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Cereal Aphid Parasitoids in Europe (Hymenoptera: Braconidae: Aphidiinae): Taxonomy, Biodiversity, and Ecology

Željko Tomanović
Nickolas G. Kavallieratos
Zhengpei Ye
Erifili P. Nika
Andjeljko Petrović
Ines M. G. Vollhardt
5 and
Christoph Vorburger
Faculty of Biology, Institute of Zoology, University of Belgrade, 16 Studentski trg, 11000 Belgrade, Serbia
Serbian Academy of Sciences and Arts, Knez Mihailova 35, 11000 Belgrade, Serbia
Laboratory of Agricultural Zoology and Entomology, Department of Crop Science, Agricultural University of Athens, 75 Iera Odos Str., 11855 Athens, Greece
Environment and Plant Protection Institute, Chinese Academy of Tropical Agricultural Sciences, Xueyuan Road 4, Haikou 571101, China
Agroecology, Department of Crop Science, Georg-August University Göttingen, Grisebachstrasse 6, 37077 Göttingen, Germany
Eawag, Swiss Federal Institute of Aquatic Science and Technology, Überlandstrasse 133, 8600 Dübendorf, Switzerland
Institute of Integrative Biology, Department of Environmental Systems Science, ETH Zürich, 8092 Zürich, Switzerland
Author to whom correspondence should be addressed.
Insects 2022, 13(12), 1142;
Submission received: 22 November 2022 / Revised: 2 December 2022 / Accepted: 7 December 2022 / Published: 12 December 2022
(This article belongs to the Collection Biology and Management of Sap-Sucking Pests)



Simple Summary

Here, we review the current knowledge on the taxonomy, biodiversity, and ecology of cereal aphid parasitoids in Europe, which include 26 cereal aphid primary parasitoids and 28 hyperparasitoids. We present approaches to studying aphid–parasitoid–hyperparasitoid food webs, as well as the secondary endosymbionts in cereal aphids that may influence parasitoid community composition via their effects on food webs. We also review the effects of agricultural practices, environmental variation, and landscape complexity, on cereal aphid food webs and biological control.


Cereals are very common and widespread crops in Europe. Aphids are a diverse group of herbivorous pests on cereals and one of the most important limiting factors of cereal production. Here, we present an overview of knowledge about the taxonomy, biodiversity, and ecology of cereal aphid parasitoids in Europe, an important group of natural enemies contributing to cereal aphid control. We review the knowledge obtained from the integrative taxonomy of 26 cereal aphid primary parasitoid species, including two allochthonous species (Lysiphlebus testaceipes and Trioxys sunnysidensis) and two recently described species (Lipolexis labialis and Paralipsis brachycaudi). We further review 28 hyperparasitoid species belonging to three hymenopteran superfamilies and four families (Ceraphronoidea: Megaspillidae; Chalcidoidea: Pteromalidae, Encyrtidae; Cynipoidea: Figitidae). We also compile knowledge on the presence of secondary endosymbionts in cereal aphids, as these are expected to influence the community composition and biocontrol efficiency of cereal aphid parasitoids. To study aphid–parasitoid–hyperparasitoid food webs more effectively, we present two kinds of DNA-based approach: (i) diagnostic PCR (mainly multiplex PCR), and (ii) DNA sequence-based methods. Finally, we also review the effects of landscape complexity on the different trophic levels in the food webs of cereal aphids and their associated parasitoids, as well as the impacts of agricultural practices and environmental variation.

1. Introduction

Cereals dominate in the European agricultural landscape, by encompassing almost one-third of the European agricultural area [1]. It is well known that aphids are one of the most important limiting factors of cereal production, through direct damage or by transmitting viral pathogens (e.g., barley yellow dwarf virus, wheat dwarf virus) [2,3,4,5,6,7]. Aphid parasitoids are diverse and effective natural enemies of cereal aphids [8,9,10,11,12,13,14,15,16]. It should be noted that the high importance of aphidiines as natural enemies of cereal aphids led to their introduction into North and South America [17,18,19] and Australia [20,21] to control cereal aphids.
Brewer and Elliott [19] reported classical biological control efforts against cereal aphids in North America using parasitoids and predators. Cereal aphids and their parasitoids are common models for numerous ecological and evolutionary studies focusing on tri-trophic food web interactions (cereal aphid-parasitoid-hyperparasitoid) [13,22,23,24,25]; landscape effects on parasitoid species diversity [26] or biological control [13,27,28]; and the importance of agricultural practices on parasitoid effectiveness in cereal agroecosystems [24,29,30]. Most studies about cereal aphids and related parasitoids consider a few economically important species, usually at a local or regional scale [12,31,32,33,34]. Hyperparasitoids have the potential to compromise biological control by primary parasitoids [35,36]. The species composition of hyperparasitoids, as a top consumer in cereal aphid–primary parasitoid systems, has not been sufficiently explored. For instance, Tomanović et al. [12] reported the relative abundance of several hyperparasitoid species feeding on aphidiines attacking four major cereal aphid species in Serbia. Ye et al. [37] explored 16 species of hyperparasitoids from European cereal crops. Surprisingly, the basic biological and taxonomic characteristics of species participating in cereal aphid food webs are poorly known, including the cereal aphids’ endosymbionts, potentially affecting parasitoids and hyperparasitoids [38]. Starý [9] and Kavallieratos et al. [10] reviewed parasitoid associations and provided keys for the identification of cereal aphid parasitoids in the western Palaearctic and Southeastern Europe, respectively. Several studies have presented the knowledge to date about cereal aphids and their natural enemies in Europe [2,10,12,39,40]. However, there have been no updates on the biodiversity, taxonomy, and ecology of cereal aphid parasitoids and their endosymbionts. Therefore, the aim of the current review was to present knowledge about the taxonomy of cereal aphid parasitoids; species composition, including allochthonous parasitoids, food webs, and the effects of agricultural practices and landscapes on parasitoid diversity and efficiency; and the role of endosymbionts in biological control.

2. Cereal Aphids in Europe

Aphids are common pests in European cereal crops. Species from the genera Sitobion (Sitobion avenae (F.)), Metopolophium (Metopolophium dirhodum (Walker)), and Rhopalosiphum (Rhopalosiphum padi (L.), Rhopalosiphum maidis (Fitch)) are the most common on wheat, barley, maize, and oat crops in Europe [40,41]. In addition, climate change leads to the increasing pest importance of Schizaphis graminum (Rondani) in European cereal crops [42]. According to Blackman and Eastop [3], 34 aphid species are known to infest cereal crops in Europe. Some of those that are attacked by primary parasitoids and hyperparasitoids are mentioned in Table 1 and Table 2.

3. Primary Parasitoid Species Identification and Their Taxonomy

Identification is based on morphological characters. Some cereal aphid parasitoids are taxonomically well-defined and identifiable morphologically with existing keys for the identification of species [9,10,31,56]. Based on current records, 13 Aphidiinae genera include species that attack cereal aphids in European cereal agroecosystems.

3.1. Aphidius Nees

Across Europe, species from the genus Aphidius are the dominant parasitoids of different cereal aphids [10,32,33,57,58]. Among them, Aphidius rhopalosiphi De Stefani and Aphidius uzbekistanicus Luzhetzki are key species for the biological control of cereal aphids. They are also taxonomically the most problematic species [9,59,60]. According to Eady [61], both species belong to a group with a costulate anterolateral part of the petiole. Based on morphological characteristics, their identification often leads to misidentification. Several papers have considered the separation and characterization of both species [9,59,60,62], but due to their overlapping morphological characters, there is a list of synonyms of these species [9,56]. Summarizing all previous research efforts on the taxonomy A. rhopalosiphi and A. uzbekistanicus, the two species can be distinguished on the basis of the following characteristics: A. uzbekistanicus has a broad triangular forewing pterostigma, a narrow yellow ring at the base of flagellomere I, and usually fewer antennal segments (15–16); while A. rhopalosiphi has a more elongated pterostigma, a variable color pattern of flagellomere 1 and 2 (with predominantly yellow parts of flagellomere 1 and occasionally flagellomere 2), and usually 16–17 segmented antennae [60]. Aphidius matricariae Haliday represents an additional species with a costulate petiole and an elongated pterostigma, but it is clearly characterized by three-segmented maxillary palps and two-segmented labial palps, while those of A. rhopalosiphi and A. uzbekistanicus are four- and three-segmented, respectively [63,64]. Aphidius avenae Haliday and Aphidius colemani Viereck belong to the group with a costate anterolateral area of the petiole, but are clearly distinguished by shallow costae and the light-colored body of A. colemani, in contrast to the deep ridges on the anterolateral part of the petiole and a dark-colored body of A. avenae [63,65]. Additionally, the number of antennal segments in A. colemani is 14–15 vs. (16) 17–18 in A. avenae [11,63,66]. Although Aphidius picipes (Nees) is a commonly used name, it is a synonym of A. avenae. Aphidius ervi Haliday is an abundant cereal aphid parasitoid and the most important parasitoid of Acyrthosiphon pisum (Harris) in European legumes [67]. This is a large species, belonging to the group with a rugose anterolateral area of the petiole, and it is clearly separated from other congeners in cereal aphid parasitoid communities [10].

3.2. Praon Haliday

Four Praon species are known to be members of the cereal aphid parasitoid community in Europe. These species are divided into two groups, with either a developed forewing medio-cubital vein (m + cu) (Praon volucre (Haliday) and Praon abjectum (Haliday)) or with an effaced m + cu vein (Praon gallicum Starý and Praon necans Mackauer). Praon volucre is the most common Praon species in cereal agroecosystems in Europe and easily recognizable, with (16)17–18(19) segmented antennae, while P. abjectum has 14–15 segmented antennae. Praon gallicum is generally yellow to light brown, with yellow F1 and F2, while P. necans is generally darker with brown F1 and F2. For more details about morphological identification see Starý [9], Kavallieratos et al. [68] and Rakhshani et al. [69].

