Liriomyza trifolii (American serpentine leafminer)
- Summary of Invasiveness
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- Risk of Introduction
- Habitat List
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Biology and Ecology
- Natural enemies
- Notes on Natural Enemies
- Pathway Vectors
- Plant Trade
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- Links to Websites
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Liriomyza trifolii Burgess in Comstock, 1880
Preferred Common Name
- American serpentine leafminer
Other Scientific Names
- Agromyza phaseolunata Frost, 1943
- Liriomyza alliivora Frick, 1955
- Liriomyza alliovora Frick, 1955
- Liriomyza phaseolunata (Frost, 1943)
- Oscinis trifolii Burgess in Comstock, 1880
International Common Names
- English: chrysanthemum leaf miner; serpentine leaf miner
- Spanish: minador pequeño del frijol
- French: mineuse du gerbera
Local Common Names
- Germany: Floridaminierfliege
- Russian Federation: American clover miner
- LIRITR (Liriomyza trifolii)
Summary of InvasivenessTop of page
L. trifolii is a leaf-mining insect, commonly known as the serpentine leafminer. It is highly polyphagous and has been recorded from 25 families. As a major pest of ornamental and vegetable crops, including beans (phaseolus), Capsicum, carnations, celery, chrysanthemums (Dendranthenum, the commercial 'Mum'), clover, Cucumis, Gerbera, Gypsophila, lettuces, lucerne, potatoes, Senecio hybridus and tomatoes it has had important biological and economic impacts across a number of countries, including North America, Africa and Europe.<_u13a_p />
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Metazoa
- Phylum: Arthropoda
- Subphylum: Uniramia
- Class: Insecta
- Order: Diptera
- Family: Agromyzidae
- Genus: Liriomyza
- Species: Liriomyza trifolii
Notes on Taxonomy and NomenclatureTop of page
Liriomyza trifolii is one of the truly polyphagous agromyzids and has been recorded in 25 families (Spencer, 1990). It was first described as Oscinis trifolii (Burgess in Comstock, 1880) in the family Chloropidae from flies attacking the leaves of Trifolium repens (white clover) in Indiana, USA. Later, it was transferred to the family Agromyzidae in the genus Agromyza by Coquillet (1898), then to Liriomyza by de Meijere (1925). Spencer, 1973 synonymized Liriomyza alliovora Frick, 1955, breeding in Allium (onions) in Iowa, USA, and in Spencer, 1986, Agromyza phaseolunata Frost (1943, as Liriomyza) attacking Phaseolus lunatus (lima beans) in New Jersey, USA with L. trifolii.
DescriptionTop of page
Descriptions of L. trifolii refer to fresh materials. Dry specimens may be distorted due to the manner in which they have been preserved. Also, the age of the specimen, when killed, will have some effect on its preservation characteristics.
For accurate identification, examination of the leaf mine and all stages of development are crucial.
L. trifolii eggs are 0.2-0.3 mm x 0.1-0.15 mm, off white and slightly translucent.
This is a legless maggot with no separate head capsule, transparent when newly hatched but colouring up to a yellow-orange in later instars and is up to 3 mm long. L. trifolii larvae and puparia have a pair of posterior spiracles terminating in three cone-like appendages. Spencer (1973) describes distinguishing features of the larvae. Petitt (1990) describes a method of identifying the different instars of the larvae of L. sativae, which can be adapted for use with the other Liriomyza species, including L. trifolii.
This is oval and slightly flattened ventrally, 1.3-2.3 x 0.5-0.75 mm with variable colour, pale yellow-orange, darkening to golden-brown. The puparium has posterior spiracles on a pronounced conical projection, each with three distinct bulbs, two of which are elongate. Pupariation occurs outside the leaf, in the soil beneath the plant.
Menken and Ulenberg (1986) describe a method of distinguishing L. trifolii from L. bryoniae, L. huidobrensis, and L. sativae using allozyme variation patterns as revealed by gel electrophoresis.
L. trifolii is very small: 1-1.3 mm body length, up to 1.7 mm in female with wings 1.3-1.7 mm. The mesonotum is grey-black with a yellow blotch at the hind-corners. The scutellum is bright yellow; the face, frons and third antennal segment are bright yellow. Male and female L. trifolii are generally similar in appearance.
L. trifolii are not very active fliers, and in crops showing active mining, the flies may be seen walking rapidly over the leaves with only short jerky flights to adjacent leaves.
The frons, which projects very slightly above the eye, is just less than 1.5 times the width of the eye (viewed from above). There are two equal ors and two ori (the lower one weaker). Orbital setulae are sparse and reclinate. The jowls are deep (almost 0.33 times the height of the eye at the rear); the cheeks form a distinct ring below the eye. The third antennal segment is small, round and noticeably pubescent, but not excessively so (vte and vti are both on a yellow ground).
