Biology and Management of Dipteran Pests

## Harmful Diptera Pests in Garlic and Onion and Their Management

*Pervin Erdogan and Zemran Mustafa*

#### **Abstract**

Garlic (*Allium sativum*) is a hardy perennial member of the onion family presumably native to Central Asia; however, it has long been naturalized in southern Europe and throughout the world. Onion, on the other hand, is used all over the world, and its consumption depends mostly on the income level of consumers. It is an indispensable vegetable in the kitchen of many homes. Onions take third place in vegetable production after potato and tomato in Turkey. Mites, nematodes, and insect species cause damage to these plants, reducing considerably their yield. Among these pests, the most destructive are *Delia platura* Meigen (Diptera: Anthomyiidae) *and Delia antiqua* Meigen (Diptera: Anthomyiidae). The crop losses can sometimes reach up to 100%, depending on the crop and density of the pest. There are different methods to control these pests that vary by the pest type and the crop being applied. *D. platura* eat the contents of newly planted seeds, leaving empty seed shells and preventing germination. Also, *D. antiqua*. Young onions are particularly vulnerable. When the hide and bulb become entangled in the damaged plant, development stops, the plant turns yellow, and it breaks. Both pests are controlled using biological and chemical methods.

**Keywords:** onion, garlic, pests, biology, management

#### **1. Introduction**

Garlic (*Allium sativum*) is a hardy perennial onion family member most likely native to Central Asia, but it has been adopted in southern Europe and many other cuisines. It is one of the oldest cultivated vegetables used for its edible parts and as traditional medicine. As it is with onions, garlic is not used alone in cooking. It is one of the indispensable vegetables of the kitchens due to its taste and flavoring properties. Garlic teeth and leaves have appetizing, diuretic, antibacterial, respiratory and digestive tract antiseptic, and antithyroid effects. In addition, garlic has the properties of purifying bile, lowering blood sugar and lipids, cough suppressant, blood pressure lowering, anti-infective, and curative [1].

Onion is used all over the world, and its consumption depends mostly on the income level of consumers. It is an indispensable vegetable in the kitchen of many homes giving flavor and taste to the dishes. It regulates metabolism and strengthens the immune system against microbial diseases. It is known as a beneficial food for breastfeeding mothers due to its stimulating milk-production properties. In addition, onion is recommended to be included in the nutrition programs for children growth as it is an important energy source and a medicinal plant whose use dates back to ancient years [2].

There are different methods to control these pests that vary by the pest type and the crop being applied. *Delia antiqua* Meigen (Diptera: Anthomyiidae), *Delia platura* Meigen (Diptera: Anthomyiidae), *Thrips tabaci* L. (Thysanoptera: Thripidae), *Frankliniella occidentalis* (Pergande) (Thysanoptera: Thripidae), *Rhyzoglyphus* spp., (Acari: Acaridae), *Tyrophagus* spp. (Acari: Acaridae), *Aceria tulipae* (Keifer) (Acari: Eriophyoidea) reduce the production of garlic. Previous studies carried out in Turkey reported garlic pests as *Bactericera tremblayi* Wagner (Hemiptera: Psylloidea), *T. tabaci, F. occidentalis* (Thysanoptera: Thripidae), *Acrolepiopsis assectella* (Zeller) (Lepidoptera: Acrolepiidae), *Agriotes* spp. (Coleoptera: Elateridae), *Liriomyza* spp. (Diptera: Agromyzidae), and *Ditylenchus dipsaci* (Kühn) (Tylenchida: Anguinidae) in garlic cultivation areas [3]. However, in recent years, garlic producers have complained about side effects of the pest on garlic production. The pests have been common and caused significant loss of garlic yield. Some producers have even maintained that they did not get any yields. The most important of these are *D. antiqua* Meigen (Diptera: Anthomyiidae and *D. platura* Meigen (Diptera: Anthomyiidae). Product loss as high as 35% on average is caused when no control is applied. The crop loss may sometimes reach up to 100%, depending on the crop and density of the pests. There are different pest control methods that vary by pest type. Besides chemical control for *D. platura* and *D. antiqua*, environmentally friendly practices such as yellow sticky traps are utilized for successful control of the Diptera pests [4].

#### **2. Harmful Diptera species in garlic and onion**

#### **2.1 Seedcorn maggots (***D. platura***)**

Seedcorn maggot or the bean seed fly, *D. platura* is a small dipterous insect. This fly is almost identical to the cabbage maggot (*Delia radicum*) and onion maggot (*D. antiqua*). In previous studies, this fly was named *Hylemia platura*, *Hylemia cana, Hylemia cilicrura, and Hylemia similis* [5]. Although *D. platura* was first reported in Germany, it is now spread all over the world [6].

The adult is about 6 mm long, gray in color, its body is covered with black hairs, and the wings are held crossed on the abdomen at rest. The egg is white in color, 0.6–0.7 mm long, and 0.2 mm wide. The larva is ivory-colored, thin on the front, wide on the back, and 5–6 mm in length. Pupa is barrel-shaped (**Figure 1a**–**d**).

There is no mandatory diapause. After the adult emerges from the pupa, it feeds on flowers and other sweet substances and lays its eggs for 10–18 days. Usually, seedcorn is found in newly plowed and irrigated fields. *D. platura* lays its eggs singly or in groups. It also lays eggs on degraded organic material in soil cracks and especially in manure piles. The hatched larvae immediately go under the soil and begin to feed on organic materials. Meanwhile, the larva searches for germinating seeds in seed beds and quarries. It feeds on the stems and cotyledons of young seedlings. Total larval development time is 8–11 days depending on temperature. *D. platura* has three larval stages. When the larva completes its development, it leaves the plant it feeds on and becomes a pupa at a depth of 5–6 cm in the soil. Pupation period is 10 days at 20°C. A female lays 40–50 eggs in her lifetime. Male flies live 20–25 days, while females

*Harmful Diptera Pests in Garlic and Onion and Their Management DOI: http://dx.doi.org/10.5772/intechopen.106862*

#### **Figure 1.**

*Delia platura, (a) adult (wikipedia), (b) egg (Kilic, T), (c) pupae (Kilic, T.), (d) larva, (Erdogan, P), (e, f) damage (Erdogan, P).*

live 30–35 days. The development period from egg to adult is 35 days at 10°C and 16–17 days at 25°C. *D. platura* produces 3–4 generations per year [7].