3.3. Binodoxys Mackauer

From this genus, only Binodoxys angelicae (Haliday) parasitizes cereal aphids in Europe. All species from this genus are characterized by possessing two prongs on the last abdominal sternite for host grasping, and two pairs of tubercles on the petiole [9]. For more details about the taxonomy of Binodoxys see Lazarević et al. [70].

3.4. Trioxys Haliday

Two Trioxys species are known to parasitize cereal aphids in Europe: Trioxys auctus (Haliday) and the North American species Trioxys sunnysidensis Fulbright and Pike. Both species share common characteristics, such as 12 segmented antennae, 4–6 setae on dorsal prongs, and two simple bristles on the top of prongs, but they are easily distinguished by the numerous longitudinal striations on the dorsal side of the petiole of T. auctus [71,72].

3.5. Lysiphlebus Förster

There are three Lysiphlebus species on cereal aphids in Europe (Lysiphlebus fabarum (Haliday), Lysiphlebus testaceipes (Cresson), and Lysiphlebus dissolutus (Nees) on root aphids—see below). Lysiphlebus fabarum is a native European species, characterized by a long forewing R1 vein (= metacarpus), thickened antennae, and setae on the forewing edge that are shorter than those on the forewing surface. This species occurs in both sexual and asexual populations [73]. The allochthonous L. testaceipes (Nearctic species) has a short R1 vein and long setae along the forewing edge.

3.6. Ephedrus Haliday

Ephedrus species have braconid-like wing venation patterns and 11 antennomeres in both sexes. Only Ephedrus plagiator (Nees) is a member of the cereal aphid parasitoid community in Europe, while Ephedrus persicae Froggatt attacks some aphids using cereals as secondary host plants, but only on their primary host plants [9]. Ephedrus plagiator has a longer three-SR vein than two-SR and a more elongated petiole than E. persicae [9,74,75].

3.7. Lipolexis Förster

Although Lipolexis gracilis Förster has been considered the only member of the genus Lipolexis attacking cereal aphids (R. padi) [9], Kocić et al. [46] recently identified an additional species, Lipolexis labialis Tomanović and Kocić, which parasitizes Anoecia corni (F.) on wheat. The genus is characterized by needle-like ovipositor sheaths and reduced wing venation. The two species can be separated from each other by the number of labial palpomeres (two in L. labialis and one in L. gracilis) and by a more elongated first flagellomere in L. labialis.

3.8. Additional Parasitoid Species in Cereal Fields across Europe

Diaeretiella rapae (M’Intosh) is an easily recognizable species with (13)14 segmented antennae, narrow areola on the dorsal side of propodeum, reduced wing venation pattern, and a R1 vein that is shorter than the pterostigma [10,76,77]. Monoctonus caricis (Haliday) is the only Monoctonus species parasitizing cereal aphids. It has ploughshare shaped ovipositor sheaths and 13 segmented filiform antennae [47]. Adialytus ambiguus (Haliday) is a specialized parasitoid of Sipha spp. aphids on cereals in Europe. Morphologically characterized by a reduced forewing venation with radial vein and R1 vein longer than the pterostigma. Its propodeum bears two divergent carinae at the base [43,78].
Toxares deltiger (Haliday) is the only known species of the genus Toxares in Europe, and it is characterized by a braconid-like wing venation pattern, deltoid shaped ovipositor sheaths and antennae with 18–20 antennomers [79].
A specific aphid parasitoid fauna is present on cereal root aphids, i.e., Aclitus obscuripennis Förster (marginal cell of the forewing closed or with narrow opening, 15 segmented antennae, ovipositor sheaths short and triangular), L. dissolutus (15–16-segmented antennae with subsquare flagellomeres and carinated propodeum), and Paralipsis enervis (Nees) (reduced wing venation with a very short triangular pterostigma without R1 vein [73,80,81]. These parasitoids exhibit very specific morphological peculiarities (small eyes, square antennal segments, short body, and strong legs), due to adaptation to their specific ecological niche.

4. Integrative Taxonomy

Although some species of cereal aphid parasitoids are morphologically well-defined, the identification and taxonomy of others are still very complicated and require an integrative approach. Over the last decade, researchers have incorporated molecular methods as additional tools, together with morphology and ecology, for the accurate identification of aphidiine wasps [82]. Cereal aphid parasitoids belonging to the genus Aphidius are among the first Aphidiinae whose molecular data (cytochrome c oxidase subunit I (COI) sequences) were used to determine their taxonomic status. Kos et al. [60] used morphological data (e.g., traditional and geometric morphometrics), molecular data (COI sequences), and ecological data (trophic relationships) to resolve the taxonomic status of A. uzbekistanicus, A. rhopalosiphi, and the American species A. avenaphis (Fitch). The authors found very distinct mitochondrial lineages within several A. rhopalosiphi populations, correlated with morphological variability, and indicating the existence of cryptic species within this taxon.
Derocles et al. [83] confirmed the utility of COI sequences as a genetic marker for Aphidiinae identification and also presented their limitations for sole use. After these studies, an integrative approach became the “gold standard” in Aphidiinae taxonomy [82], which resulted in numerous and interesting new findings in cereal aphid parasitoids. For example, by combining the mitochondrial COI (cytochrome c oxidase subunit I) barcoding gene and geometric morphometrics, Mitrovski-Bogdanović et al. [84] found that P. abjectum represents a complex of sibling species. It was determined that P. abjectum, similarly to wasps parasitizing Aphis sambuci L. on common elder (Sambucus nigra), a very common plant in seminatural habitats in cereal fields, represents a different species named Praon sambuci Tomanović and Starý. Besides genetic and ecological differences, P. sambuci has morphological traits that separate it from P. abjectum, such as a broader pterostigma. Furthermore, the use of an integrative approach in studies of cereal aphid parasitoids led to the discovery of previously undiscovered allochthonous species and of species that were new to science.

5. Allochthonous Parasitoid Species

Two allochthonous species of cereal aphid parasitoids, whose role in multitrophic food webs is still poorly known, appear in European cereal agroecosystems. Lysiphlebus testaceipes was introduced from Cuba to southern France in 1973, for the control of citrus aphids [85]. However, this parasitoid attacked numerous aphid hosts, including cereal aphids, in this new environment [10,34,44,86] and spread along the Mediterranean coast, also colonizing inland habitats [87,88,89]. Trioxys sunnysidensis has been described from the USA in association with R. padi infesting wheat near irrigation channels [71]. The authors found T. sunnysidensis in open wheat fields in Germany attacking R. padi [26,50]. Due to its morphological similarity with T. auctus and probably also B. angelicae, this species has been overlooked for a long time in European cereal agroecosystems; therefore, all former records need to be revisited for a thorough re-examination. Although we considered this species as allochthonous and probably unintentionally introduced from North America to Europe, our findings on haplotype diversity of T. sunnysidensis imply a possible European origin and Holarctic distribution [50].

6. New Aphid Parasitoids in European Cereal Agroecosystems

Two new species of cereal aphid parasitoids have been described in the last couple of years. The previously mentioned L. labialis was newly described as a parasitoid of A. corni and has been found in several European countries [46]. Paralipsis spp. parasitize root aphids [80,81]. Little is known about the economic effects of root aphids on cereals [90], even though they exhibit considerable diversity in European cereal fields [91]. Paralipsis enervis is a specialized parasitoid of root aphids that has been recorded on Geoica utricularia (Passerini) [44]. After careful examination of many Paralipsis specimens, a new sibling species was described from this genus, Paralipsis brachycaudi Tomanović and Starý, emerging from Tetraneura ulmi (L.) [48].

7. Species Complexes

Cryptic speciation in aphid parasitoids is a very common phenomenon. Especially within what had seemed to be oligophagous species exhibiting broad host ranges, some host associated lineages had to be newly described as cryptic species, after applying modern integrated taxonomic approaches [73,92]. Additionally, it seems that widely distributed generalist species often cover a narrower host range at a local scale, driven by trophic specialization and environmental factors in specific areas [93]. For example, the broadly oligophagous D. rapae attacks only a few aphid hosts in the continental part of Southeastern Europe, mainly Brevicoryne brassicae (L.) in the lowlands, and exceptionally Myzus persicae (Sulzer) under glasshouse conditions. Additionally, D. rapae parasitizes some specific non-crop aphids occurring in the mountains of Southeastern Europe (e.g., Aphis cadiva Walker, Hayhurstia atriplicis (L.), and Pseudobrevicoryne leclanti Petrović-Obradović and Remaudière) [44]. In contrast, the host range of this species in the Mediterranean is very diverse and composed of >20 aphid species, including some cereal aphids (Diuraphis noxia (Kurdjumov), R. padi, Rhopalosiphum maidis (Fitch)) [44,45]. A second example is A. avenae, which is more frequently recorded as a member of the cereal parasitoid spectrum in Central and Western Europe than in Southern Europe, where it is more abundant in high mountains as a parasitoid of some Sitobion aphid species [44,63]. Therefore, it has become evident that local specialization plays an important role, since generalist species behave as specialists at local scale in broader geographic areas [93]. Further research is needed to confirm the status of host-associated lineages in broadly oligophagous parasitoid species.
Several authors have considered the taxonomic status of A. rhopalosiphi and A. uzbekistanicus as two sibling species (“rhopalosiphi-uzbekistanicus” species complex). Apart from their overlapping morphological characters, they also share a similar host range pattern (Table 1). Kos et al. [60] found that A. uzbekistanicus shows very restricted variation, both genetically and morphologically. In A. rhopalosiphi, on the other hand, several mitochondrial lineages exist, forming a highly diverse haplotype network [60]. The status of this large number of haplotypes should be re-evaluated in future research. Electromorph variability, host range, and breeding experiments indicated A. ervi as a species complex consisting of several sibling taxa [94]. Until now, only Aphidius microlophii Pennacchio and Tremblay has been split from A. ervi and described as a specialized parasitoid of Microlophium carnosum (Buckton), associated with Urtica spp., which are common in cultivated areas [95,96]. Molecular markers did not support any significant differentiation within the “ervi” complex [97]. This issue should be further investigated with more robust taxonomic and geographic samples. Mitrovski-Bogdanović et al. [84] demonstrated that P. abjectum represents a species complex and described P. sambuci and Praon longicaudus Tomanović and Starý. In addition, further research on the various P. abjectum biotypes (e.g., of Aphis spp. hosts) should be conducted. Praon volucre is a broadly oligophagous species attacking a wide spectrum of aphid hosts [98] and also exhibiting possible biotype diversification [92]. However, a previous investigation did not reveal any specific genetic or morphological lineages within several P. volucre biotypes [99]. Rakhshani et al. [78] recognized by revealing morphological diversification within A. ambiguus,Adialytus cf. ambiguus” (Haliday) and “Adialytus arvicola” (Starý) emerging from Sipha aphid hosts. Both phenotypes required further taxonomic treatment.