Acrostical bristles occur irregularly in 3-4 rows at the front, reducing to two rows behind. There is a conspicuous yellow patch at each hind-corner. The pleura are yellow; the meso- and sterno-pleura have variable black markings.
Length 1.3 -1.7 mm, discal cell small. The last section is M<(sub)3+4> from 3-4 times the length of the penultimate one.
The shape of the distiphallus is fairly distinctive but could be mis-identified for L. sativae. Identification using the male genitalia should only be undertaken by specialists.
The head (including the antenna and face) is bright yellow. The hind margin of the eye is largely yellow, vte and vti always on yellow ground.
The mesopleura is predominantly yellow, with a variable dark area, from a slim grey bar along the base to extensive darkening reaching higher up the front margin than the back margin. The sternopleura is largely filled by a black triangle, but always with bright yellow above.
The femora and coxa are bright yellow, with the tibia and tarsi darker; brownish-yellow on the fore-legs, brownish-black on the hind legs. The abdomen is largely black but the tergites are variably yellow, particularly at the sides. The squamae are yellowish, with a dark margin and fringe.
Although individual specimens may vary considerably in colour, the basic pattern is consistent.
DistributionTop of page
L. trifolii has not yet been reported from many countries where it is actually present. It is generally recognized that all the countries bordering the Mediterranean have L. trifolii in varying degrees and that it occurs in all mainland states of the USA. L. trifolii has been recorded from the Juan Fernandez Islands (an offshore territory of Chile; Martinez and Etienne, 2002; EPPO, 2009). See also CABI/EPPO (1998, No. 96). L. trifolii is apparently unable to overwinter in the open in the north European EPPO countries. However, the current regulations to prevent entry and spread in non-Mediterranean areas were found to be only partially effective, as interceptions are still being reported (EFSA, 2012).
The record for Argentina has been changed to 'Absent, unreliable record' as Martinez and Etienne (2002) and EPPO (2006) are based on Burgess (in Comstock, 1880 (1879)) and there have been no other reports of the pest in Argentina. L. trifolii is a quarantine pest for Argentina (SENASA, personal communication, 2008).
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.Last updated: 30 Jun 2021
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|South Africa||Present, Few occurrences|
|Cambodia||Absent, Unconfirmed presence record(s)|
|-Jammu and Kashmir||Present|
|Laos||Absent, Unconfirmed presence record(s)|
|Malaysia||Absent, Unconfirmed presence record(s)|
|Taiwan||Present, Few occurrences|
|Thailand||Absent, Unconfirmed presence record(s)|
|United Arab Emirates||Present|
|Austria||Present, Localized||First reported: 198*|
|Bosnia and Herzegovina||Present|
|Denmark||Absent, Intercepted only|
|Estonia||Absent, Confirmed absent by survey|
|Finland||Present, Few occurrences|
|Germany||Absent, Formerly present||First reported: 197*|
|Lithuania||Absent, Confirmed absent by survey|
|Montenegro||Absent, Formerly present|
|Poland||Absent, Formerly present||1980|
|Romania||Present, Few occurrences|
|-Central Russia||Present, Localized|
|-Southern Russia||Present, Few occurrences|
|Serbia and Montenegro||Present, Localized|
|Slovakia||Absent, Invalid presence record(s)|
|-Canary Islands||Present, Localized|
|Switzerland||Present, Few occurrences||First reported: 198*|
|United Kingdom||Absent, Eradicated||1977|
|British Virgin Islands||Present|
|Netherlands Antilles||Present, Localized|
|Saint Kitts and Nevis||Present, Localized|
|Trinidad and Tobago||Present|
|U.S. Virgin Islands||Present|
|United States||Present, Localized|
|-District of Columbia||Present|
|Australia||Absent, Intercepted only|
|-New South Wales||Absent, Intercepted only|
|-Victoria||Absent, Intercepted only|
|Federated States of Micronesia||Present|
|Northern Mariana Islands||Present|
|Argentina||Absent, Unconfirmed presence record(s)||Original citation: SENASA, personal communication, 2008|
|-Rio Grande do Norte||Present|
Risk of IntroductionTop of page
L. trifolii is listed as an A2 quarantine pest by EPPO (OEPP/EPPO, 1984). It is one of the most important recent introductions to the EPPO region.
It is a major pest of a wide variety of ornamental or vegetable crops grown under glass (Lactuca, Dendranthema, Gypsophila, Dahlia) or as protected crops in the EPPO region. It could also cause damage to these crops grown in the open in the warmer parts of the EPPO region. It is widely distributed in the region and the success of eradication programmes which have been conducted cannot be confirmed.
