12 species captured, only 4% of individuals were A. ludens (143). Similar results have been reported for Costa Rica (90) and Brazil (126).

BEHAVIORAL ECOLOGY

Diel Rhythms of Activity and Resource Utilization Patterns

Under laboratory conditions, A. suspensa larvae have been shown to feed within fruit continously over a 24-h period (189). Behavioral partitioning of adult daily activities in Anastrepha (i.e. feeding, mating, oviposition,

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ANASTREPHA BIONOMICS AND MANAGEMENT 163

and resting) is quite different in the few species where information is available, i.e. A. ludens, A. Jraterculus, A. obliqua, A. serpentina, A. striata, and A. suspensa (5-7, 10, 13, 44, 80, 107). An exception to this seems to be time of adult emergence, which usually takes place during morning hours. For example, in A. ludens, 95.7% emerged between 0600 and 1000 hs (115) and in A. striata, 62.3% emerged between 0900 and 1200 hs (10). Other behaviors seem to follow species-specific patterns. Some species only call (Le. emission of a series of courtship sounds through wing fanning and release of a sexual pheromone) in the early morning, such as A. robusta (4), and others, such as A. ludens, A. grandis, and A. pseudoparallela (6, 44, 52, 146), restrict their calling activities to the late afternoon. Other species such as A. obliqua call both during the morning and afternoon (5). In the case of behaviors such as feeding, resting and oviposition, flies exhibit plasticity and can adapt to local microhabitat conditions (5). For example, A. obliqua females were never seen ovipositing in Spondias purpurea fruit during midday when ambient temperature could reach 45°C. However, if members of the same population were transferred to a more sheltered environment, oviposition was observed at precisely the period of time avoided under environmentally harsh conditions (5).

Trivial Movements and Migration

LARVAE Movement of larvae is related to the maturity of the host fruit. In citrus, A. suspensa larvae hatch in the flavedo (the part of the fruit where eggs are deposited) and move to the albedo and pulp as they grow and the fruit ripens (39).

ADULTS Research on Anastrepha adult movement includes laboratory stud­ies of tethered A. suspensa and A. ludens individuals aimed at establishing differences between irradiated and unirradiated or between wild and labo­ratory-reared insects (159), as well as field studies aimed at establishing short (intraorchard) and long-range (interorchard) displacement patterns. Under laboratory conditions, I-day-old flies had a significantly greater mean flight propensity than older flies (15, 160). Furthermore, individual flies differed significantly in their mean flight times when measured repeatedly over periods of 9-10 days. For example, in females the time between the shortest and longest flight increased 30-fold (15).

Christenson & Foote (45) have described long-range displacements and report that A. ludens fly -135 km from breeding sites in Mexico to invade citrus groves in southern Texas. Also, Shaw et al (169) trapped tepa-sterilized A. ludens up to 36 km from their release site. Work on this aspect of Anastrepha biology has been marred by the lack of a theoretical framework,

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and the literature is rife with anecdotal accounts about the "migratory" capacity of adult flies (3). In my opinion, the use of the term migration

should be avoided unless threshold responses by flies to vegetative stimuli are tested. At the orchard level, release-recapture studies with A. obliqua and A. ludens show that wind affects displacements of flies (mean fly movements are oriented to directions similar to those of the prevailing wind) (16, 17). Overall fly mobility after release is low (16, 30), but it is strongly correlated with environmental conditions. If flies are released in a place

where vegetation, food, water, and oviposition substrates are plentiful, they will not leave the release site (145). But if flies are released in an unfavorable

environment (e.g. dry conditions, lack of host plants, or adult food sources)

they will quickly leave the release site (58, 169). Within a certain area, Anastrepha species such as A. obliqua or A. ludens are known to move back and forth between native vegetation and orchards or between two habitats that harbor two sets of essential resources. Such patterns have been documented using release-recapture methods (58) and through direct obser­

vations (5), but these can also be inferred from trapping studies that show that most flies are captured in border rows (2).