8. Host Location, Specialization, and Exploitation of Aphid Colonies

Cereal aphids belong to several aphid tribes in Europe (i.e., Aphidini, Macrosiphini, Pterocommatini, Anoecini, Eriosomatini, Fordini) that are characterized by weak mobility, and feeding upon leaves, green stems, ears, or roots [100,101,102,103]. Aphid colonies are located on the open surface of cereals, except D. noxia which grows within curled leaves. All cereal aphids are medium-sized (except the small-sized D. noxia) and develop dense colonies. Most cereal aphids are monoecious and feed only on cereals and other Poaceae, except for M. dirhodum and R. padi, which are heteroecious. They migrate in autumn to primary woody host plants other than Poaceae, such as Rosa spp. and Prunus padus, respectively, where they mate and deposit their overwintering eggs [3]. It is well known that the most abundant and the most effective Aphidiinae parasitoids in European cereal agroecosystems are A. rhopalosiphi, A. uzbekistanicus, A. ervi, P. volucre, and E. plagiator [12,13,26,33,57,104], with the addition of A. matricariae, D. rapae, and allochtonous L. testaceipes in the Mediterranean area [10,34,44,45,105]. Since a measure of the efficiency of parasitoids in exploiting their hosts is their abundance on the hosts they attack [106,107], we can conclude that the most important cereal aphid parasitoid assemblages in Europe consist of oligophagous specialists (A. uzbekistanicus, A. rhopalosiphi, and A. ervi) and broadly oligophagous generalists (P. volucre and E. plagiator), with additional oligophagous generalists from the Mediterranean areas (A. matricariae, D. rapae, and L. testaceipes). According to Straub et al. [108], there are two hypotheses explaining the relationships between host-range breadth and the abundance of parasitoids. According to the resource–breadth hypothesis, generalist parasitoids are more abundant than specialists on common hosts, exhibiting an advantage under environmental instability (e.g., agroecosystems, urbanization) and low available resources. The trade-off hypothesis gives more advantages to specialists under a more stable environment [109]. In the case of cereal aphid parasitoids, a mixed complex of parasitoids (oligophagous specialists and oligophagous generalists) could provide adequate biological control and ecosystem stability through control of aphid populations [110].
According to van Baaren et al. [111,112], the cereal aphid parasitoid A. rhopalosiphi attacks S. avenae, as the most abundant parasitoid species present in the field prior to the arrival of A. ervi and A. avenae, by exploiting aphid colonies when they start to produce cornicle secretions. Upon arrival, A. ervi and A. avenae parasitize those aphid colonies that have not been exploited by A. rhopalosiphi.
There has been little research about the ability of cereal parasitoids to distinguish between parasitized and non-parasitized aphid hosts, an issue that is extremely important for the aphid host selection in parasitoids. Recognition of already parasitized aphid hosts by parasitoids is done using chemical cues in their cornicle secretion and exuviae, in combination with visual cues [113,114,115]. Outreman et al. [116] reported that A. rhopalosiphi recognizes already parasitized aphid hosts using external factors such as aphid alarm pheromone in cornicle secretion or by internal factors after the insertion of its ovipositor into the aphid’s body. However, these host discriminations are species-specific [117,118]. Van Baaren et al. [112] showed that A. rhopalosiphi recognizes S. avenae parasitized by A. ervi on the basis of cornicle secretion, which is released after parasitization. However, aphid hosts already parasitized by A. avenae do not alert others in the aphid colony [112]. The exploitation of aphid colonies by aphid parasitoids is mediated by many different factors, including host quality and the presence of different host defenses (e.g., mobility, wax production, and microorganisms) [119,120,121].

9. Primary Parasitoids and their Hyperparasitoids

Secondary parasitoids represent the fourth trophic level in multitrophic aphid–parasitoid communities and have the potential to exert top-down control in such food webs [122,123,124]. According to Sullivan and Völkl [123], there are several categories and subcategories of secondary parasitoids, of which two are relevant for aphidiines. “True” hyperparasitoids attack primary parasitoids at the larval stage within the living aphid and develop inside the primary parasitoid host, without arresting its development; hence, they are referred to as koinobiont endohyperparasitoids. Idiobiont ectohyperparasitoids, also referred to as mummy parasitoids, attack primary parasitoid prepupae and pupae, on which they develop externally within the empty aphid cuticule (the mummy). In Table 2, we review 28 hyperparasitoid species belonging to three hymenopteran superfamilies and four families (Ceraphronoidea: Megaspillidae; Chalcidoidea: Pteromalidae, Encyrtidae; Cynipoidea: Figitidae).
Generally, idiobiont ectohyperparasitoids on cereals have a broad host range, while koinobiont endohyperparasitoids exhibit host specificity, except for Syrphophagus aphidivorus (Mayr), Alloxysta brevis (Thomson), Alloxysta victrix Westwood, and Phaenoglyphis villosa (Hartig) (Table 2). However, cryptic speciation abounds in these wasps, especially in Alloxysta spp. [125], and field records of their aphid-primary parasitoid hosts are usually missing, such that the true extent of their host specificity remains elusive.
Hyperparasitoids disrupt biological control directly, by affecting primary parasitoids at variable levels of specialization [36,53,54,126] or as intraguild competitors (facultative hyperparasitoids) [127]. However, there is a huge debate about the role of hyperparasitoids as a fourth trophic level in the entire trophic systems. Some authors point out the positive role of hyperparasitoids in stabilizing the whole trophic chain through density-dependent interactions [126,128,129]. Due to higher thermal requirements and the time-lag inherent in consumer–resource relationships, hyperparasitoids appear later in the season in comparison with aphids and their primary parasitoids [36,122]. However, due to climate change and land-use changes through landscape simplification, higher temperatures could increase and positively affect the hyperparasitoid community, leading to pest increase [130]. Additionally, the increasing temperature causes primary parasitoids and hyperparasitoids to lose their diapause, an issue that negatively affects the abundance of primary parasitoids in the spring, due to the winter activities of hyperparasitoids [131,132]. However, the effects of climate change on the whole cereal aphid–parasitoid trophic system are still unpredictable and require further research.

10. Methods for Determining Trophic Interactions of Cereal Aphids and their Associated Parasitoids

Traditionally, the methods for determining trophic interactions among cereal aphids and their parasitoids have mainly included morphological approaches such as dissection and rearing. Direct host dissection provides an unequivocal estimate of parasitism rate, as well as multiparasitism, superparasitism, and hyperparasitism [55,133,134,135]. However, identifying the eggs or larvae of parasitoids inside their hosts is difficult and often impossible, due to a lack of distinguishable morphological characteristics [136,137]. On the other hand, the primary parasitoid and hyperparasitoid adults reared from field-collected hosts can be morphologically identified at the species level and linked directly with the aphid species they emerge from [138]. To link the hyperparasitoids to their real host (i.e., primary parasitoids), it is possible to identify the hyperparasitoid species and determine their host (i.e., primary parasitoids) at the genus level based on the morphological characteristics of aphid mummies [124,139]. Nevertheless, species-specific links between primary parasitoids and hyperparasitoids still remain obscure [140,141]. Additionally, rearing approaches are affected by delayed parasitoid emergence, as well as host and parasitoid mortality. There are also hurdles when facing multiparasitism and superparasitism [136,137].
Molecular approaches, mainly DNA-based approaches, can overcome these limitations [137]. They have been shown to allow for species-specific examination of trophic interactions between primary parasitoids (e.g., multiparasitism), as well as between primary and secondary parasitoids (e.g., hyperparasitism) [22,24,142]. These approaches remain unaffected by delayed parasitoid emergence, as well as host and parasitoid mortality, and can be applied to each developmental stage [136]. To date, there have been two kinds of DNA-based approaches used to determine aphid–parasitoid–hyperparasitoid food webs: (i) diagnostic PCR methods (mainly multiplex PCR), and (ii) DNA sequence-based methods [137]. Multiplex PCR allows the amplification of several targets in parallel within a single reaction [22,25,142,143], compared to singleplex PCR [144]. However, diagnostic PCR approaches require prior knowledge of all potential species in the study area [145,146], and might be affected by unexpected trophic interactions, such as alternative food sources and invasive species [147,148]. DNA sequence-based methods, on the other hand, can overcome these limitations and have been used to identify parasitoid DNA using universal primers, followed by Sanger sequencing [83] or next-generation sequencing (NGS) [149]. The issue with Sanger sequencing is that, when there are multiple species in the same DNA extract, it either obtains one DNA sequence or even sequencing failure, as in the case of hyperparasitim and multiparasitim [83]. The NGS approach can overcome this hurdle [150], but it is affected by different target DNA proportions among test samples [145]. Compared to rearing and morphological identification of parasitoids, another limitation of molecular approaches is the difficulty in estimating the superparasitism and real biocontrol efficiency of parasitoids, as they can detect the presence of parasitoid eggs and larvae within the host even when they would not have survived to complete their development [52,135,151].