The seedcorn maggot is a polyphagous pest that attacks over 40 different host plants [8]. *Allium* species have been reported to host *D. platura* [9, 10]. The larvae feed by opening galleries in the stems and newly formed cotyledons of seeds germinating in seed beds or quarries such as squash, melon, cucumber, beans, garlic, and other vegetables [11]. When the seedling emerges from the soil, only the cotyledons attract attention because the shoot tip is eaten. During the seedling stage, plants turn yellow, and dry. *D. platura* is the most destructive in garlic. Seedcorn maggots, according to Bessin [12], damage newly planted seeds by feeding on seed contents, often

leaving empty seed shells and preventing germination (**Figure 1e** and **f**). Seedlings that germinate despite the damage are spindly and have few leaves, and they die before maturing. Seedcorn maggots can sometimes be found tunneling within stems and germinating seeds [13]. Early planting dates, heavy cover crops, and cool-wet weather all contribute to seedcorn maggot damage [12]. *D. platura* also feeds in garlic. *D. platura* is the most harmful species in garlic fields, and it was spread throughout the entire area with the rate of harmful infestation increasing to 41% in some areas in Turkey [14]. In Ecuador, seedling losses of more than 60% have been reported 2 weeks after sowing. Plants that survive are weaker and less resistant to subsequent pest and pathogen attacks [15].

#### *2.1.1 Management*

### *2.1.1.1 Cultural control*


Kessing and Mau [11] suggested reusing manure and thoroughly incorporating it into the soil before planting. Planting when the soil and weather conditions favor rapid germination is a preventive measure against seed corn maggot infestation. According to Basin [12] planting, shallow planting, higher seeding rates, a wellprepared seedbed, and turning the cover early are all preventative measures for seed corn maggots (which renders the field less attractive to egg laying by flies). In another study, it was discovered that no-till fields are less likely to have seedcorn maggot problems because germinating seeds alone do not attract large populations of egglaying females [17].

### *2.1.1.2 Biological control*

Seedcorn maggots do not appear to have many natural enemies because they spend much of their life cycle underground. Isolated incidences of predation by

#### *Harmful Diptera Pests in Garlic and Onion and Their Management DOI: http://dx.doi.org/10.5772/intechopen.106862*

spiders, ants, and birds upon adults and of fungal diseases infecting larvae have been reported, but none of these predators or pathogens is considered significant in controlling the population of seedcorn maggots [16]. There are some studies on the biological control of *D. platura*. Ellis and Scatcherd [4] revealed that two nematode parasitic species, *Steinernema feltiae* and *S. krausseri*, can be used to control seed fly and onion fly. The same study determined that *S. feltiae* reduced numbers of bean seed fly larvae by about 50%. Moreover, onion seedlings treated with the entomopathogenic fungus *Beauveria bassiana* (strain ANT-03) based insecticides were less attacked by the pest [18]. Recently, entomopathogenic nematodes (EPNs) have been proposed as a potential control method for *Delia* species [19, 20].

#### *2.1.1.3 Chemical control*

In problematic areas, seed spraying should be performed or seed beds should be sprayed before planting seeds. If seedlings are infected after planting, spraying should be done to prevent new contamination [11]. Before planting, the seeds should be sprayed with drugs containing active substances Imidacloprid, Thiamethoxam, and Acetamiprid. If there is a need to spray during the green period, a chemical pesticide containing cyromazine as active ingredient can be applied.

#### **2.2 Onion fly (***Delia antiqua***)**

Onion maggot*, D. antiqua*, is a major underground agricultural pest, found throughout Asia, Europe, and North America due to its high and low temperature tolerance [21, 22].

Adult of *D. antiqua* is similar to adult houseflies but slightly smaller. The adult is 6 mm long and gray in color. There are five dark bands on the thorax of the adults. Wings of adults are transparent and unblemished; legs and antennae are black. *D. antiqua* eggs are matte, white in color, 1.5 mm long, and banana-shaped. There are 12 longitudinal lines on the egg [23]. Larva is white and legless. The head of the body is narrow and shaped like a carrot that widens toward the end. Mature larva is 6–8 mm. *D. antiqua* pupae are reddish, 6 mm long, and barrel-shaped (**Figure 2a**–**d**).

Adults begin to appear in mid-March and early April, depending on climatic conditions. Adults emerge gradually throughout the following month. Temperature affects the length of the preoviposition period. Flies kept at 25°C laid eggs in 10–24 days, with a 15-day average. Adults lay their eggs in groups of 10–20 or singly, usually in the place where they meet with the onion, on leaves, tuber bark, and cracks in the soil, within 10–20 days following emergence. Flies in captivity lay up to 123 eggs at the rate of 1–45 per day. Eggs hatch in 3–6 days, depending on temperature and proportional humidity. The emerging larvae enter from the junction of the onion and the beet and move toward the inside of the onion. The larval period is 15–20 days. The mature larvae pupate in the soil near the onion plant [24]. The pupal period is 10–15 days. In addition to the damage it causes by feeding on the plant tissue, the larva causes decay with various bacteria it carries. The damage is especially greater in young onions. In the damaged plant, the development stops, the plant turns yellow, and it breaks when the hide and bulb are caught. Since the shallots that come out of the onion seeds planted for the purpose of growing shallots are frequent, the damage is more common (**Figure 2e** and **f**) [25]. The seed value of shallots is lost, and they rot. Onions, garlic, and bulbous flowers are their hosts [26, 27].