HabitatTop of page
L. trifolii'sdevelopment is strictly connected with temperature. Consequently, at a uniform temperature of 28°C one generation cycle can be accomplished in 14-15 days, but at lower temperatures the time taken is progressively longer. At 16°C puparial diapause begins and winter generations of puparia will remain in the soil until warmer conditions occur again. The adult can survive temperatures down to about 12°C but does not appear to feed or lay eggs.
In heated glasshouses where suitable hosts may be grown throughout the year, the breeding and development of L. trifolii will be virtually continuous. In cool glasshouses generation rates will be different throughout the seasons, with fairly rapid development during the summer and puparia remaining undeveloped in the soil during the coldest periods.
In the moderate and variable temperatures of open-field cultivation there will be only a few (perhaps three) generations produced throughout the growing season because of the longer time required to complete each cycle (Süss et al., 1984).
Habitat ListTop of page
Hosts/Species AffectedTop of page
The host range of L. trifolii includes over 400 species of plants in 28 families including both ornamental crops (Bogran, 2006) and vegetables (Cheri, 2012). The main host families and species include: Apiaceae (A. graveolens); Asteraceae (Aster spp., Chrysanthemum spp., Gerbera spp., Dahlia spp., Ixeris stolonifera, Lactuca sativa, Lactuca spp., Zinnia spp.); Brassicaceae (Brassica spp.); Caryophyllaceae (Gypsophila spp.); Chenopodiaceae (Spinacia oleracea, Beta vulgaris); Cucurbitaceae (Cucumis spp., Cucurbita spp.); Fabaceae (Glycine max, Medicago sativa, Phaseolus vulgaris, Pisum sativum, Pisum spp., Trifolium spp., Vicia faba); Liliaceae (A. cepa, Allium sativum) and Solanaceae (Capsicum annuum, Capsicum frutescens, Petunia spp., Solanum lycopersicum, Solanum spp.) (EFSA, 2012).
It is now considered to be the most important pest of cowpea (Vigna uniguilata), towel gourd (Luffa cylindrica), cucumber (Cucumis sativus) and many other vegetable crops in southern China (Gao, 2014). In Europe, L. trifolii is a major pest of lettuce, beans, cucumber and celery, Capsicum sp., carnations, clover, Gerbera sp., Gypsophila sp., lucerne, Senecio hybridus, potatoes and tomatoes (EFSA, 2012). It is now a major pest of the Compositae worldwide, particularly chrysanthemums (including Dendranthenum, the commercial 'Mum') in North America, Colombia, and elsewhere. It also causes severe damage to different open field crops, such as chili peppers in Mexico.
Host Plants and Other Plants AffectedTop of page
Growth StagesTop of page
SymptomsTop of page
L. trifolii feeding punctures appear as white speckles between 0.13 and 0.15 mm in diameter. Oviposition punctures are usually smaller (0.05 mm) and are more uniformly round.
L. trifolii leaf mines can vary in form with the host plant, but when adequate leaf area is available they are usually long, linear, narrow and not greatly widening towards the end. They are usually greenish white.
In very small leaves the limited area for feeding results in the formation of a secondary blotch at the end of the mine, before pupariation. In Kenya, Spencer (1985) notes the growth of many L. trifolii from mines which began with a conspicuous spiral. This is not a characteristic associated with L. trifolii on other continents.
The frass is distinctive in being deposited in black strips alternately at either side of the mine (like L. sativae), but becomes more granular towards the end of the mine (unlike L. sativae) (Spencer, 1973).
Fungal destruction of the leaf may also occur as a result of infection introduced by L. trifolii from other sources during breeding activity. Wilt may occur, especially in seedlings.
List of Symptoms/SignsTop of page
|Leaves / abnormal colours|
|Leaves / abnormal forms|
|Leaves / abnormal leaf fall|
|Leaves / external feeding|
|Leaves / internal feeding|
|Leaves / necrotic areas|
|Leaves / wilting|
Biology and EcologyTop of page
L. trifolii eggs are inserted just below the leaf surface. Eggs hatch in 2-5 days according to temperature. Harris and Tate (1933) give 4-7 days at 24°C. Many eggs may be laid on a single leaf.
The duration of larval development also depends on temperature and probably host plant. Several generations can occur during the year, breeding only being restricted by the temperature and the availability of fresh plant growth in suitable hosts (Spencer, 1973).
L. trifolii pupariation occurs outside the leaf, in the soil beneath the plant. Puparial development will vary according to season and temperature. Adult emergence occurs 7-14 days after pupariation at temperatures between 20 and 30°C (Leibee, 1982). Wolfenbarger (1947) gives 24-28 days for the complete cycle, in Florida during December-January (winter period).