11. Cereal Aphid–Primary Parasitoid–Hyperparasitoid Food Webs

Host–parasitoid food webs are among the most studied trophic interactions. They have been investigated as models to address important ecological questions, such as robustness and restoration of ecological networks [152], bottom-up effects and habitat effects on food web structures [153,154], as well as apparent competition [155,156].
The cereal aphid–primary parasitoid–hyperparasitoid food webs across different habitats have been broadly surveyed and analyzed [157,158]. These networks have also been investigated to understand important questions in trophic ecology, such as parasitoid host specificity [100,159], intraguild predation [160,161], and the effects of climate change on trophic networks [162,163,164]. Further studies were concerned with how farming practices affect these trophic interactions, including the effects of agriculture intensification [23,24,124,165], landscape changes [139,166], habitat modification [167,168], and pesticide application [169]. To date, several cereal aphid–primary parasitoid–hyperparasitoid food webs have been reconstructed in different geographic regions around the world [22,143], exhibiting partial spatiotemporal variation [159].
Aphid–primary parasitoid–hyperparasitoid networks are linked to other trophic levels, such as plants, plant pathogens, and predators, forming wider and much more complex trophic interaction networks. Plants can have cascading effects on aphid–primary parasitoid–hyperparasitoid food webs. For example, negative impacts of aphid resistant soybeans on the development, mummification, and adult parasitoid emergence have been observed [170]. Among plant-derived defense chemicals, beta-aminobutyric acid has a negative impact on the size of emerging A. ervi [171]. On the other hand, acetylsalicylic acid, oxalic acid, and glucosinolates can positively affect aphid parasitoids [172,173]. Genetically-modified crops seem not to have strong effects on the non-target pests, aphids and their associated parasitoids. For cereal, the impact of powdery mildew-resistant GM wheat on cereal aphid–parasitoid–hyperparasitoid food webs seems negligible and limited [174]. Moreover, in other crop systems, transgenic Cry1Ac + CpTI cotton cultivars do not affect aphid–parasitoid food webs and biological control [142]. Although transgenic potatoes with nematode resistance can reduce the density of aphids, no effect on higher trophic levels (i.e., primary- and hyperparasitoid) has been found [175].
Besides plant traits, plant-associated symbionts or pathogens can affect aphid–primary parasitoid–hyperparasitoid food webs. Plant protective symbiotic fungal infection can depress both aphids and parasitoids [176,177,178,179,180]. Arbuscular mycorrhizal fungi of plants have a negative impact on the cereal aphid R. padi, but positive effects on related plants and parasitoids [181]. Plant virus infection, on the other hand, can increase aphid size [182], as well as parasitoid larvae mortality and developmental times [183].
Intraguild predation by generalist predators has been found to disrupt the cereal aphid control exerted by primary parasitoids [160]. In contrast, other studies also found intraguild predation of predators on parasitoids in cotton aphids and soybean aphids, but in this case the intraguild predation seemed not to disrupt the aphid biological control [184,185]. Additionally, parasitoids can respond to intraguild predation by avoiding chemical traces of predators [161]. Furthermore, the mutualism of aphids with tending ants has been found to affect aphid–primary parasitoid–hyperparasitoid food webs, mainly by reducing the generalist hyperparasitoids [186].

12. The Role of Secondary Endosymbionts

In addition to the factors discussed above, the presence of secondary endosymbionts in their hosts is expected to influence the community composition and biocontrol efficiency of cereal aphid parasitoids. Secondary or facultative endosymbionts of aphids are maternally transmitted bacteria that, in contrast to the primary or obligate endosymbiont Buchnera aphidicola Munson et al. 1991 (Enterobacterales: Erwiniaceae), are not strictly required for host survival. It was first demonstrated by Oliver et al. [187], in pea aphids, that two secondary endosymbionts, Hamiltonella defensa Moran et al. 2005 (Enterobacterales: Enterobacteriaceae) and Serratia symbiotica Moran et al. 2005 (Enterobacterales: Yersiniaceae) [188], increased resistance to the parasitoid A. ervi. Further research has shown that endosymbionts conferring resistance to parasitoids are widespread in many species of aphids [189,190,191,192,193] and that “defensive” symbionts may also provide protection against natural enemies other than parasitoids, such as entomopathogenic fungi [194,195,196]. The secondary endosymbionts of cereal aphids have been increasingly surveyed over recent years, and the knowledge available so far suggests that the “usual suspects” [197,198] are present. H. defensa, S. symbiotica, Regiella insecticola Moran et al. 2005 and Rickettsia sp. have been reported from S. avenae [25,199,200,201,202,203]. Interestingly, in addition to H. defensa, R. insecticola, and Rickettsia sp., Li et al. [204,205,206] detected and described a previously unknown secondary endosymbiont (SMLS) (Rickettsiales: Rickettsiaceae) in Sitobion miscanthi (Takahashi) (Hemiptera: Aphididae) [204,205,206], which was also found in R. padi [204]. Additional secondary symbionts reported from R. padi and R. maidis include S. symbiotica, H. defensa, R. insecticola, Rickettsia sp., Spiroplasma sp., Arsenophonus sp., and Wolbachia sp. [207,208]. Telesnicki et al. [209] found a high frequency of infection with H. defensa in Argentinean M. dirhodum, which was mainly a host for R. insecticola in Chile [210].
Considering that H. defensa is associated with resistance to parasitoids in several aphid species [211], it was surprising that four strains of H. defensa from S. avenae did not provide any protection against the parasitoids A. ervi and E. plagiator [199]. However, it is known that different strains of H. defensa vary in the strength of resistance they provide [212], that protection can be specific against certain parasitoid species or even certain genotypes of the same parasitoid [192,213,214,215,216,217], and that parasitoids can evolve counteradaptations to overcome symbiont-conferred resistance [218,219,220]. It would thus be premature to assume a general absence of H. defensa-conferred protection in S. avenae or other cereal aphids based on one negative experimental result. Indeed, some evidence for H. defensa-mediated resistance to cereal aphid parasitoids has since been accumulated [25,193], but it is by no means universal [221,222].
Could hyperparasitoids be affected by aphids’ endosymbionts as well? This has never been tested in cereal aphids, but it seems unlikely, because in symbiont-protected aphids, the development of primary parasitoids is often arrested very early [223]. On the other hand, defensive symbionts have the potential for upward cascading effects to the trophic level of hyperparasitoids, by reducing the abundance of primary parasitoids in the food web [224]. This can alter food web composition. A field experiment with a non-cereal aphid, Aphis fabae, not only showed a strong reduction in parasitism in Hamiltonella-protected aphids, but also showed a dramatic change in the community composition of primary and secondary parasitoids, because some primary parasitoids were virtually excluded from plots with aphids harboring H. defensa. It will, thus, be important to consider the secondary endosymbionts harbored by cereal aphids when studying the parasitoid communities they support [25].

13. Effect of Agricultural Practices and Environment

The abundance of cereal aphids in field crops is affected by biotic and abiotic conditions that are linked to the environment and the agricultural practices followed. Temperature and precipitation affect the abundance of certain cereal aphid species. For example, the population growth of the grain aphid S. avenae increased when the mean fall temperature increased, but when the mean spring temperature and precipitation increased, populations decreased [225]. Similarly, wheat aphid, S. graminum populations decreased when the mean winter temperature and total fall precipitation increased [225]. Furthermore, Thierry et al. [226] reported that high temperatures during spring and winter caused a reduction of the bird cherry-oat aphid R. padi numbers, whereas high temperatures during fall increased the abundance of R. padi. Increases of wind speed, relative humidity, and precipitation decreased the abundance of cereal aphids caught in suction traps [227]. In a recent study, high levels of CO2, which was used as a fertilizer, and high temperatures decreased R. padi development and fecundity [228]. Soil fertilization with inorganic fertilizer accelerated the development and increased the fecundity of S. avenae, when the infested wheat plants were sufficiently watered [229]. Furthermore, 0.4 NH4NO3 g/plant of four different types of nitrogen fertilizers, enhanced the fecundity and the weight of adults of R. padi and S. avenae in contrast with 0.1 or 0.2 NH4NO3 g/plant [230]. The population densities of R. padi and M. dirhodum were higher when the fertilization with nitrogen was high (80 kg N/ha as manure vs. 80 kg N/ha as manure + 110 kg N/ha as mineral fertilizer) [231]. In some cases, nitrogen was not a favorable fertilizer for cereal aphid growth, in contrast to phosphorus, which enhanced the aphid infestation [232]. In addition, a 69-46-25 kg/acre N-P-K fertilization reduced cereal aphid populations, while simultaneously increasing the yield of the crop [233]. Nitrogen fertilization with maize straw amendment caused a reduction in the number of the aphids [234]. Interestingly, fertilization can alter certain aphid characteristics. For example, nitrogen fertilization caused R. padi and S. avenae individuals to gain body weight [235].
Since cereal aphid populations can be controlled by beneficial organisms, their impact is highly affected by environmental conditions [236,237,238,239]. For instance, nitrogen fertilizer led to an increase of parasitism of A. colemani and A. rhopalosiphi on S. avenae and R. padi, respectively [239]. Furthermore, the abundance of S. avenae, S. graminum, and R. padi, as well as the parasitoid species A. avenae, Praon rhopalosiphum Takada, and Aphelinus albipodus Hayat and Fatima (Hymenoptera: Aphelinidae), were increased by landscape complexity [237]. Similarly, Plećaš et al. [13] documented that large non-crop habitats, with high landscape heterogeneity, increased the population of cereal aphids, their parasitoids, and hyperparasitoids. Winqvist et al. [240] documented that landscape homogeneity resulted in less predation on cereal aphids. The authors also found that predation in non-organic (conventional) fields was higher than in organic fields within simple landscapes. In a recent study, the abundance of S. miscanthi and R. padi was decreased when five different varieties of winter wheat were sown in a field [241]. Different wheat genotypes can also determine aphid abundance. Batyrshina et al. [242] reported that the tetraploid wild emmer Triticum turgidum ssp. dicoccoides cv. Zavitan was the most resistant among three varieties, while the hexaploid spring bread Triticum aestivum cultivar Rotem was the most susceptible. The authors documented that aphids did not colonize juvenile plant tissues, preferring the matured spikes and flag leaves; therefore, the aphid population was higher on plants that matured earlier. R. padi heavily colonized maize but not grasslands that surrounded wheat seedlings, indicating that neighboring landscapes aid or deteriorate aphid infestation [243]. However, not all neighboring plants are useful, since certain species enhance the presence of cereal aphids. For instance, flower strips in-between fields increased the abundance of cereal aphids [244]. However, wildflower strips enhanced aphid parasitism and predation [245].
Pesticide applications are of major importance, since their irrational use may cause the evolution of resistance in exposed aphids [246,247,248]. Due to the continuous use of pyrethroids [247,248], some S. avenae strains exhibit the L1014F (kdr) mutation that makes them resistant to these insecticides [249]. Similarly, Wang et al. [247] documented that a strain of R. padi resistant to lambda-cyhalothrin exhibited cross-resistance to another nine active compounds. On the other hand, some insecticides, such as imidacloprid, controlled R. padi, S. graminum, and S. avenae aphids, resulting in a yield enhancement [250]. The application of imidacloprid should be handled cautiously, since it does not only affect cereal aphids, but their parasitoids as well [251]. Furthermore, insecticides did not reduce aphid densities in the presence of their natural enemies, while in their absence, insecticides were effective [252]. Apart from aphids, insecticides also kill the aphids’ natural enemies, and therefore aphid populations resurge more quickly after insecticidal treatments [252]. The year of insecticidal treatments should also be taken into consideration, since insecticides can be more or less efficient due to the prevalent environmental conditions. For instance, an experiment that took place during 2008–2009 revealed that cereal aphid densities were not reduced by treatment with the pyrethroid insecticide lambda-cyhalothrin. However, when the same experiment was repeated during 2010–2011, the same insecticide killed the exposed aphids [253]. This fact could be attributed to the high precipitation levels occurring in 2008–2009 vs. 2010–2011 [253]. The efficacy of insecticides may differ among strains of the same aphid species. For instance, Zuo et al. [254] reported that twelve geographical strains of R. padi exhibited variable levels of susceptibility or resistance to ten insecticidal active ingredients. Recently, Cao et al. [255] documented that short exposures of S. avenae to higher temperatures altered the tolerance of this species to imidacloprid.
Overall, the effects of the environment and agricultural practices are multifarious and can strongly affect the biological traits of aphids and their associated natural enemies in cereal fields, consequently increasing or decreasing the yield of the infested crops. These parameters should be taken into account when insecticidal management strategies are employed against cereal aphids. The use of registered insecticides at label doses and the rotation of the active ingredients can be suggested as management strategies, to ensure delayed development of the insecticidal resistance of cereal aphids [247,254].