#### **Figure 2.**

*Delia antiqua, (a) adult (Pest and diseases image library, (b) egg (Pest and diseases image library), (c) larva (Rasbak), (d) pupa (Rasbak), (e, f) damage (wikipedia.org).*

#### *2.2.1 Management*

#### *2.2.1.1 Cultural control*

There are numerous non-chemical approaches to *D. antiqua* management. Crop rotation, crop sanitation, delayed planting, protective barrier installation, biological control, and sterile insect technique are examples of these.


#### *2.2.1.2 Biological control*

Pests of *Delia* spp. are vulnerable to a variety of natural enemies and biocontrol agents. Many (60–100) species of staphylinid and carabid ground beetles, generalists that feed on eggs and early instars, prey on *Delia* spp. [32]. In addition to feeding on eggs, some staphylinid beetles, such as *Aleochara bilineata* Gyllenhal (**Figure 3a**) and *A. bipustulata* Linnaeus, parasitize *Delia* pupae [32]. *Aphaereta pallipes* Say (**Figure 3b**) is a braconid fly, which has a diverse host range and successfully parasitizes *D. antiqua.* Moreover, other biocontrol agents of *Delia* include predators and parasitoids as well as entomopathogenic fungi (EPFs) and nematodes (EPNs). *Entomophthora muscae* (**Figure 3c**), *Metarhizium anisopliae* (**Figure 3d**), and *B. bassiana* (**Figure 3e**) have all been found to be capable of infecting and killing *Delia* sp. pests in EPF screenings [33–35]. Similarly, EPNs, including commercially (Nemaplus, Biotem, Larvanem, Capsanem, etc.) available species such as *S. feltiae* (**Figure 3f**) and *Heterorhabditis bacteriophora* (**Figure 3g**) infect *D. antiqua* [36].

#### *2.2.1.3 Sterile insect technique*

Sterile insect technique (SIT) is a pest management technique that involves rearing and sterilizing large numbers of a pest before releasing them into the wild population. Sterile males compete with fertile males for the opportunity to mate with fertile females, reducing the number of viable offspring [37].

#### *2.2.1.4 Chemical management*

Growers rely heavily on chemical management tactics due to the practical and economic limitations of non-chemical management tools. Currently, *Delia antiqua* is managed principally with insecticides applied as seed treatments and in-furrow applications of chlorpyrifos at planting [38]. Because flies move and disperse during the day, sprays targeting flies are unlikely to contact the pest and are not recommended for fly management, although some growers continue to use them [29].

#### **2.3 Leaf miners (***Liriomyza* **spp.)**

Adults are 1–2 mm long, gray-black in color. Larvae are up to 3 mm, white yellow in color, and clear (**Figure 4a**–**d**). Adults are on all leaves of the plant, and larvae are gallery life miner adults and live 25–30 days. A female lays about 400 eggs between two epidermises of the leaf. Larvae that emerge from egg feed by opening various galleries in the leaf epidermis. The mature larva, which has completed its development, leaves itself the soil to become pupae [39].

#### **Figure 3.**

*(a) Aphaereta pallipes, (b) Aleochara bilineata, (c) Entomophthora muscae, (d) Metarhizium anispliae, (e) Beauveria bassiana, (f) Steinernema feltiae, (g) Heterorhabditis bacteriophora (wikipedia.org).*

*Harmful Diptera Pests in Garlic and Onion and Their Management DOI: http://dx.doi.org/10.5772/intechopen.106862*

#### **Figure 4.**

*Liriomyza spp. (a) Adult, (b) egg, (c) larva, (d) pupa (Payne J.A.), (e, f) damage (wikivand.com).*

The females inflict small wounds on leaves, feed on the sap that comes out of it, and cause cell disruption. Feeding causes loss of healthy leaf tissue, so that plant cannot capture enough sunlight and often becomes infected with disease. Plants often fail to grow or produce crops. Then damaged areas turn yellow and dry and the leaves fall (**Figure 4e** and **f**). They delay development in young plants and seedlings. They cause quality and yield loss. They are harmful to tomatoes, eggplant, peppers, beans, peas, broad beans, lettuce, zucchini, cucumbers, spinach, onions, and leeks [39].

#### *2.3.1 Management*

#### *2.3.1.1 Cultural control*


#### *2.3.1.2 Biotechnical control*

It can be used as yellow sticky traps to control adults of *Liriomyza* spp.

### **3. Conclusions**

Garlic and onion foods have been known for their health benefits since ancient times and are frequently used for both health promotion and flavor purposes. The positive effects of garlic and onion also called a natural antibiotic among people are innumerable, and new benefits continue to be discovered every day. Both play a huge role in the health of people, such as reducing the risk of various types of cancer, improving mood, and maintaining skin and hair health. There are many pests that limit the production of these plants. For example, *D. platura, Delia antiqua, Liriomyza* spp., and *Ditylenchus dipsaci*. Especially, *D. platura, D. antiqua* cause a high rate of product loss when these two pests are not controlled. In order to prevent product loss caused by *D. platura* and *D. antiqua*, correct control methods must be applied. Onion and garlic farmers should be informed about these pests and their control.

### **Author details**

Pervin Erdogan1 \* and Zemran Mustafa2

1 Faculty of Agricultural Sciences and Technology, Plant Protection Department, Sivas University of Science and Technology, Sivas, Turkey

2 Faculty of Agricultural Sciences and Technology, Plant Production and Technologies Department, Sivas University of Science and Technology, Sivas, Turkey

\*Address all correspondence to: pervinerdogan@sivas.edu.tr

© 2022 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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#### **Chapter 5**

## Walnut Husk Fly (*Rhagoletis completa* Cresson), the Main Burden in the Production of Common Walnut (*Juglans regia* L.)