Peak emergence of adult L. trifolii occurs before midday (McGregor, 1914). Males usually emerge before females. Mating takes place from 24 hours after emergence and a single mating is sufficient to fertilize all a female's eggs.
Female L. trifolii flies puncture the leaves of the host plants causing wounds which serve as sites for feeding or oviposition. Feeding punctures cause the destruction of a large number of cells and are clearly visible to the naked eye. About 15% of oviposition punctures made by L. trifolii contain viable eggs (Parrella et al., 1981). Male L. trifolii are unable to puncture the leaves but have been observed feeding at punctures made by females. Both male and female L. trifolii feed on dilute honey (in the laboratory) and take nectar from flowers (OEPP/EPPO, 1990).
In the southern USA, the L. trifolii life cycle is probably continuous throughout the year. There is a noticeable first generation which reaches a peak in April (Spencer, 1973). In southern Florida, L. trifolii has two or three generations followed by a number of incomplete, overlapping generations (Spencer, 1973).
On celery L. trifolii completes its life cycle (oviposition to adult emergence) in 12 days at 35°C, 26 days at 20°C, and 54 days at 15°C (Leibee, 1982). On chrysanthemums the life-cycle is completed in 24 days at 20°C but on Vigna sinensis and Phaseolus lunatus it takes only 20 days at this temperature (Poe, 1981).
Adults of L. trifolii live between 15 and 30 days. On average, females live longer than males.
Both male and female L. trifolii may act as vectors for disease by transference during feeding or egg laying, but are not inherent carriers of disease.
Natural enemiesTop of page
|Natural enemy||Type||Life stages||Specificity||References||Biological control in||Biological control on|
|Bacillus thuringiensis kurstaki||Pathogen|
|Chrysocharis clarkae||Parasite||Hawaii; USA||chrysanthemums|
|Chrysocharis oscinidis||Parasite||Arthropods|Larvae||USA; Hawaii; Senegal||chrysanthemums; okras; potatoes; Solanum aethiopicum; vegetables ì ì|
|Chrysonotomyia punctiventris||Parasite||Arthropods|Larvae||Hawaii; Senegal; USA; Hawaii||chrysanthemums; okras; potatoes; Solanum aethiopicum; watermelons|
|Closterocerus utahensis||Parasite||USA; Hawaii||watermelons|
|Cothonaspis pacifica||Parasite||Guam; USA; Hawaii||Phaseolus vulgaris; Vigna unguiculata; watermelons|
|Diaulinopsis callichroma||Parasite||Hawaii; Senegal; Trinidad and Tobago||aubergines; chrysanthemums; okras; potatoes; Solanum aethiopicum; tomatoes|
|Diglyphus begini||Parasite||Arthropods|Larvae||California; Colombia; Guam; Hawaii; USA; Hawaii||beans; chrysanthemums; watermelons|
|Diglyphus intermedius||Parasite||Arthropods|Larvae||Hawaii; Senegal; USA; USA; California||chrysanthemums; Gerbera; okras; potatoes; Solanum aethiopicum|
|Diglyphus isaea||Parasite||Arthropods|Larvae||Hawaii; Netherlands; Poland||chrysanthemums; tomatoes|
|Diglyphus pulchripes||Parasite||Hawaii; USA||chrysanthemums|
|Eucoilidea guamensis||Parasite||Guam||Phaseolus vulgaris; Vigna unguiculata|
|Eucoilidea micromorpha||Parasite||Guam||Phaseolus vulgaris; Vigna unguiculata|
|Ganaspidium hunteri||Parasite||Arthropods|Larvae||Hawaii; USA; Hawaii||chrysanthemums; watermelons|
|Ganaspidium utilis||Parasite||Arthropods|Larvae||Guam; Tonga||beans; vegetables|
|Halticoptera||Parasite||Aamer and Hegazi (2014)|
|Halticoptera circulus||Parasite||Arthropods|Larvae||Hawaii; Senegal; Trinidad and Tobago; USA; Hawaii||chrysanthemums; okras; potatoes; Solanum aethiopicum; watermelons|
|Neochrysocharis formosa||Parasite||Arthropods|Larvae||Guam; Hawaii|
|Opius dimidiatus||Parasite||Arthropods|Larvae||Hawaii; Netherlands; Senegal; USA||chrysanthemums; okras; potatoes; Solanum aethiopicum; tomatoes|
|Opius dissitus||Parasite||Arthropods|Larvae||Hawaii; Senegal; USA; Hawaii||chrysanthemums; okras; potatoes; Solanum aethiopicum; watermelons|
|Pseudopezomachus masii||Parasite||Aamer and Hegazi (2014)|
|Zagrammosoma||Parasite||Aamer and Hegazi (2014)|
Notes on Natural EnemiesTop of page
Numerous parasitic wasps (Hymenoptera) occurring naturally, may be used for control of L. trifolii. These wasps are difficult to isolate or identify and local agricultural advisory services should be consulted about which species are available and natural in that locality, and their artificial introduction.