14. Landscape Complexity

As already mentioned above, the structure of a whole landscape can influence aphids and their parasitoids. A structurally complex landscape can improve cereal aphid abundance and the primary parasitism rate, by providing more overwintering sites, alternative hosts, food sources, and shelter after harvest [27,256]. Such positive effects were found in several studies [13,237], while no effect of landscape complexity was shown in other studies [24,166,257]. Additionally, such impacts of landscape structure can vary during the seasons [256]. Differences in the species diversity of primary parasitoids in simple and complex structured landscapes were observed [13] or not [26,138,139]. Higher diversity of cereal aphid parasitoids may lead to higher biological control, as different species invade fields at different times during the season to parasitize aphids (cp. section “Host location, specialization, and exploitation of aphid colonies” in this review).
Hyperparasitoids can also be affected by landscape structure, even more strongly than primary parasitoids [138,237,258]. Furthermore, it was shown that more hyperparasitoid species hatched from both stinging nettle aphids on the field margin and cereal aphids in the adjacent field than primary parasitoid species [138]. Derocles et al. [157] also found that aphids inside and outside of agricultural fields shared almost no primary parasitoid species. Therefore, hyperparasitoids can link different habitats and may gain more advantages from the provision of semi-natural habitats in agricultural landscapes than primary parasitoids [138], which may increase the effect of hyperparasitoids on primary parasitoids, due to hyperparasitoid individuals invading fields from surrounding semi-natural habitats.
Landscape complexity may also affect cereal aphid–primary parasitoid and primary parasitoid–hyperparasitoid food webs, as shown, for example, by the reduced complexity of such food webs in structurally rich landscapes compared to simple ones [24,139]. This shows that the effect of landscape structure can be variable and different at different trophic levels. This has to be taken into account when landscape structure is changed (e.g., by sowing flowering strips adjacent to fields) to enhance biological control of cereal aphids.

15. Conclusions

Aphids are one of the most important limiting factors of cereal production, by causing direct damage or by transmitting viral pathogens to cereal crops. Aphid parasitoids are diverse and effective natural enemies of cereal aphids. Surprisingly, the basic biological and taxonomic characteristics of species participating in the cereal aphid food webs are poorly known, including cereal aphids endosymbionts that potentially affect parasitoids and hyperparasitoids. Most existing studies have considered a few economically important aphid species (e.g., S. avenae, R. padi, and M. dirhodum) and a few parasitoid species (e.g., A. ervi, A. rhopalosiphi, and P. volucre), not considering the full complexity of cereal aphid food webs, including hyperparasitoids and endosymbionts. Predicting the stability and dynamics of cereal aphid food webs requires understanding the food web diversity of all belonging trophic members and their interactions. One of the most problematic issues for parasitoid food web research is the common phenomenon of cryptic speciation and the ensuing problems for taxonomy. The community composition of secondary endosymbionts and their role in the biocontrol efficiency of cereal aphid parasitoids are further important issues to consider in biocontrol strategies. Cereal aphid food webs are also contingent on the surrounding habitats and landscape complexity, which can also affect cereal aphids and their associated parasitoids in very different ways. Finally, a proper combination of insecticidal treatments with agricultural practices suited for the prevailing environmental conditions can lead to the optimization of cereal aphid management with minimum detrimental impacts on natural enemies and the environment. Although we tried to integrate fundamental research in the ecology, taxonomy, and biodiversity of cereal aphid food webs and applied research about the effective control of cereal aphid pests, we are aware that this approach needs further justification from future research, especially in the context of climate change.

Author Contributions

Conceptualization, Ž.T., N.G.K., Z.Y., E.P.N., A.P., I.M.G.V. and C.V.; Methodology, Ž.T., N.G.K., Z.Y., E.P.N., A.P., I.M.G.V. and C.V.; Validation, Ž.T., N.G.K., Z.Y., E.P.N., A.P., I.M.G.V. and C.V.; Investigation, Ž.T., N.G.K., Z.Y., E.P.N., A.P., I.M.G.V. and C.V.; Resources, Ž.T. and N.G.K.; Data Curation, Ž.T., N.G.K., Z.Y., E.P.N., A.P., I.M.G.V. and C.V.; Writing—Original Draft Preparation, Ž.T., N.G.K., Z.Y., E.P.N., A.P. and C.V.; Writing—Review and Editing, Ž.T., N.G.K., Z.Y., E.P.N., A.P., I.M.G.V. and C.V.; Visualization, Ž.T., N.G.K., Z.Y., E.P.N., A.P., I.M.G.V. and C.V.; Supervision, Ž.T.; Project Administration, Ž.T., N.G.K., Z.Y., E.P.N., A.P. and C.V.; Funding Acquisition, Ž.T. and Z.Y. All authors have read and agreed to the published version of the manuscript.


The research was partially funded by the Serbian Ministry of Science and Education (451-03-68/2022-14/200178), Grant Serbian Academy of Sciences F131 (Ž.T. and A.P.), and by Central Public-Interest Scientific Institution Basal Research Fund for the Chinese Academy of Tropical Agricultural Sciences (NO. 1630042021014, 1630042022006 and 1630042020002) (Z.Y.).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Data are contained within the article.

Conflicts of Interest

The authors declare no conflict of interest.