*Aljaz Medic, Metka Hudina, Robert Veberic and Anita Solar*

#### **Abstract**

The walnut husk fly (*Rhagoletis completa* Cresson) is the most important pest of walnuts (*Juglans regia* L.). It causes economically significant crop losses (up to 80% yield loss) in many growing regions, including the United States and most European walnut-producing countries. This chapter describes the impact of pest infestation on yield quantity and quality along with the current geographic distribution of the pest. Its bionomy and infestation symptoms are described in detail. An overview of monitoring and control methods used is also provided, and new methods that may prove useful for walnut husk fly control are listed. Monitoring the occurrence of the pest is the most important part of controlling the walnut husk fly, as only with an effective monitoring system can insecticides be applied at the appropriate time. Emphasis is placed on biotic protection and the possible role of phenolic compounds in cultivar resistance to walnut husk fly. Other control methods (non-chemical, mechanical, and biological control) are also gaining importance in pest management as more and more active substances in pesticides are regulated or phased out each year. Mechanical control methods are more or less only suitable for walnuts grown in the protected areas.

**Keywords:** biotic protection, phenolic compounds, cultivar resistance, yield quality, yield quantity, bionomy, control methods

#### **1. Introduction**

The walnut husk fly (*Rhagoletis completa* Cresson) belongs to the group of fruit flies (Diptera: Tephritidae), which are the most important pests worldwide [1]. Fruit flies are considered one of the most important pests in the world due to the large economic impact and strict quarantine restrictions imposed by several countries. Larval feeding and oviposition by females render fruits or vegetables unusable and inedible [2], and losses can be as high as 80% of the yield [1]. Apart from the economic level, the fruit fly is also important at the ecological level, as it can displace native species or compete with them for resources [2].

*Rhagoletis* is a genus of Tephritidae fruit flies, which includes about 60 species. The walnut husk fly is the most destructive pest of the genus *Rhagoletis*, along with the European cherry fruit fly (*Rhagoletis cerasi* Loew) and the apple maggot (*Rhagoletis pomonella* Walsh) [1]. Fruit flies of the genus *Rhagoletis* have a specific combination of wing patterns consisting of pointed black or yellow postocular setae, four or five transverse bands, and posteroapical lobe along the anal vein. The male specimens have a long, saber-like surstyli genital, whereas the female specimens have a terminalia with a short oviscape that forms a soft, desclerotized T-shaped area at the apex [3]. Flies of the genus *Rhagoletis* are distributed throughout North and South America, Europe, and parts of Eurasia [4]. *Rhagoletis* species are usually found in temperate, mesic environments where rainfall is abundant. Most *Rhagoletis* species are univoltine, and only a few are multivoltine, producing a small second generation. Most species spend 8 to 10 months in the soil under host trees, from where they emerge from pupae between May and August (Northern Hemisphere). The female specimens emerge before the males. After emerging, they mate within 1 to 2 weeks on or near the fruit of their host plants. Both sexes are opportunistic feeders as their foods include fruit juice, exudates from extrafloral nectaries, plant leachates, bird excretions, homopteran honeydew, and possibly yeasts and bacteria. Male specimens are territorial and wait on host fruits, which they defend against other males, for the arrival of female specimens to mate with them. *Rhagoletis* species do not exhibit true courtship, as there are no predictable or elaborate behaviors that lead to mating and mounting. To mate, males simply attempt to hump the back of females. Males attempt to mate with females that are ovipositioning. The females then individually deposit the eggs directly under or on the skin of the fruit. In addition to depositing the eggs, females also mark the fruit with pheromones that prevent further oviposition and thus competition for resources within the species. After the larvae hatch from the eggs, they begin to feed on the pulp of the fruit, causing the fruit to rot and become unmarketable and inedible. There are three larval stages in *Rhagoletis* species. The final larval stage usually leaves the damaged fruit and burrows into the soil, where it pupates. It is believed that adults can survive up to 1 month in the wild, while the duration of the stage from egg to pupation depends on temperature and usually lasts between 4 and 10 weeks [5–9].

#### **1.1 Hosts, origin, and distribution**

Walnut husk fly belongs to the *suavis* species group. It has been classified there along with *Rhagoletis suavis* Loew, *Rhagoletis juglandis* Cresson, *Rhagoletis boycei* Cresson, *Rhagoletis ramosae* Hernández-Ortiz, and *Rhagoletis zoqui* Bush [9, 10]. Ten plants have been identified as hosts for the walnut husk fly. English or Persian walnut (*Juglans regia* L.) is the most commercially important, while black walnut (*Juglans nigra* L.) is the native host and is of lesser commercial importance. Other hosts include little or Texas walnut (*Juglans microcarpa* Berlandier), Arizona walnut (*Juglans major* (Torrey) Heller), Nuevo León walnut (*Juglans hirsuta* Manning), Hinds' black walnut (*Juglans hindii* Rehder), California black walnut (*Juglans californica* S. Watson) [6, 9], and Mexican walnut (*Juglans mollis* Engelm) [11]. Walnut husk fly has also been reported to infrequently attack peach (*Prunus persica* L. Batsch) [12], and Midland or English hawthorn (*Crataegus laevigata* (Poir.) DC.) [13].

The walnut husk fly is native to North America, to be more precise Midwestern USA and northeastern Mexico [9, 10]. It was first described and characterized by Cresson in the late 1920s [14]. From its native region in the Midwestern United States (Iowa, Kansas, Oklahoma, Minnesota, and Texas), it gradually spread to the other regions. In 1922, it

*Walnut Husk Fly (*Rhagoletis completa *Cresson), the Main Burden in the Production… DOI: http://dx.doi.org/10.5772/intechopen.106046*

was reported in California [12], from where it spread to the southern areas of California and northward to Washington State [15] and later to southern British Columbia (Canada) [5]. In Mexico, however, it is still restricted to the northeast, particularly to the states of Tamaulipas, Nuevo León, and Coahuila. In Nuevo León, walnut husk fly is restricted to *Janirella hirsuta* in higher-elevation areas within the canyons of the Sierra Madre Oriental [16], whereas in Tamaulipas and Coahuila it is infesting *J. mollis* [11].