There has been considerable work on natural enemies in relation to biological control introduction programmes. Waterhouse and Norris (1987) give a detailed list of the natural enemies of Liriomyza spp. and a summary of the results of the biological control introductions against L. trifolii.
Foliar applications of the entomophagous nematode, Steinernema carpocapsae, significantly reduced adult development of L. trifolii (Harris et al., 1990).
Pathway VectorsTop of page
Plant TradeTop of page
|Plant parts liable to carry the pest in trade/transport||Pest stages||Borne internally||Borne externally||Visibility of pest or symptoms|
|Leaves||eggs; larvae||Yes||Pest or symptoms usually visible to the naked eye|
|Seedlings/Micropropagated plants||eggs; larvae||Yes||Pest or symptoms usually visible to the naked eye|
|Plant parts not known to carry the pest in trade/transport|
|Growing medium accompanying plants|
|Stems (above ground)/Shoots/Trunks/Branches|
|True seeds (inc. grain)|
ImpactTop of page
L. trifolii is an economically important key pest of both ornamental crops (Bogran, 2006) and vegetables (Cheri, 2012).
In Kenya, chrysanthemums were grown commercially before 1976, but L. trifolii was thought to have been introduced in contaminated cuttings from Florida (USA) in 1976, at a large propagating nursery at Masongaleni. By 1979 the nursery was closed, but the establishment of the pest in local wild hosts, and the dissemination of cuttings from the nursery to other parts of the country as well as abroad, has added L. trifolii to the other pests of East Africa. It has caused considerable crop losses and loss of overseas markets due to quarantine requirements (IPPC Secretariat, 2005).
Vegetable losses in the USA are also considerable. For example, losses for celery were estimated at US$ 9 million in 1980 (Spencer, 1982). It was noted, however, that damage to celery during the first 2 months of the 3-month growing season was insignificant and largely cosmetic, whereas considerable yield loss resulted from pest presence during the final month (Foster et al., 1988). 1.5 million larval mines per hectare were recorded from onions in Iowa (Harris et al., 1933).
Damage is caused by L. trifolii larvae mining into leaves and petiole. The photosynthetic ability of the plants is often greatly reduced as the chlorophyll-containing cells are destroyed. Severely infested leaves may fall, exposing plant stems to wind action, and flower buds and developing fruit to scald (Musgrave et al., 1975). The presence of unsightly larval mines and adult punctures caused by L. trifolii in the leaf palisade of ornamental plants, such as chrysanthemums, can further reduce plant value (Smith et al., 1962; Musgrave et al., 1975). In young plants and seedlings, L. trifolii mining may cause considerable delay in plant development, even leading to plant loss. The level of damage depends on many factors, including climate suitability, host resistance, crop distribution, growing conditions, control methods in place and the degree of infestation (EFSA, 2012).
L. trifolii is also known to be a vector of plant viruses (Zitter et al., 1980).
Detection and InspectionTop of page
L. trifolii are small black and yellow flies which may be detected flying closely around host plants or moving erratically and rapidly upon the leaf surfaces. Inspection of the leaf surface will reveal punctures of the epidermis and the obvious greenish-white mines with linear grains of frass along their length. For accurate identification, examination of the leaf mine and all stages of development are crucial.
L. trifolii larvae will be found feeding at the end of the mine, or the mine will end with a small convex slit in the epidermis where the larva has left the mine to pupariate on the ground. Sometimes the puparium may be found adhering to the leaf surface, although in most cases the fully-fed larva will have found its way to the ground beneath the plant to pupariate. This is especially true in hot, dry conditions where the larva/puparia would quickly desiccate if exposed on the leaf surface. Empty puparial cases are split at the anterior end, but the head capsule is not usually separated from the rest of the case.
Mined leaves should be collected into polythene bags and transferred to a press as soon as possible. Leaves containing larvae intended for breeding should be collected into individual polythene bags, which on return to the laboratory should be slightly over-pressurized by blowing into them before sealing the end. Blowing up the bag by mouth and sealing it adds valuable carbon dioxide to the moist air mix. Constant attention is required to ensure that puparia are transferred to individual tubes until the fly emerges. If the plant material begins rotting, good material with feeding larvae must be removed to more sanitary conditions.