  1. European Commission. EU Cereal Farms Report Based on 2017 FADN Data European Union; Agriculture and Rural Development, European Union: Brussels, Belgium, 2019. [Google Scholar]
  2. Dixon, A.F.G. Cereal aphids as an applied problem. Agric. Zool. Rev. 1987, 2, 1–57. [Google Scholar]
  3. Blackman, R.L.; Eastop, V.E. Aphids on the World’s Crops. An Identification and Information Guide; John Wiley and Sons Ltd.: Chichester, UK, 2000. [Google Scholar]
  4. Van Emden, H.F.; Harrington, R. Aphids as Crop Pests; CABI Publishing: Wallingford, UK, 2017. [Google Scholar]
  5. Liu, Y.; Khine, M.O.; Zhang, P.; Fu, Y.; Wang, X. Incidence and distribution of insect-transmitted cereal viruses in wheat in China from 2007 to 2019. Plant Dis. 2020, 104, 1407–1414. [Google Scholar] [CrossRef] [PubMed]
  6. Brewer, M.J.; Peairs, F.B.; Elliott, N.C. Invasive cereal aphids of North America: Ecology and pest management. Annu. Rev. Entomol. 2019, 64, 73–93. [Google Scholar] [CrossRef]
  7. Stell, E.; Meiss, H.; Lasserre-Joulin, F.; Therond, O. Towards predictions of interaction dynamics between cereal aphids and their natural enemies: A review. Insects 2022, 13, 479. [Google Scholar] [CrossRef] [PubMed]
  8. Starý, P. Aphidius uzbekistanicus Luzhetzhi (Hym.: Aphidiidae), a parasite of graminicolous pest aphids. Annot. Zool. Bot. 1972, 85, 1–7. [Google Scholar]
  9. Starý, P. Biosystematic synopsis of parasitoids on cereal aphids in the western Palaearctic (Hymenoptera: Aphidiidae, Homoptera: Aphidoidea). Acta Entomol. Bohemoslov. 1981, 78, 382–396. [Google Scholar]
  10. Kavallieratos, N.G.; Tomanović, Ž.; Athanassiou, C.G.; Starý, P.; Žikić, V.; Sarlis, G.P.; Fasseas, C. Aphid parasitoids infesting cotton, citrus, tobacco, and cereal crops in southeastern Europe: Aphid-plant associations and keys. Can. Entomol. 2005, 137, 516–531. [Google Scholar] [CrossRef]
  11. Rakhshani, E.; Talebi, A.A.; Starý, P.; Tomanović, Ž.; Kavallieratos, N.G.; Manzari, S. A review of Aphidius Nees (Hymenoptera: Braconidae: Aphidiinae) in Iran: Host associations, distribution and taxonomic notes. Zootaxa 2008, 1767, 37–54. [Google Scholar] [CrossRef]
  12. Tomanović, Ž.; Kavallieratos, N.G.; Starý, P.; Petrović-Obradović, O.; Athanassiou, C.G.; Stanisavljević, L. Cereal aphids (Hemiptera: Aphidoidea) in Serbia: Seasonal dynamics and natural enemies. Eur. J. Entomol. 2008, 105, 495–501. [Google Scholar] [CrossRef] [Green Version]
  13. Plećaš, M.; Gagić, V.; Janković, M.; Petrović-Obradović, O.; Kavallieratos, N.G.; Tomanović, Ž.; Thies, C.; Tscharntke, T.; Ćetković, A. Landscape composition and configuration influence cereal aphid parasitoid hyperparasitoid interactions and biological control differentially across years. Agric. Ecosyst. Environ. 2014, 183, 1–10. [Google Scholar] [CrossRef]
  14. Alvarez-Baca, J.K.; Alfaro-Tapia, A.; Lavandero, B.; Le Lann, C.; Van Baaren, J. Suitability and profitability of a cereal aphid for the parasitoid Aphidius platensis in the context of conservation biological control of Myzus persicae in Orchards. Insects 2020, 11, 381. [Google Scholar] [CrossRef] [PubMed]
  15. Dos Santos, C.D.R.; Lau, D.; Redaelli, L.R.; Jahnke, S.M.; Pereira, P.R.; Engel, E.; Sampaio, M.V. Aphid-parasitoids trophic relationship in a cereal crop succession system: Population oscillation and food webs. Agric. Forest Entomol. 2022, 24, 516–530. [Google Scholar] [CrossRef]
  16. Woolley, V.C.; Yolice, L.B.; Tembo, B.N.; Obanyi, J.N.; Arnold, S.E.J.; Belmain, S.R.; Ndakidemi, P.A.; Joshua, O.O.; Stevenson, P.C. The diversity of aphid parasitoids in East Africa and implications for biological control. Pest Manag. Sci. 2022, 78, 1109–1116. [Google Scholar] [CrossRef] [PubMed]
  17. Zuniga, E. Biological control of cereal aphids in the Southern Cone of South America. In Proceedings of the International Workshop World Perspectives on Barley Yellow Dwarf, Udine, Italy, 6–11 July 1987; Burnett, P.A., Ed.; DCAS/CIMMYT: Mexico City, Mexico, 1990; pp. 362–367. [Google Scholar]
  18. Altieri, M.A.; Trujillo, J.; Campos, L.; Klein-Koch, C.; Gold, C.S.; Quezada, J.R. El control biologico clasico en America Latina en su contexto historico. Manejo Int. Plagas 1989, 12, 82–107. [Google Scholar]
  19. Brewer, M.J.; Elliott, N.C. Biological control of cereal aphids in North America and mediating effects of host plant and habitat manipulations. Annu. Rev. Entomol. 2004, 49, 219–242. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  20. Waterhouse, D.F.; Sands, D.P.A. Classical Biological Control of Arthropods in Australia; Australian Centre for International Agricultural Research: Canberra, Australia, 2001. [Google Scholar]
  21. Heddle, T.; Van Helden, M.; Nash, M.; Muirhead, K. Parasitoid communities and interactions with Diuraphis noxia in Australian cereal production systems. BioControl 2020, 65, 571–582. [Google Scholar] [CrossRef]
  22. Traugott, M.; Bell, J.R.; Broad, G.R.; Powell, W.; Van Veen, F.J.F.; Vollhardt, I.M.G.; Symondson, W.O.C. Endoparasitism in cereal aphids: Molecular analysis of a whole parasitoid community. Mol. Ecol. 2008, 17, 3928–3938. [Google Scholar] [CrossRef] [PubMed]
  23. Gagić, V.; Hanke, S.; Thies, C.; Scherber, C.; Tomanović, Ž.; Tscharntke, T. Agricultural intensification and cereal aphid-parasitoid-hyperparasitoid food webs: Network complexity, temporal variability and parasitism rates. Oecologia 2012, 170, 1099–1109. [Google Scholar] [CrossRef] [Green Version]
  24. Vollhardt, I.M.G.; Ye, Z.; Parth, N.; Rubbmark, O.; Fründ, J.; Traugott, M. Influence of plant fertilisation on cereal aphid-primary parasitoid-secondary parasitoid networks in simple and complex landscapes. Agric. Ecosyst. Environ. 2019, 281, 47–55. [Google Scholar] [CrossRef]
  25. Ye, Z.; Vollhardt, I.M.G.; Girtler, S.; Wallinger, C.; Tomanovic, Z.; Traugott, M. An effective molecular approach for assessing cereal aphid-parasitoid-endosymbiont networks. Sci. Rep. 2017, 7, 3138. [Google Scholar] [CrossRef] [Green Version]
  26. Vollhardt, I.M.G.; Tscharntke, T.; Wäckers, F.L.; Bianchi, F.J.A.; Thies, C. Diversity of cereal aphid parasitoids in simple and complex landscapes. Agric. Ecosyst. Environ. 2008, 126, 289–292. [Google Scholar] [CrossRef]
  27. Thies, C.; Roschewitz, I.; Tscharntke, T. The landscape context of cereal aphid–parasitoid interactions. Proc. Royal Soc. B 2005, 272, 203–210. [Google Scholar] [CrossRef] [Green Version]
  28. Bosem Baillod, A.; Tscharntke, T.; Clough, Y.; Batáry, P. Landscape-scale interactions of spatial and temporal cropland heterogeneity drive biological control of cereal aphids. J. Appl. Ecol. 2017, 54, 1804–1813. [Google Scholar] [CrossRef] [Green Version]
  29. Thies, C.; Haenke, S.; Scherber, C.; Bengtsson, J.; Bommarco, R.; Clement, L.W.; Ceryngier, P.; Dennis, C.; Emmerson, M.; Gagić, V.; et al. The relationship between agricultural intensification and biological control: Experimental tests across Europe. Ecol. App. 2011, 21, 2187–2196. [Google Scholar] [CrossRef] [PubMed]
  30. Altieri, M.; Nicholls, C. Biodiversity and Pest Management in Agroecosystems; CRC Press: Boca Raton, FL, USA, 2018. [Google Scholar]
  31. Powell, W. The identification of hymenopterous parasitoids attacking cereal aphids in Britain. Syst. Entomol. 1982, 7, 465–473. [Google Scholar] [CrossRef]
  32. Pankanin-Franczyk, M.; Ceryngier, P. Cereal aphids, their parasitoids and coccinellids on oats in central Poland. J. Appl. Entomol. 1995, 119, 107–111. [Google Scholar] [CrossRef]
  33. Sigsgaard, L. A survey of aphids and aphid parasitoids in cereal fields in Denmark, and the parasitoids’ role in biological control. J. Appl. Entomol. 2002, 126, 101–107. [Google Scholar] [CrossRef]
  34. Lumbierres, B.; Starý, P.; Pons, X. Seasonal parasitism of cereal aphids in a Mediterranean arable crop system. J. Pest Sci. 2007, 80, 125–130. [Google Scholar] [CrossRef]
  35. Schooler, S.S.; De Barro, P.; Ives, A.R. The potential for hyperparasitism to compromise biological control: Why don't hyperparasitoids drive their primary parasitoid hosts extinct? Biol. Control 2011, 58, 167–173. [Google Scholar] [CrossRef]
  36. Tougeron, K.; Tena, A. Hyperparasitoids as new targets in biological control in a global change context. Biol. Control 2019, 130, 164–171. [Google Scholar] [CrossRef]
  37. Ye, Z.; Vollhardt, I.M.G.; Tomanovic, Z.; Traugott, M. Evaluation of three molecular markers for identification of European primary parasitoids of cereal aphids and their hyperparasitoids. PLoS ONE 2017, 12, e0177376. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  38. Vorburger, C. The evolutionary ecology of symbiont-conferred resistance to parasitoids in aphids. Insect Sci. 2014, 21, 251–264. [Google Scholar] [PubMed] [Green Version]