Feeding

While feeding, adult Anastrepha flies exhibit the following behaviors:

dabbing (repeated lowering of proboscis to touch the surface of leaves while walking at increased rates of turning), sucking (absorbing water droplets or liquid oozing from a fruit), bubbling (formation of a drop of liquid of varying sizes at the tip of proboscis while fly sits motionless), and regur­gitating (deposition of a series of regurgitated drops on a leaf or fruit and reabsorption of those drops after varying intervals of time) (10). In studying dabbing (or grazing) behavior in another tephritid fly (Rhagoletis pomonella), Hendrichs and collaborators (81) determined that through grazing on leaf

surfaces adult flies could accrue small amounts of carbohydrates and some proteins. These essential nutrients were made available by the plant through

leaching and guttation processes. Regurgitation behavior has been reported in A. fraterculus, A. obliqua, A. ludens, A. serpentina, A. striata, and A. suspensa (6, 10,80, 175). In A. striata, individuals deposit a mean of 23.5 drops and reingest them within 12 min (10).

Oviposition

Oviposition behavior in Anastrepha is a dynamic process that follows a pattern closely resembling other frugivorous tephritids (149). Apparently both olfactory and visual stimuli lead gravid females to suitable oviposition sites. Factors influencing final choice are poorly understood. Females typically land on fruit, inspect it by walking on the surface (exhibiting

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ANASTREPHA BIONOMICS AND MANAGEMENT 165

repeated head butting bouts), bore, and oviposit. As discussed earlier, eggs are deposited singly or in clutches, with clutch size strongly affected by host size (24). In the few species studied so far (A. suspensa, A. fraterculus, A. ludens, A. serpentina, A. obliqua, A. striata), females deposit a water­soluble host-marking pheromone (HMP) after ovipositing (147, 148; F. Diaz, personal communication). Although they deter females prior to egg­laying, HMPs actually seem to stimulate continued host-marking behavior after egg laying (139). Work with A. ludens has suggested that host-marking is regulated by sensory adaptation or habituation to HMP in conjunction with dosage-dependent restoration of inhibition of the motor pattern (139).

Mating

Anastrepha males follow two distinct mating strategies: (a) resource defense polygyny exhibited by species such as A. bistrigata that are strictly mo­nophagous and patrol and defend clumps of fruit that are attractive to receptive females (121) and (b) lek polygyny exhibited by polyphagous species such as A. fraterculus, A. obliqua, A. ludens, and A. suspensa (7, 33, 80, 120, 170).

In lekking species, males establish territories preferably on the underside of leaves of host and nonhost trees (5, 107). Males deposit pheromone on leaf territories and defend them with a mixture of postures, sounds, and

- physical encounters (34, 55, 174). Within a lek (or outside of it) males typically emit a series of courtship and aggressive songs via rapid wing vibrations (190, 191) and release a sex pheromone through the mouth and anus (127, 130). Pheromone release is achieved by puffing expanded pleural abdominal membranes and an everted proctiger (129). The males make a precopulatory sound as they mount the back of a potential mate or if a female becomes restless during copulation (172). A typical characteristic of the Anastrepha lekking system is that females make discriminating mate decisions, preferring larger males (35). Larger males sing better-quality songs and presumably pass more sperm to females (172). In species such as A. striata, the process of female choice can take up to two hours. During this time, elaborate close-range courtship displays and interactions, including repeated mating attempts and labellum-to-Iabellum contacts (during which actual trophalaxis takes place), occur (10).

MANAGEMENT

Anastrepha control and management has, in my opinion, been conceptually and technically stagnant for the past 35 years. From a historic perspective, control strategies can be summarized as follows: placement of McPhail traps, application of poisoned bait sprays for monitoring and control, and

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heavy reliance on postharvest treatments with fumigants. The recent banning

of the fumigant ethylene dibromide (EDB) by the US Environmental Pro­tection Agency has finally prompted an upsurge in research aimed at identifying new postharvest treatments and monitoring tools (lures, traps) .and overall management schemes. Here, owing to space restrictions, I only refer to relevant work in each of the areas that are currently being applied

to manage Anastrepha populations.

Adult Detection Mechanisms

Without question the most widely used traps to monitor and in some cases control Anastrepha populations are glass and plastic versions of the McPhail trap, which is baited with a mixture of protein (occasionally molasses or fermented fruit juices are also used) and water (2, 18, 32, 103, 186). This trap, even though widely used, has several drawbacks: it is expensive, breaks easily, and is cumbersome to service (2); it works better in dry climates (50); it preferentially captures females (88); and, most importantly, it is very inefficient. For example, working in a mixed mango orchard with large numbers of A. ludens, A. obliqua, and A. serpentina, we (6) found that out of 665 flies that landed on the exterior of the trap, only 3 1 . 1 %

were caught. Furthermore, when fly populations are small, high densities of traps are needed to detect them (38).