When puparia are observed they can be very carefully removed to tubes containing a layer of fine sand, or a small strip of blotting paper or filter paper. This should be kept damp (never wet) until the adult emerges.
On emergence, the fly should be kept for at least 24 hours to harden up. Do not allow condensation to come into contact with the fly, or it will stick to the water film and be damaged.
Field collection of the adult L. trifolii is done by netting. The use of sticky traps, especially yellow ones, placed near host plants is a very effective method of collection and estimation of infestation.
If the puparial stage is collected from the soil, care must be taken not to damage the puparial skin or death will almost certainly follow. The pupae should be stored in glass tubes on a layer of clean sand or, better still, thick filter paper. The tube must have high humidity, but be free of condensation.
When the fly emerges, it must be allowed to harden for 24 hours before killing for identification purposes. Ensure that the tube has no condensation present.
Newly emerged adult L. trifolii are generally softer than specimens aged for several days and may crinkle as drying proceeds, especially the head. The ptilinal sac may still protrude from the suture between the frons and face obliterating some important characteristics. Adults should be dried slowly in the dark in a sealed receptacle over blotting paper. If preserving wet is preferred, the live specimen should be dropped into 20-40% alcohol, and transferred to 70-90% alcohol after 2 days.
Similarities to Other Species/ConditionsTop of page
Liriomyza species, in general, may be recognized by their black (sometimes brilliantly black) and yellow colouring. Particularly, the scutellum is usually yellow and distinctive.
Several pests in this genus are similar and may be mistaken for each other on quick examination. These are L. sativae (shining black mesonotum without yellow at the hindquarters, vte always and vti usually on black ground, origins probably in South America); L. huidobrensis (which has a larger discal cell, origins in South America); L. trifolii (origins probably Caribbean/Florida); L. brassicae (origins probably South America/Caribbean); L. bryoniae (origins in Europe); L. congesta (Origins in Europe/western Asia); L. strigata (origins in Europe).
The spread of Liriomyza species through international commerce and the similarities between the seven species means that identification of individual infestations must be confirmed by specialists (Spencer, 1973).
Prevention and ControlTop of page
Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.
The use of glue traps can be effective for assessing the presence of adult L. trifolii, gauging the best time to apply control measures on a population, and as a direct method of pest suppression (Valenzuela, 2010). Yellow sticky traps (YSTs) attracted significantly more adult L. trifolii than blue, purple or white traps; the average percentage of damaged leaves and damage severity (number of mines per leaf) were significantly lower in fields with YSTs after 50 days (Arida et al., 2013).
L. trifolii has developed resistance to most commonly used insecticides that were recommended for its control before 1990 (Parella et al., 1984; Nuessly and Webb, 2013), including carbamates, organophosphates, pyrethroids, avermectins, spinosyns and moulting disruptors, such as cyromazina (Hernandez, 2009). However, its susceptibility to insecticides varies widely between agricultural regions and populations. In Florida, USA, the lifetime of an insecticide’s effectiveness is often only two to four years, and is then usually followed by a strong resistance in treated populations (Reitz et al., 2013; Capinera, 2014).
The insecticides (active ingredients) abamectin, acephate, acephate + fenpropathrin, acetamiprid, bifenazate + abamectin, bifenthrin, carbaryl, chlorpyrifos, clothianidin, cyantraniliprole, cyromazine, deltamethrin, diazinon, diflubenzuron, dimethoate, dinotefuran, emamectin benzoate, fenpropathrin, fenoxycarb, gamma-cyhalothrin, imidacloprid, indoxacarb, lambda-cyhalothrin, lambda-cyhalothrin + chlorantraniliprole, malathion, novaluron, naled, permethrin, phosmet, rynaxypyr (chlorantraniliprole), spinetoram, spinosad, thiamethoxam, thiamethoxam + chlorantraniliprole, and the natural insecticides azadirachtin, extract of Chenopodium ambrosioides, Isaria fumosorosea Apopka strain 97, mineral oils, potassium salts of fatty acids and pyrethrins have been cited for the control or suppression of immature or adult L. trifolii in agricultural and ornamental crops (Price and Nagle, 2012; Webb et al., 2012; Webb and Stansly, 2012; Misra, 2013; Nuessly and Webb, 2013; Webb, 2013).
For the control of L. trifolii, effective insecticides with different modes of action (and with different site of action) should be rotated during the growing season (IRAC, 2014).