  39. Carter, N.; McLean, I.F.G.; Watt, A.D.; Dixon, A.F.G. Cereal aphids: A case study and review. In Applied Biology; Coaker, T.H., Ed.; Academic Press: London, UK, 1980; pp. 271–348. [Google Scholar]
  40. Kröber, T.; Carl, K. Cereal aphids and their natural enemies in Europe-a literature review. Biocontrol News Inf. 1991, 12, 357–371. [Google Scholar]
  41. Vickerman, G.P.; Wratten, S.D. The biology and pest status of cereal aphids (Hemiptera: Aphididae) in Europe: A review. Bull. Entomol. Res. 1979, 69, 1–32. [Google Scholar] [CrossRef]
  42. Aljaryian, R.; Kumar, L. Changing global risk of invading greenbug Schizaphis graminum under climate change. Crop Prot. 2016, 88, 137–148. [Google Scholar] [CrossRef]
  43. Stanković, S.S.; Petrović, A.; Ilić Milošević, M.; Starý, P.; Kavallieratos, N.G.; Žikić, V.; Tomanović, Ž. Morphological and molecular characterization of common European species of Adialytus (Hymenoptera: Braconidae: Aphidiinae) based on the mtCOI barcoding gene and geometric morphometrics of forewings. Eur. J. Entomol. 2015, 112, 165–174. [Google Scholar] [CrossRef] [Green Version]
  44. Kavallieratos, N.G.; Tomanović, Ž.; Starý, P.; Athanassiou, C.G.; Sarlis, G.P.; Petrović, O.; Niketić, M.; Veroniki, M.A. A survey of aphid parasitoids (Hymenoptera: Braconidae: Aphidiinae) of Southeastern Europe and their aphid-plant associations. Appl. Entomol. Zool. 2004, 39, 527–563. [Google Scholar] [CrossRef] [Green Version]
  45. El-Mali, M.U.; Starý, P.; Sahbaz, A.; Ozsermeci, F. A review of aphid parasitoids (Hym.: Braconidae: Aphidiinae) of Turkey. Egypt. J. Biol. Pest Control 2004, 14, 355–370. [Google Scholar]
  46. Kocić, K.; Petrović, A.; Čkrkić, J.; Kavallieratos, N.G.; Rakhshani, E.; Arnó, J.; Aparicio, J.; Hebert, P.D.N.; Tomanović, Ž. Resolving the taxonomic status of potential biocontrol agents belonging to the neglected genus Lipolexis Förster (Hymenoptera: Braconidae: Aphidiinae) with descriptions of six new species. Insects 2020, 11, 667. [Google Scholar] [CrossRef]
  47. Čkrkić, J.; Petrović, A.; Kocić, K.; Kavallieratos, N.G.; Hebert, P.D.; Tomanović, Ž. Review of the world Monoctonina Mackauer 1961 (Hymenoptera: Braconidae: Aphidiinae): Key for their identification and descriptions of five new species. Zootaxa 2019, 4691, 359–385. [Google Scholar] [CrossRef]
  48. Mitrović, M.; Starý, P.; Jakovljević, M.; Petrović, A.; Žikić, V.; Hidalgo, N.P.; Tomanović, Ž. Integrative taxonomy of root aphid parasitoids from the genus Paralipsis (Hymenoptera: Braconidae: Aphidiinae) with description of new species. ZooKeys 2019, 831, 49–69. [Google Scholar] [CrossRef] [PubMed]
  49. Höller, C.; Borgemeister, C.; Haardt, H.; Powell, W. The relationship between primary parasitoids and hyperparasitoids of cereal aphids—An analysis of field data. J. Anim. Ecol. 1993, 62, 12–21. [Google Scholar] [CrossRef]
  50. Čkrkić, J.; Petrović, A.; Kocić, K.; Ye, Z.; Vollhardt, I.M.G.; Hebert, P.D.N.; Traugott, M.; Tomanović, Ž. Hidden in plain sight: Phylogeography of an overlooked parasitoid species Trioxys sunnysidensis Fulbright & Pike (Hymenoptera: Braconidae: Aphidiinae). Agric. Forest Entomol. 2019, 21, 299–308. [Google Scholar]
  51. Kos, K.; Petrović-Obradović, O.; Žikić, V.; Petrović, A.; Trdan, S.; Tomanović, Ž. Review of interactions between host plants, aphids, primary parasitoids and hyperparasitoids in vegetable and cereal ecosystems in Slovenia. J. Entomol. Res. Soc. 2012, 14, 67–78. [Google Scholar]
  52. Ferrer-Suay, M.; Selfa, J.; Tomanović, Ž.; Janković, M.; Kos, K.; Rakhshani, E.; Pujade-Villar, J. Revision of Alloxysta from the north-western Balkan Peninsula with description of two new species (Hymenoptera: Figitidae: Charipinae). Acta Entomol. Musei Natl. Pragae 2013, 53, 347–368. [Google Scholar]
  53. Ferrer-Suay, M.; Starý, P.; Selfa, J.; Puraje-Villar, J. Revision of charipine aphid hyperparasitoids (Hymenoptera: Cynipoidea: Figitidae) from central Europe. Entomol. Fenn. 2017, 28, 113–147. [Google Scholar] [CrossRef] [Green Version]
  54. Evenhuis, H.H.; Barbotin, F. Studies on Cynipidae Alloxystinae. 6. Phaenoglyphis villosa (Hartig) and Alloxysta arcuata (Kieffer). Entomol. Ber. 1977, 37, 184–190. [Google Scholar]
  55. Starý, P. Dendrocerus hyperparasites of aphids in Czechoslovakia (Hymenoptera: Ceraphronoidea). Acta Entomol. Bohemoslov. 1977, 74, 1–9. [Google Scholar]
  56. Pungerl, N.B. Morphometric and electrophoretic study of Aphidius species (Hymenoptera: Aphidiidae) reared from a variety of aphid hosts. Syst. Entomol. 1986, 11, 327–354. [Google Scholar] [CrossRef]
  57. Starý, P. Parasite spectrum and relative abundance of parasites of cereal aphids in Czechoslovakia (Hymenoptera: Aphidiidae, Homoptera: Aphidodea). Acta Entomol. Bohemoslov. 1976, 73, 216–233. [Google Scholar]
  58. Tomanović, Ž.; Kavallieratos, N.G.; Starý, P.; Stanisavljević, L.Z.; Ćetković, A.; Stamenković, S.; Jovanović, S.; Athanassiou, C.G. Regional tritrophic relationship patterns of five aphid parasitoid species (Hymenoptera: Braconidae: Aphidiinae) in agroecosystem-dominated landscapes of Southeastern Europe. J. Econ. Entomol. 2009, 102, 836–854. [Google Scholar] [CrossRef] [PubMed]
  59. Tomanović, Ž.; Kos, K.; Petrović, A.; Starý, P.; Kavallieratos, N.G.; Žikić, V.; Jakše, J.; Trdan, S.; Ivanović, A. The relationship between molecular variation and variation in the wing shape of three aphid parasitoid species: Aphidius uzbekistanicus Luzhetzki, Aphidius rhopalosiphi De Stefani Perez and Aphidius avenaphis (Fitch) (Hymenoptera: Braconidae: Aphidiinae). Zool. Anz. 2013, 252, 41–47. [Google Scholar] [CrossRef]
  60. Kos, K.; Petrović, A.; Starý, P.; Kavallieratos, N.G.; Ivanović, A.; Toševski, I.; Jakše, J.; Trdan, S.; Tomanovič, Ž. On the identity of cereal aphid parasitoid wasps Aphidius uzbekistanicus, Aphidius rhopalosiphi, and Aphidius avenaphis (Hymenoptera: Braconidae: Aphidiinae) by examination of COI mitochondrial gene, geometric morphometrics, and morphology. Ann. Entomol. Soc. Am. 2011, 104, 1221–1232. [Google Scholar] [CrossRef]
  61. Eady, R.D. A new diagnostic character in Aphidius (Hymenoptera: Braconidae) of special significance in species on pea aphid. Proc. R. Entomol. Soc. Lond. B Taxon. 1969, 38, 165–173. [Google Scholar] [CrossRef]
  62. Höller, C.A. Evidence for the existence of a species closely related to the cereal aphid parasitoid Aphidius rhopalosiphi De Stefani-Perez based on host ranges, morphological characters, isoelectric focusing banding patterns, cross-breeding experiments and sex pheromone specificities (Hymenoptera, Braconidae, Aphidiinae). Syst. Entomol. 1991, 16, 15–28. [Google Scholar]
  63. Starý, P. A review of the Aphidius species (Hymenoptera: Aphidiidae) of Europe. Annot. Zool. Bot. 1973, 84, 1–85. [Google Scholar]
  64. Tomanović, Ž.; Kavallieratos, N.G.; Starý, P.; Athanassiou, C.G.; Žikić, V.; Petrović-Obradović, O.; Sarlis, G.P. Aphidius Nees aphid parasitoids (Hymenoptera: Braconidae: Aphidiinae) in Serbia and Montenegro: Tritrophic associations and key. Acta Entomol. Serbica 2003, 8, 15–39. [Google Scholar]
  65. Tomanović, Ž.; Petrović, A.; Mitrović, M.; Kavallieratos, N.G.; Starý, P.; Rakhshani, E.; Rakhshanipour, M.; Popović, A.; Shukshuk, A.H.; Ivanović, A. Molecular and morphological variability within the Aphidius colemani group with redescription of Aphidius platensis Brethes (Hymenoptera: Braconidae: Aphidiinae). Bull. Entomol. Res. 2014, 104, 552–565. [Google Scholar] [CrossRef]
  66. Pennacchio, F. The Italian species of the genus Aphidius Nees (Hymenoptera, Braconidae, Aphidiinae). Boll. Lab. Entomol. Agrar. Portici 'Filippo Silvestri' 1989, 46, 75–106. [Google Scholar]
  67. Starý, P. Impact of an indigenous parasite, Aphidius ervi Hal., on pea aphid, Acyrthosiphon pisum (Harris) populations on alfalfa in Czechoslovakia. Boll. Lab. Entomol. Agrar. Portici 'Filippo Silvestri' 1968, 26, 293–313. [Google Scholar]
  68. Kavallieratos, N.G.; Tomanović, Ž.; Starý, P.; Athanassiou, C.G.; Fasseas, C.; Petrović, O.; Stanisavljević, L.Ž.; Anagnou, M.V. Praon Haliday (Hymenoptera: Braconidae: Aphidiinae) of Southeastern Europe: Key, host range and phylogenetic relationship. Zool. Anz. 2005, 243, 181–209. [Google Scholar] [CrossRef]