Recent work on increasing the effectiveness of Anastrepha monitoring mechanisms has progressed along three lines: developing alternatives to standard hydrolyzed protein baits in McPhail traps, designing color-based traps, and attempting to identify Anastrepha sexual pheromones in order to develop a pheromone trap. Human urine has been found to be 10 times more effective than torula yeast in attracting A. striata and A. obliqua flies to McPhail traps in guava groves in Costa Rica (78). The odor of fermented yellow chapote (Sargentia greggii), the preferred native host of A. ludens, is 3.6 times more attractive than yeast hydrolysate to hungry, laboratory reared A. ludens (153). Nevertheless, chapote odor almost completely inhibited attraction of sexually active A. ludens females to male pheromone (152).

Colors such as yellow and orange, reflecting maximally within a narrow spectral region (Le. 500-590 nm), have proven effective at capturing several Anastrepha species (e.g. A. fraterculus, A. ludens, A. suspensa) when used in spherical, rectangular or cylindrical traps (31, 51, 54, 73, 111, 117). For example, orange panels (reflecting maximally at 590 nm) attracted numerous mature A. suspensa females (73). Orange balls, 20 cm in diameter, were also effective at attracting sexually mature A. suspensa females; sphere attractiveness was increased further by placing calling males close to the balls (171).

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Identification of Anastrepha sexual pheromones has been a slow process

that started in the 1970s and has been mostly restricted to two species: A.

suspensa and A. ludens (77, 127, 130). To date, several compounds have been isolated and their structures elucidated. Some of these compounds have been synthesized (22, 46, 122, 128, 154, 155, 187). Both females and males are attracted to calling males, and male-baited traps catch five to ten times more flies than food-baited traps in field trials using laboratory-reared,

virgin A. suspensa flies (141). Similar results have been reported for A. ludens (151).

Biological Control

Both classical biological control and repeated augmentative releases of mass-reared parasitoids have been used to suppress Anastrepha populations. Classical biological control has been attempted repeatedly in various coun­tries. Parasitoid species such as Diachasmimorpha longicaudata, Doryctobracon crawfordi, Ganaspis pelleranoi, Biosteres giffardi, B. vandenboschi, and Aceratoneuromyia indica have been imported and re­leased in the US, Mexico, Costa Rica, Brazil, Peru, and Argentina for the control of A. suspensa, A. iudens, and A. fraterculus (47, 89, 192). Of these, D. longicaudata and D. crawfordi have become firmly established from the US to Argentina. Unfortunately, the impact that these release programs had on native fly populations was never adequately documented .

Examples of augmentative releases of parasitoids are rare with Anastrepha. These include efforts in Florida where thousands of D. longicaudata have caused significant reductions in wildA. suspensa populations (19; J. Sivinski, personal communication). Other efforts in Mexico and Costa Rica have also apparently been successful in controlling A. obliqua and A. ludens but have never been published (D. Enkerlin & T.T.Y. Wong, unpublished data; H. Camacho, unpublished data). Such an approach has proven to be highly successful against Ceratitis capitata in Hawaii (195) under conditions similar to those encountered in Latin America. Higher success rates in the future are, in my view, contingent on prerelease studies, a more critical selection of natural enemies to be released, and a better understanding of parasitoid behavior and ecology.

Sterile Insect Technique

Even though release of sterile insects alone has proven effective at eradicating or substantially suppressing populations of A. suspensa, A. ludens, and A.

fraterculus in the US (86, 87), Mexico (48, 167), and Peru (69), the current trend is to explore the possibility of joint sterile fly and parasitoid releases. Computer models suggest that the use of sterile pest releases and inundative release of parasitoids are more efficient together than either method alone

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(20, 94), because sterile releases are most effective at low pest density, whereas the inundative releases of parasitoids are more efficient at high pest densities (20, 94). Also, parasitoids can be used to reduce fly popu­lations prior to sterile fly releases. To date, no field tests reporting results of these new strategies applied to Anastrepha have been published, but work is underway (J. Sivinski, personal communication).

Chemical Control

Anastrepha flies are susceptible to almost all insecticides (2, 157). Quick­acting poisons are preferred to prevent adult females from ovipositing after exposure to the poison. In the early parts of the century, naturally occurring and inorganic insecticides such as nicotine, rotenone, arsenic, and copper salts ( 13), as well as soil toxicants ( 168), were used. Since then, the most widely used mechanism has been the use of bait sprays applied from the ground or air ( 102, 166). Because bait sprays have been shown to cause unacceptably high environmental damage, surges of secondary pests, and reductions in parasitization rates (56, 102, 179), research is underway to find more acceptable alternatives. Current studies are searching for toxicants that are more fly specific such as cyro_mazine (1 1 0) and avermectin Bl ( 1)

and products that are less harmful to the environment such as borates (59).