It is believed that walnut husk fly spread from North America to Europe *via* global trade routes. In 1988, the walnut husk fly was documented for the first time in Europe, in Switzerland in the Ticino region [1, 17]. Later, the fly subsequently spread to neighboring countries. In 1991, it was reported near Venice and in the Friuli region [18], from where it rapidly spread throughout Italy. By 1992, it was confirmed in Milano, Novara, Varese, Pavia, and Sondrio [19]. From Italy, the walnut husk fly then spread across walnut-growing areas in Italy, and in 1997, the walnut husk fly was documented for the first time in Slovenia near Nova Gorica [20]. Documented observations of the walnut husk fly then increased each year: Croatia (2004), France (2007), Austria and Germany (2008), Albania (2010), Hungary (2011), Bosnia and Herzegovina, and Spain (2013) [1, 21–23]. Lacking natural predators, the walnut husk fly is expected to spread to all walnut-growing areas in Europe and Asia in the next few years, severely affecting walnut production [1].

#### **1.2 Biology**

To determine the species morphologically, a binocular microscope is recommended for diagnosis. Magnification for adult specimens is x10 and for larvae and aculeus x200. Only adult specimens can be reliably identified, while identification of eggs, larvae, and pupae is not reliable. Morphological identification of adult specimens can be done according to the EPPO (European and Mediterranean Plant Protection Organization) protocol [24], and that of larvae according to the protocol for the identification of larvae (third instar stage) [25]. The walnut husk fly life cycle, along with the damage caused can be seen in **Figure 1**.

Similar to other species of Tephritidae, the adult specimen of walnut husk fly is small, reaching a length of 4.0–6.5 mm. Both female and male specimens are of about the same size. The head of the walnut husk fly is completely yellow. It has a yellow dot on the posterior base of the thorax. Its wings are transparent and have three characteristic dark brown stripes. The last stripe is an elongated L that begins at the leading edge of the wing. Female specimens can be distinguished from males by a pointed abdomen with an ovipositor and by the color of the first leg segment. On males, the first leg segment is brown to black, whereas on females, it is pale yellow [9, 24]. A detailed morphology to distinguish *Rhagoletis* species is available at EPPO [24].

The walnut husk fly is univoltine (has only one generation per year). It usually overwinters in the soil under or near the host tree, as a pupa, however in years of high population density, individual specimens migrate elsewhere. Passive dispersal (wind, transport vehicles, etc.) is also likely. The life span of adult specimens is similar to other *Rhagoletis* species and is up to 40 days. Adult specimens emerge from the ground from mid-July to late September. Peak emergence is between late July and late August. In the host tree, they usually stay in the shaded part of the canopy where there are plenty of fruit. As previously seen with other *Rhagoletis* species, male specimens are territorial. They wait on the host fruit, for the arrival of the female specimens to mate with them. Mating occurs within 6 to 8 days after emergence from the soil. Females begin laying eggs 10 days after emerging from the soil.

#### **Figure 1.**

*Adult specimen of walnut husk fly (A), early signs of larvae damage (early: Bottom left, late: Bottom right, uninfested fruit: Above) (B), larvae (C), life cycle of walnut husk fly (D), and damage to shells and kernels (healthy left and damaged right) (E). \*Photo of pupae provided by AGES/A. Egartner.*

A female can lay between 300 and 400 eggs in her lifetime. They usually lay eggs in groups of 15 to 20 per fruit. As with other *Rhagoletis* species, female specimens release a pheromone after laying eggs that prevents other females from laying eggs on the same fruit (walnut husk), which thus reduce larval competition for resources. One female can infest up to 20 fruits per season. After the females lay eggs on the fruit, the larvae hatch from the eggs in 3 to 10 days, depending on climatic conditions, especially temperature. The larvae are dirty white in color and have neither head nor legs. They reach a length of 8–10 mm. Once the larvae hatch, they bore into the walnut husk, where they make burrows and feed on the flashy pericarp (inner husk). Larval development takes between 30 and 40 days, during which they strip twice. Once they reach the last strip and are fully mature, they drop out of the husk onto the ground. They berry into the soil and pupate at a depth of a few centimeters. The pupation allows them to overwinter. The following year, more than 90% of the adults emerge from the soil and repeat this cycle again, while less than 10% of the pupae spend another season in diapause [1, 9, 26–29].

#### **1.3 Economic damage**

Economic damage is caused by the larvae of the walnut husk fly when they feed on the pericarp. This softens and blackens the husk, making it soft, sticky, and black. Once the damaged husk dries, it sticks to the endocarp (maturing nut) and is very difficult to remove or wash off. In severe infestations, the larvae may completely destroy the pericarp, leaving only the withered black exocarp around the shell [1]. Infestation causes the release of tannins from the damaged husks, resulting in black spots on the

#### *Walnut Husk Fly (*Rhagoletis completa *Cresson), the Main Burden in the Production… DOI: http://dx.doi.org/10.5772/intechopen.106046*

shells and reducing their commercial value. Apart from the damaged shell, infestation alters the internal quality of the nut, affecting its flavor and metabolic composition, making it bitter and thus reducing its commercial value [30, 31]. In early infestations, kernel (seed) development is severely inhibited as larvae attack the conductive tissues of the fruit stalk, resulting in malnutrition of the fruit. As a result, these fruits fall off before they reach maturity, or they do not fall off the tree at all and remain on the tree through the winter. Heavily infested fruit can also facilitate the entry of pathogens into the edible interior of the nut, particularly the bacterium *Xanthomonas campestris* pv. *juglandis* (Pierce (Dye)) and the fungus *Marssonina juglandis* (Lib.). This causes the kernel to shrink, lose weight, rot, mold, and deform the kernel, resulting in significant or total yield loss [30–33]. For this reason, the early emergence (mid-July to mid-August) of the walnut husk fly is the most dangerous, whereas the later emergences are not as dangerous because the nuts ripen before the larvae complete their development and cause serious damage. Because the larvae do not complete their development, fewer tannins are released from the husk that would blacken the shell or affect the internal quality of the nuts [30, 31]. Although late infestation usually does not damage the kernel, it interferes with the natural separation of the pulp from the nutshell, making marketing cumbersome and impractical. In addition, black stains must be removed from the shell with high water pressure or the nuts must be bleached because consumers are unwilling to buy stained nuts [4, 32]. In years when vegetation is delayed by cold spring temperatures, the walnut husk fly also reaches its peak attacks later, even in the first days of September. In these cases, even the September emergence of the walnut husk fly causes major damage, especially to late ripening cultivars.