Natural enemies periodically suppress leaf-miner populations (Spencer, 1972). Parasitoids, and to a lesser extent to nematodes, bacteria and fungi, are used for biological control of leafminers (Cikman y Comelkcloglu, 2006; Sher et al. 2000; Abd El-Salam et al., 2012; Capinera, 2014). Although several predatory species have been found feeding on Liriomyza, predators are not considered to be important as biological control agents (Liu et al., 2009; Capinera 2011, 2014). There are several successful cases of classical biological control with parasitoids to different species of leaf miners, both in open fields and greenhouses (Abd-Rabou, 2006; Salvo y Valladares, 2007; Liu et al., 2011;).
There has been considerable work on natural enemies in relation to biological control introduction programmes. Waterhouse and Norris (1987) gave a detailed list of the natural enemies of Liriomyza spp. and a summary of the results of the biological control introductions against L. trifolii. In Hawaii, several parasitoids were already present as immigrant species, presumably accidentally introduced with their hosts. More of these parasitoids were introduced from the USA, to broaden the genetic base, as well as Chrysonotomyia punctiventris and Ganaspidium hunteri, and have proved a substantial control, at least on water melons when natural enemies are not eliminated by pesticide sprays (Johnson, 1987). Subsequently, introductions of species established in Hawaii were made in Pohnpei (Mariana Islands) and G. utilis and C. oscinidis became established and are credited with achieving substantial control (Suta and Esguerra, 1993). These two parasitoids have also been successfully established in Tonga where control is reported as complete (Johnson, 1993). Earlier unsuccessful introductions were made in the Caribbean islands (Cock, 1985) and a biological control programme has been carried out in Senegal: the results of this require re-assessment but it is unlikely that any beneficial results were obtained (Neuenschwander et al., 1987).
Foliar applications of the entomophagous nematode Steinernema carpocapsae significantly reduced adult development of L. trifolii (Harris et al., 1990).
Extensive global research has reported more than 150 species of parasitoids associated with Liriomyza sp. (Liu et al. 2011). For L. trifolii,Hernandez et al. (2010) listed 20 genera of parasitoids in various chili crops during autumn 2007 and spring 2008 in Weslaco, Texas, USA: Neochrysocharis formosa, Closterocerus cinctippenis Ashmead, Diglyphus isaea, Cirrospilus variegatus Masi, Asecodes spp., Pnigalio spp., Zogrammosoma spp., Chrysocharis spp. (Eulophidae); Opius dissitus Muesebeck, O. dimidiatus (Ashmead), O. nr. brownsvillensis Fischer, O. thoracosema Fischer, O. bruneipes Gahan, O. spp. (Braconidae); Ganaspidium pusillae Weld, G. nigrimanus (=utilis) (Kieffer), Disorygma pacifica (Yoshimoto), Agrostocynips robusta (Ashmead) (Figitidae) and Halticoptera nr. circulus Walker (Pteromalidae). N. formosa was the most common, comprising 60% of the natural enemies.
In Tamaulipas, Mexico, Arcos-Cavazos et al. (2011) found six larval parasitoid hymenopterid natural enemies: Opius sp., Chysocharis sp., Diglyphus sp. (Eulophidae), Gronotoma sp. (Hymenoptera: Figitidae), and two unidentified species. Of these, Chrysocharis was the primary regulator of L. trifolii populations, with an average of 79.5% larval parasitism of L. trifolii and in some samples of 100%. Fadl and El-Khawas (2009) found five species of hymenopterid parasitoids of L. trifolii on tomato in Qalyubia, Egypt, during two growing crop seasons: Cirrospilus sp., Diglyphus crassinervis, D. isaea, Chrysocharis sp. and Neochrysocharis.Neochrysocharis had the highest recorded total numbers.
Currently, mass rearing of leaf miner parasitoids for augmentative biological control includes the simultaneous use of three trophic levels: host plant, phytophagous insects and parasitoids, which may be difficult and costly. Therefore, the idea should be carefully considered (Cortez-Mondaca and Valenzuela-Escoboza, 2013).
The impact of insecticides on parasitoids of leaf miners is complex and further studies are needed to determine which insecticides are least damaging to natural enemies of L. trifolii (Hernandez 2009). Field studies suggested that cyromazine has the least impact on parasitoid populations, followed by abamectin and spinosyns, which in turn were not as detrimental as carbamates, organophosphate or pyrethroids (Reitz et al., 2013). Nuessly and Webb (2013) reported that the use of selective insecticides, such as spinosad and emamectin benzoate, for armyworm and cabbage looper control also provided some control of L. trifolii populations, as well as being gentle to most beneficial insects. Novaluron had the least impact on adult parasitoids in laboratory bioassays compared to other treatments (abamectin, spinetoram, lambda-cyhalothrin) (Hernandez, 2009). The insecticide lambda-cyhalothrin showed negative effects only for the parasitoid Ganaspidium nigrimanus (in topical application assays), but residual tests had negative effects on G. nigrimanus and on Neochrysocharis formosa. Abamectin showed no ill effects on N. formosa or G. nigrimanus in topical bioassays. In contrast, spinetoram showed negative effects on N. formosa and G. nigrimanus in all bioassays in the laboratory.