Legal Control

Anastrepha is considered a major threat to the fruitgrowing industry in almost every country in the world. Thus a series of quarantine regulations limit the movement of fruit from infested areas to areas considered to be pest-free. Legal control encompasses quarantines, issuing of certificates and limited permits for use when fruit are moved from infested areas to places considered fly-free, orchard certification, authorizing and strictly enforcing fruit disinfestation procedures, and establishment of fly-free zones. Most advances have been made in the two last areas (i.e. fruit disinfestation procedures and establishment of fly-free zones) in the past 10 years. Several alternative disinfestation methods against A. suspensa, A. /udens, A. serpentina, and A. obliqua larvae have recently been developed: e.g. hot-water or hot-air treatments in mangoes, guavas, and carambolas (72, 125, 158, . 161, 163); use of radiation (37); and cold-storage (71). Also, a flexible acoustical device was developed to detect feeding sounds of A. suspensa larvae in such fruit as mango, grapefruit, and guava (164). Generally, disinfestation treatments must effect a mortality of 99.9968% (probit 9) of eggs and larvae in the fruit (97). Landolt et al (97) have proposed an alternative to this unrealistic value based on calculations of the probability of a mating pair surviving a shipment.

Another recent alternative is the establishment of fly-free zones. Such

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ANASTREPHA BIONOMICS AND MANAGEMENT 169

zones are currently operating in the US, Mexico, Ecuador, and Brazil. Certification procedures are site specific and strictly enforced following a series of rules negotiated by both exporting and importing countries [150; see also USDA-PPQ-APHIS Fly-free Area Regulations (unpublished infor­mation), for a working example].

Alternative Methods

A series of new approaches have been proposed recently. For instance, plant-growth regulators can cause the plant to produce more of a given resistance agent or sustain the innate resistance properties of the plant. One such substance is gibberellic acid, which has been shown to reduce grapefruit susceptibility to attack by A. suspensa (74). Other approaches currently under research are border trapping, trap cropping, and habitat manipulation (2, 1 1).

DIRECTIONS OF FUTURE RESEARCH

In my view, three areas of research warrant our immediate attention: taxonomy, basic biology'and ecology, and management. A thorough taxo­nomic revision of the genus (including identification keys to immature stages and males) is long overdue. More emphasis needs to be placed on studying Anastrepha spp. under natural conditions and, if at all possible, in unper­turbed environments. This effort might lead us away from species of economic importance but will undoubtedly add new depth to our under­standing of basic patterns in Anastrepha biology and ecology. Our under­standing of factors that regulate population numbers in the field and of how individuals can track down unpredictable and ephemeral resources is still only marginal. Comparative studies between monophagous and polyphagous, or primitive and derived, species in areas such as demography, behavior, life-history traits, and chemical ecology should be especially rewarding. In the area of Anastrepha management we need to develop better traps and lures, identify more efficient biological-control agents, and most importantly, develop economic thresholds tailored to meet both the stringent quality requirements of international fresh fruit markets and the less rigorous local markets. Finally, much more attention should be paid to the development of truly integrated (Le. integration of all available control methods and consideration of other pests and diseases that attack the same crop) and regional management schemes that are based on ecologically sound principles (e.g. trap cropping, habitat manipulation) and that are within the cultural and economic reach of the recipient (11). As part of a regional management strategy, countries could be divided up according to biogeographic charac­teristics into large eradication zones, localized fly-free zones, integrated

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management zones, and zones where any control action is technically and economically unfeasible (see 11).

ACKNOWLEDGMENTS

I am most indebted to RJ Prokopy and RT Carde for their support. I sincerely thank all colleagues who provided reprints and copies of unpub­lished manuscripts, but because of space restrictions, I cannot list their names. I thank P Greany, J Sivinski, A Norrbom, V Hernandez, P Liedo, and G Dieringer for extremely useful comments on a previous version of the manuscript. I acknowledge support from Programa MOSCAMED (DGSV-SARH), CONACyT, SEP, and IFS (projects 051/93, Dl1 1-903537, DGICSA-902467, and Cl174 1-1, respectively).

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