In orchards where walnut husk fly is present, and if left uncontrolled, the damage can be visible on 74 to 91% of the husks [27] and yield loss can be as high as 80% [1]. However, yield losses vary between cultivars.

#### **2. Influence of walnut cultivar on resistance to the walnut husk fly**

Some cultivars were found to be more tolerant to walnut husk fly attacks, with first studies reaching in the 1930s, just after the emergence of the pest in California [34]. At the start of the research and with the lack of analyzing equipment that is available nowadays, cultivar resistance was believed to be depended on the hardiness of walnut husk at the time of oviposition activity of the fly [34]. Later on, it was found that walnut husk flies prefer cultivars that produce larger and heavier fruit. Overall, fruit weight was also correlated with the pupal weight and diapause length, as offspring that developed in larger fruit likely accrued fitness advantages over offspring that developed in smaller fruit. Adult fly longevity was reported to be influenced by cultivars and not any particular physical fruit characteristic [35]. Fruit weight has also been documented to influence infestation rates in other Tephritidae species [36, 37]. There are, however, inconsistent and contradicting results between various authors that were comparing the same cultivars across different orchards and years [35, 38]; therefore, no clear conclusion on which cultivar is more or less susceptible to walnut husk fly attack could be drawn. It was, however, suggested that environmental conditions (soil moisture levels, favorable temperatures, etc.) could affect infestation patterns, by simultaneously influencing fly development rate (delaying or accelerating sexual patterns) or walnut growth (delay in the phenophases) [33, 35, 38, 39].

One possibility of cultivar resistance could be the content and composition of phenolic compounds in the husk at the time of infestation. Phenolic compounds are secondary metabolites found in all plant tissues, as they play a major role in physiological processes, growth, and durability of the plant. Their key role in the plant is related to plant defense against biotic and abiotic stresses [40]. In the past, higher total phenolic content was associated with higher stress tolerance in plants [41, 42]. However, in recent years, increasing attention has been focused on individual phenolic compounds and groups as more and more studies [40, 43–45] show that plants respond to pathogens with only selected individual phenolic compounds and groups. It is hypothesized that some individual phenolic compounds and groups respond more rapidly and are better adapted to short-term stress conditions, whereas others require more time to form and are better adapted to long-term stress conditions. In addition, there is a possibility that plant/cultivar resistance is not related to the total phenolic compound content, but rather is due to the reaction time of the plant and the speed with which it recognizes the pathogen and responds quickly to infection containing it while it has not yet spread. Cultivar resistance could be due to the composition of phenolic compounds or to the reaction time of the plant to the infection, as already observed in walnut anthracnose (*Ophiognomonia leptostyla*) [43] and walnut blight (*X. campestris* pv. *juglandis*), where flavonols, flavanols, and naphthoquinones were observed to have the biggest role in walnuts defense mechanisms [40]. Unfortunately, to date, there have been no investigations of this claim, only studies examining the effects of walnut husk fly infestation on kernel quality and its composition [30, 31].

Because cultivar resistance is usually associated with a higher total phenolic content or the phenolic content of certain groups, breeding and selection of new cultivars usually involves breeding cultivars with a higher content of phenolic compounds. Since breeding is a long-term process that can take more than 20 years for walnuts, and farmers cannot tolerate the damage caused by the walnut husk fly for that long duration, the use of pesticides and other technical measures is crucial and necessary. Since walnut husk fly causes up to 80% yield loss, we cannot imagine walnut cultivation without these measures. However, yield losses vary between cultivars.

#### **3. Walnut husk fly control**

Governments, agencies, and policymakers in the agricultural sector are constantly faced with the risk of epidemics and pest outbreaks. In the context of global climate change and world trade, there is growing concern about the environmental and economic impacts of non-indigenous invasive species of parasites and pathogens on crops. With globalization and the growing number of trade routes, the problem is increasing every year. Alien insect species have become an increasing problem worldwide due to their significant ecological and economic impacts [1]. According to some estimates, invasive species cause around €19.64 billion in damage and losses each year in Europe alone [46]. Strict regulations and quarantines are in place to prevent these outbreaks, but ultimately the pathogen finds its way across national borders. At the same time, scientific advances and growing public concern about human and environmental health are prompting legislators and policymakers to enact and enforce pest control regulations [1, 4]. Every year, more and more active substances in the plant protection products are regulated or withdrawn from use. And in the event of an outbreak of an invasive pathogen, the effective response can be very limited. To respond effectively to pathogen outbreaks, alternative plant protection methods

must be applied. To be successful, accurate data on the pathogen itself (persistence, overwintering, developmental stages, timing of infestation and extent of damage, reproduction, etc.) are required in addition to varietal resistance. Only with all this detailed information can we successfully control the pathogen and its damage [1].

#### **3.1 Monitoring methods**

Walnut husk fly control is primally based on successful monitoring of the pathogen to adapt phytosanitary treatments to the occurrence of the pest. Occurrence of the walnut husk fly is usually determined by capturing the adults, but larvae can also be detected. Monitoring of the adult walnut husk fly begins in mid-July. At this time, yellow sticky traps must be suspended in the tree canopy. According to field observations, the efficiency of observations is better when the traps are suspended higher in the tree canopy [1]. There are a number of different traps from different vendors that can be used. Typically, yellow rectangular PVC sticky traps are used that have been shown to be effective in capturing a range of dipterans [27]. The yellow sticky traps are used alone or in combination with attractant, which has been shown to be more effective [27]. Yellow sticky traps were first used in the 1980s in the U.S. and were proven successful in monitoring walnut husk fly populations [47]. Traps are baited with 3 g of ammonium carbonate to increase trap attraction (**Figure 2**) [27].