It is possible to increase the action of leafminer natural enemies through habitat management (Musundire et al., 2012). Weed patches near crops may be important as possible reservoirs of parasitoids (Altieri and Nichols, 2009). For this reason, there have been suggestions of increasing weed diversity or improving the availability of pollen and nectar for natural enemies in agroecosystems affected by L. trifolii (Altieri et al., 2005; Altieri and Nichols, 2009). The combined use of cultural practices and low- or reduced-impact insecticides on non-target species might favour populations of parasites (Cortez-Mondaca and Valenzuela-Escoboza, 2013; Reitz et al., 2013).
In chrysanthemum cuttings, L. trifolii survived cold storage at 1.7°C for at least 10 days. Newly laid eggs of L. trifolii in chrysanthemums survived for up to 3 weeks in cold storage at 0°C (Webb et al., 1970). Webb et al. (1970) therefore proposed that chrysanthemum cuttings should be maintained under normal glasshouse conditions for 3-4 days after lifting to allow eggs to hatch. Subsequent storage of plants at 0°C for 1-2 weeks should then kill the larvae.
Gamma irradiation of eggs and first larval stages at doses of 40-50 Gy provided effective control (Süss et al., 1986; Yathom et al., 1991). The release of sterile L. trifolii males significantly reduced the number of offspring (Kaspy and Parrella, 2006). When the release of sterile males was combined with a release of the parasitoid Diglyphus isaia, the damage caused by L. trifolii and the size of the adult population were significantly reduced.
Yildrim and Unay (2011) noted that foliar fertilizers of fulvic acid and calcium nitrate combinations in tomato had a negative effect on L. trifolii population. Mortezaiefard et al. (2012) found that foliar applications of potassium silicate reduced L. trifolii populations on Gerbera jamesonii.
Lei et al. (2008) found that L. trifolii were found more often and made more feeding punctures on non-Bt transgenic cotton plants than on Bt cotton plants. Females oviposited more eggs on non-Bt cotton plants, but larval and puparial survival did not differ between Bt and non-Bt plants.
Sahu et al. (2006) reported that the leaf area of the lower leaves of the tomato plants was positively correlated with the percentage of leaves affected by L. trifolii, indicating that L. trifolii infestation increases with increasing leaf surface area. Thus, genotypes with narrow leaves would be less preferred by the species. Studies with similar purposes have been made ??in the castor oil plant Ricinus communis and cowpea Vigna unguiculata, among others (Eid, 2008; Hedge et al., 2009).
The nitrogen content in the leaves of a host plant of L. trifolii is an important component that influences in the susceptibility to attack (Altieri et al., 2005). Potassium and phosphorus reduce the susceptibility in potato crops and cause negative effects on pest (Minkenberg and van Lenteren, 1986; Facknath & Lalljee, 2005).
Integrated Pest Management
The selection, integration and implementation of different control tactics of the leafminer, based on the conservation of biological control, is sufficient for adequate management of L. trifolii (Liu et al., 2009, 2011; Cortez-Mondaca and Valenzuela-Escoboza, 2013). One of the most important steps is the use of selective or specific biorational insecticides, such as botanical extracts, soaps, minerals, entomopathogenic insecticides and growth regulators (Hernandez, 2009; Hernández 2011; Liu et al., 2009, 2011; Yildirim and Baspinar 2012). It is also important to apply insecticides so that they cause the least impact to natural enemies; for instance, some systemic insecticides can be applied in the seed or irrigation system (Cikman and Comelkcloglu, 2006; Nath and Singh, 2006; Kumar, 2010; El-Wakeil et al., 2013).
To avoid the introduction of L. trifolii (and other leaf miner species L. huidobrensis, L. sativae and Amauromyza maculosa [Nemorimyza maculosa]), EPPO recommends that propagating material (except seeds) of Capsicum, carnations, celery, chrysanthemums, Cucumis, Gerbera, Gypsophila, lettuces, Senecio hybridus and tomatoes from countries where L. trifolii occurs must have been inspected at least every month during the previous 3 months and found free from the pests (EOPP/EPPO, 1990).
Regulations could be tightened in the EU by including additional commodities under regulatory control, clearly prescribing the inspection procedures and the appropriate treatments to be used, and combining these with other measures, such as screening (EFSA, 2012). The application of protected zones to areas where L. trifolii is not yet present could help prevent further spread of the pest.
A phytosanitary certificate should be required for cut flowers and for vegetables with leaves.
ReferencesTop of page
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