For an orchard of 1 ha, one to two yellow sticky traps have proven sufficient. It is recommended to place one of the two traps 2 m above the ground and the second 5–6 m above the ground in the canopy. The traps are visually inspected twice a week or every 3 days for the presence of the pest. It is important to remove the trapped flies and re-coat the sticky layer or change the attractant according to the manufacturer's instructions. Normally, the traps are renewed every 3 weeks, while ammonium carbonate is added every week. When the first adult specimens are observed on the traps, it is important to begin the phytosanitary treatments. After the first insecticide application, the pests must be removed from the traps and the traps monitored once a week for further pest occurrence [27, 48]. Monitoring costs associated with walnut husk fly are estimated to be about €75 per hectare, including labor and materials [27, 48].

Larvae can also be detected, but the method is not as reliable and the major initial infestations that cause the most damage are missed because the larvae are already developing in the husk. To detect the larvae, you must visually inspect the

#### **Figure 2.**

*Two types of traps that are typically in use, PHEROCON® Trécé trap (A), and REBELL® Amarillo trap (B).*

fruit surface and locate the damage where the adult females have poked the holes and laid the eggs in the husk. These holes are not easy to see and can be easily missed. Later observation may be easier as the husk begins to blacken from larval feeding activity [1].

#### **3.2 Control methods**

#### *3.2.1 Insecticide use*

Following the detection of the first walnut husk fly specimens, walnut producers must begin chemical control. Commercially available insecticides containing the active ingredients dimethoate and fenitrothion are considered efficient in controlling the walnut husk fly. Their efficacy is comparable and both showed efficacy when applied at 1500 g of active ingredient per hectare. Apart from the two mentioned, imidacloprid, cyclaniliprole, thiacloprid, zeta-cypermethrin, chlorpyrifos, spinosad, fosmet, and bifenthrin as active ingredients are considered successful in walnut husk fly control. Considering nut prices, average yields, and insecticide application costs, 1 to 2 insecticide treatments are considered economically viable. When pesticides are sprayed at the right time, two applications are considered sufficient to maintain infestations at acceptable levels. However, effective timing of pesticide application can only be achieved with the effective monitoring techniques. A single insecticide application can save about 50% of production in orchards [27]. Current guidelines assume that an attractant (protein bait) is added to insecticide applications to improve their effectiveness. In the canopy, walnut husk flies feed on the protein baits and also ingest the insecticide. Wider nozzles are recommended for easier application because they form larger droplets. Because of the difficulty of applying insecticides to walnuts, it is currently recommended that only 1/3 of the tree canopy from the north to east sides be sprayed with the insecticide and attractant combination. If walnut husk fly is observed again in the yellow sticky traps after the first insecticide application, the insecticide application may be repeated up to two times depending on country guidelines. The last application must be made at least 3 weeks before the walnuts ripen to avoid residues of the insecticide [48]. Because insecticide application to walnut trees is difficult, research is being conducted on different application techniques: special nozzles, drone application, direct trunk injection. One of these is direct trunk injection of abamectin, which has recently been shown to be a viable method of controlling walnut pests [49].

In addition to traditional insecticidal crop protection methods, a walnut husk fly trap has recently become available. It contains an attractant and an insecticide that first attracts the pathogen, and once in the trap, the insecticide kills the pathogen. The lure is hung in the canopy, after the presence of the walnut husk fly is confirmed with the yellow tapes. The traps are a closed system where the insecticide is impregnated on the inside of the lid, while a bag containing the attractant is in the lower container. However, this method is more suitable for gardeners or protection of individual plants, as 50 to 100 traps are needed for a 1-hectare orchard [48].

#### *3.2.2 Non-chemical control methods*

With the absence of registered phytosanitary products for walnut husk fly control, more and more alternative methods are being tested. In addition to chemical ones, there are also some non-chemical control methods that are particularly useful in

*Walnut Husk Fly (*Rhagoletis completa *Cresson), the Main Burden in the Production… DOI: http://dx.doi.org/10.5772/intechopen.106046*

protected areas, urban areas, and areas where the use of insecticides is not appropriate or the use of pesticides is not considered a safe and viable option.

In organic orchards, the use of clay (calcinated kaolinite) has proven effective as it physically protects the fruit. However, even in low-rainfall regions, four to five applications per year are required. In addition, the application is only suitable for small walnut trees, as it is not possible for larger walnut trees [1].

One of the most efficient methods is ground cover under the canopy. This prevents adult specimens from emerging from the ground and thus prevents further oviposition on the husk. However, this method is considered effective only when the entire ground under the tree is covered; otherwise, females from neighboring trees will disperse to these trees [48, 50].

To control walnut husk fly populations, regular removal and burning of infected fallen fruit along with shallow tillage in the spring and fall under tree canopies may be effective. In the fall, shallow tillage of the soil must be done immediately after collecting walnuts. This will destroy the larvae before they can crawl into the soil and pupate. In the spring, it is recommended that the soil be tilled in April to destroy the pupae before the adult emerge. Tillage in the fall must be done 5 to 10 cm deep in the soil and 10 to 15 cm in the spring [48, 50].

#### *3.2.3 Biological control*

Among other methods, biotic methods of walnut husk fly control are also an option. Studies have shown that two entomopathogenic fungi are available to control adult specimens of the genus *Rhagoletis*, one is *Beauveria bassiana* and the other is *Metarhizium anisopliae* [51, 52]. The effect of entomopathogenic fungi on larvae and pupae is limited, but they prove effective against adult specimens. To date, no studies of walnut husk fly control with entomopathogenic fungi are known. However, there are some data on the control of cherry fruit fly (*Rhagoletis cerasi*) with the entomopathogenic fungus *Beauveria bassiana*. Application of *Beauveria bassiana* has been shown to reduce cherry fruit damage by 25–30% when applied to larvae and pupae and by up to 65% when applied to walnut husk fly adults foliar [53]. In addition to entomopathogenic fungi, the use of entomopathogenic nematodes of the genus *Steinernema* in *Heterorhabditis* has also been mentioned as a successful biotic agent in the control of various insects [54]. As with entomopathogenic fungi, the effect of entomopathogenic nematodes on walnut husk fly control has not been studied. However, there were some studies on the effect of entomopathogenic nematodes of the genus *Steinernema* on the control of species related to the walnut husk fly, *Rhagoletis indifferens*. The results showed that entomopathogenic nematodes of the genus *Steinernema* were considered successful biotic agents in controlling larvae (up to 80% mortality) and adults (up to 50% mortality), whereas no effect was obtained on pupae [55]. The first optimal time to apply biotic agents against walnut husk fly would be when the walnut husk fly begins to molt from pupae and fly out of the ground (late spring). To determine the proper time to apply biotic agents, the walnut husk fly must be monitored using the yellow sticky panels. The second effective time to apply biotic agents to walnut husk fly would be when the larvae begin to move into the soil (fall) to pupate [1, 56]. Recent work has shown that entomopathogenic nematodes are compatible with insecticides, so combining the two methods could improve efficiency in controlling walnut husk fly [56].

Previously, the possibility of a natural predator, the parasitic wasp *Coptera occidentalis* Muesebeck (Hymenoptera: Diapriidae), was considered. *C. occidentalis* is native

to California, USA, and parasitizes the pupae of some species of the genus *Rhagoletis* [57]. *C. occidentalis* has been reported to parasitize the walnut husk fly, along with *Rhagoletis cingulata*, *R. indifferens*, and *Ceratitis capitata*. In the late 1970s, a massive propagation of *C. occidentalis* began in California to control the walnut husk fly. In the 1980s, the parasitoid was released into the wild for the first time. Although *C. occidentalis* has been continuously released for 30 years, its efficacy in controlling walnut husk fly is considered insufficient [58]. This is thought to be due to the particular bionomy of *C. occidentalis*, as it is a parasitic predator that parasitizes pupae in the soil. Due to the insufficient concentration of attractants (kairomones) emitted from the soil by the pupae, the results are not optimistic [54, 59]. However, the species has been introduced in Slovakia as a biotic agent for control of *R. cerasi* [59]. The second parasitic wasp-parasitizing species of the genus *Rhagoletis* is *Diachasmimorpha juglandis* Muesebeck (Hymenoptera: Braconidae). A solitary parasitoid that parasitizes the pupae and larvae of the genus *Rhagoletis*. *Dorcaschema juglandis* finds its prey by sensing the volatile compounds released by infested fruit. However, *D. juglandis* has not been included in the biotic protection program for walnut husk fly control [60]. Close monitoring of native parasitoids in regions where walnuts are grown has been suggested. The species of natural enemies (*Coptera occidentalis*, *Diachasmimorpha juglandis*) of the walnut husk fly could be considered as sufficient alternatives to the use of insecticides in the control of walnut husk fly [54].

#### **4. Conclusions**

The walnut husk fly is not a new pest in the United States, but it is fairly new to Europe (1988). Since the introduction of this invasive species, it has spread to almost all walnut-growing areas. Where it is not currently present, it will most likely emerge in the next few years. In the absence of natural predators, it reproduces at a very high rate and causes very high yield losses (80%) in orchards. Monitoring the occurrence of the pest is the most important part of controlling the walnut husk fly. Only with an effective monitoring system, we can apply insecticides at the appropriate time. Since the application of these insecticides is very difficult for adult walnut trees, new methods for insecticide application are being researched (drone application, special nozzles, trunk injection, etc.). As more and more active substances in pesticides are regulated or phased out each year, other control methods are also gaining importance in the pest management. Mechanical control methods are more or less only suitable for walnuts grown in the protected areas, in urban areas and in areas where the use of insecticides is not appropriate or the use of pesticides is not considered a safe and viable option. The use of biotic control agents needs to be further investigated as it may also be an option, especially in organically managed orchards. In addition, the role of phenolic compounds needs further investigation, as little or no research has been conducted. The implications of this research would greatly benefit our understanding of pathogen control, as well as benefit breeders who could easily determine which walnut cultivars have the ability to resist walnut husk fly attacks.

#### **Acknowledgements**

This study is a part of programme P4-0013-0481, which is funded by the Slovenian Research Agency (ARRS).

*Walnut Husk Fly (*Rhagoletis completa *Cresson), the Main Burden in the Production… DOI: http://dx.doi.org/10.5772/intechopen.106046*

#### **Conflict of interest**

The authors declare no conflict of interest.

### **Author details**

Aljaz Medic\*, Metka Hudina, Robert Veberic and Anita Solar Biotechnical Faculty, Department of Agronomy, University of Ljubljana, Ljubljana, Slovenia

\*Address all correspondence to: aljaz.medic@bf.uni-lj.si

© 2022 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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### *Edited by Sarita Kumar*

This volume, *Advances in Diptera - Insight, Challenges and Management Tools*, highlights the biology, life stages, physiology, and significance of dipteran flies. Their importance as agricultural pests and disease vectors, and their management using various conventional and advanced tools is examined. Each of the five chapters covers a significant aspect in the field of diptera, including the physiology of dipteran diapause, the biology and management of mosquitoes using various traditional and innovative approaches, and the bionomics of onion, garlic and walnut pests. We hope that this book will interest students and researchers, and help them to recognize potential research areas.

Published in London, UK © 2022 IntechOpen © Backiris / iStock

Advances in Diptera - Insight, Challenges and Management Tools

Advances in Diptera

Insight, Challenges and Management Tools

*Edited by Sarita Kumar*