  69. Rakhshani, E.; Talebi, A.A.; Manzari, S.; Tomanović, Ž.; Starý, P.; Rezwani, P. Preliminary taxonomic study of genus Praon Haliday (Hymenoptera: Braconidae: Aphidiinae) and its host associations in Iran. J. Entomol. Soc. Iran 2007, 26, 19–34. [Google Scholar]
  70. Lazarević, M.; Stanković, S.S.; Petrović, A.; Ilić Milošević, M.; Tomanović, Ž.; Ivanović, A.; Žikić, V. Comparative morphometric analysis of petioles and forewings of the European Binodoxys Mackauer species (Hymenoptera: Braconidae: Aphidiinae). Zool. Anz. 2020, 284, 7–15. [Google Scholar] [CrossRef]
  71. Fulbright, J.L.; Pike, K.S. A new species of Trioxys (Hymenoptera: Braconidae: Aphidiinae) parasitic on the bird cherry-oat aphid, Rhopalosiphum padi (L.) (Hemiptera: Aphididae) in the Pacific Northwest. Proc. Entomol. Soc. Wash. 2007, 109, 541–546. [Google Scholar]
  72. Fulbright, J.L.; Pike, K.S.; Starý, P. A key to North American species of Trioxys Haliday (Hymenoptera: Braconidae: Aphidiinae), with a summary of the geographic distribution, hosts, and species diagnostic features. Proc. Entomol. Soc. Wash. 2007, 109, 779–790. [Google Scholar]
  73. Tomanović, Ž.; Mitrović, M.; Petrović, A.; Kavallieratos, N.G.; Žikić, V.; Ivanović, A.; Rakhshani, E.; Starý, P.; Vorburger, C. Revision of the European Lysiphlebus species (Hymenoptera: Braconidae: Aphidiinae) on the basis of COI and 28SD2 molecular markers and morphology. Arthropod Syst. Phylogeny 2018, 76, 179–213. [Google Scholar]
  74. Gärdenfors, U. Taxonomic and biological revision of Palearctic Ephedrus Haliday (Hymenoptera: Braconidae: Aphidiinae). Entomol. Scand. Suppl. 1986, 27, 1–95. [Google Scholar]
  75. Tomanović, Ž.; Petrović, A.; Starý, P.; Kavallieratos, N.G.; Žikić, V.; Rakhshani, E. Ephedrus Haliday (Hymenoptera: Braconidae: Aphidiinae) in Serbia and Montenegro: Tritrophic associations and key. Acta Entomol. Serbica 2009, 14, 39–53. [Google Scholar]
  76. Starý, P. A revision of the genus Diaeretiella Starý (Hymenoptera: Aphidiidae). Acta Entomol. Musei Natl. Pragae 1961, 34, 383–397. [Google Scholar]
  77. Singh, R.; Singh, G. Systematics, distribution and host range of Diaeretiella rapae (McIntosh) (Hymenoptera: Braconidae, Aphidiinae). Int. J. Res. Stud. Biosci. 2015, 3, 1–36. [Google Scholar]
  78. Rakhshani, E.; Starý, P.; Tomanović, Ž. Species of Adialytus Förster, 1862 (Hymenoptera: Braconidae: Aphidiinae) in Iran: Taxonomic notes and tritrophic associations. ZooKeys 2012, 221, 81–95. [Google Scholar] [CrossRef] [PubMed]
  79. Koponen, M.; Halme, J. New finds of Ephedrus and Toxares species (Hymenoptera: Braconidae: Aphidiinae) from Finland. Entomol. Fenn. 1993, 4, 31–36. [Google Scholar] [CrossRef] [Green Version]
  80. Starý, P. Notes on the parasites of the root aphids (Hymenoptera, Aphidiidae). Acta Soc. Entomol. Cechosloveniae 1961, 58, 228–238. [Google Scholar]
  81. Van Achterberg, C.; de Zugasti Carrón, N.F.O. Revision of the genus Paralipsis Foerster, 1863 (Hymenoptera: Braconidae), with the description of two new species. ZooKeys 2016, 606, 25–39. [Google Scholar] [CrossRef] [Green Version]
  82. Petrović, A. Sizing the knowledge gap in taxonomy: The last dozen years of Aphidiinae research. Insects 2022, 13, 170. [Google Scholar] [CrossRef]
  83. Derocles, S.A.P.; Plantegenest, M.; Simon, J.C.; Taberlet, P.; Le Ralec, A. A universal method for the detection and identification of Aphidiinae parasitoids within their aphid hosts. Mol. Ecol. Res. 2012, 12, 634–645. [Google Scholar] [CrossRef]
  84. Mitrovski-Bogdanović, A.; Petrović, A.; Mitrović, M.; Ivanović, A.; Žikić, V.; Starý, P.; Vorburger, C.; Tomanović, Ž. Identification of two cryptic species within the Praon abjectum group (Hymenoptera: Braconidae: Aphidiinae) using molecular markers and geometric morphometrics. Ann. Entomol. Soc. Am. 2014, 106, 170–180. [Google Scholar] [CrossRef]
  85. Starý, P.; Lyon, J.P.; Leclant, F. Biocontrol of aphids by the introduced Lysiphlebus testaceipes (Cress.) (Hym.: Aphidiidae) in Mediterranean France. J. Appl. Entomol. 1988, 105, 74–78. [Google Scholar] [CrossRef]
  86. Starý, P.; Lyon, J.P.; Leclant, F. Post-colonisation host range of Lysiphlebus testaceipes in the Mediterranean area (Hymenoptera: Aphidiidae). Acta Entomol. Bohemoslov. 1988, 85, 1–11. [Google Scholar]
  87. Starý, P.; Lumbierres, B.; Pons, X. Opportunistic changes in the host range of Lysiphlebus testaceipes (Cr.), an exotic aphid parasitoid expanding in the Iberian Peninsula. J. Pest Sci. 2004, 77, 139–144. [Google Scholar] [CrossRef]
  88. Hughes, G.E.; Sterk, G.; Bale, J.S. Thermal biology and establishment potential in temperate climates of the aphid parasitoid, Lysiphlebus testaceipes. BioControl 2011, 56, 19–33. [Google Scholar] [CrossRef]
  89. Kavallieratos, N.G.; Tomanović, Ž.; Petrović, A.; Kocić, K.; Janković, M.; Starý, P. Parasitoids (Hymenoptera: Braconidae: Aphidiinae) of aphids feeding on ornamental trees in Southeastern Europe: Key for identification and tritrophic associations. Ann. Entomol. Soc. Am. 2016, 109, 473–487. [Google Scholar] [CrossRef]
  90. Shrivastava, S.K. Status and management of wheat root aphid. JNKVV Res. J. 2012, 46, 166–171. [Google Scholar]
  91. Petrović, O. Aphids (Homoptera: Aphididae) on cereal crops. Rev. Res. Work Fac. Agr. 1996, 41, 159–168. [Google Scholar]
  92. Derocles, S.A.; Plantegenest, M.; Rasplus, J.Y.; Marie, A.; Evans, D.M.; Lunt, D.H.; Le Ralec, A. Are generalist Aphidiinae (Hym.: Braconidae) mostly cryptic species complexes? Syst. Entomol. 2016, 41, 379–391. [Google Scholar] [CrossRef]
  93. Derocles, S.A.; Navasse, Y.; Buchard, C.; Plantegenest, M.; Le Ralec, A. “Generalist” aphid parasitoids behave as specialists at the agroecosystem scale. Insects 2019, 11, 6. [Google Scholar] [CrossRef] [Green Version]
  94. Cameron, P.J.; Powell, W.; Loxdale, H.D. Reservoirs for Aphidius ervi Haliday (Hymenoptera: Aphidiidae), a polyphagous parasitoid of cereal aphids (Hemiptera: Aphididae). Bull. Entomol. Res. 1984, 74, 647–656. [Google Scholar] [CrossRef]
  95. Pennacchio, F.; Tremblay, E. Biosystematic and morphological study of two Aphidius ervi Haliday (Hymenoptera: Braconidae) biotypes with the description of a new species. Boll. Lab. Entomol. Agrar. Portici 'Filippo Silvestri' 1986, 43, 105–117. [Google Scholar]
  96. Petrović, A.; Tomanović, Ž.; Žikić, V. Wahlgreniella ossiannilssoni Hille Ris Lambers, a new host for Aphidius microlophii Pennacchio and Tremblay (Hymenoptera: Braconidae: Aphidiinae). Arch. Biol. Sci. 2006, 58, 41–42. [Google Scholar] [CrossRef]
  97. Postic, E.; Outreman, Y.; Derocles, S.; Granado, C.; Le Ralec, A. Genetics of wild and mass-reared populations of a generalist aphid parasitoid and improvement of biological control. PLoS ONE 2021, 16, e0249893. [Google Scholar] [CrossRef]
  98. Rakhshani, E.; Barahoei, H.; Ahmad, Z.; Starý, P.; Ghafouri-Moghaddam, M.; Mehrparvar, M.; Kavallieratos, N.G.; Čkrkić, J.; Tomanović, Ž. Review of Aphidiinae parasitoids (Hymenoptera: Braconidae) of the Middle East and North Africa: Key to species and host associations. Eur. J. Taxon. 2019, 552, 1–132. [Google Scholar] [CrossRef]
  99. Mitrovski-Bogdanović, A.; Tomanović, Ž.; Mitrović, M.; Petrović, A.; Ivanović, A.; Žikić, V.; Starý, P.; Vorburger, C. The Praon dorsale–yomenae s.str. complex (Hymenoptera: Braconidae: Aphidiinae): Species discrimination using geometric morphometrics and molecular markers with description of a new species. Zool. Anz. 2014, 253, 270–282. [Google Scholar] [CrossRef]
  100. Gagić, V.; Petrović-Obradović, O.; Frund, J.; Kavallieratos, N.G.; Athanassiou, C.G.; Starý, P.; Tomanović, Ž. The effects of aphid traits on parasitoid host use and specialist advantage. PLoS ONE 2016, 11, 14. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  101. Heie, O.E. Fauna Entomologica Scandinavica, 17. The Aphidoidea of Fennoscandia and Denmark (III). The Family Aphididae: Subfamily Pterocommatinae & Tribe Aphidini of Subfamily Aphididae; Lubrecht & Cramer: Port Jervis, NY, USA, 1986. [Google Scholar]
  102. Heie, O.E. Fauna Entomologica Scandinavica, 25. The Aphidoidea of Fennoscandia and Denmark (IV). Family Aphididae: Part 1 of the Tribe Macrosiphini of subfamily Aphidinae; Lubrecht & Cramer: Port Jervis, NY, USA, 1992. [Google Scholar]
  103. Heie, O.E. Fauna Entomologica Scandinavica, 28. The Aphidoidea of Fennoscandia and Denmark (V). Family Aphididae: Part 2 of the Tribe Macrosiphini of subfamily Aphidinae; Lubrecht & Cramer: Port Jervis, NY, USA, 1994. [Google Scholar]
  104. Pankanin-Franczyk, M.; Sobota, G. Relationships between primary and secondary parasitoids of cereal aphids. J. Appl. Entomol. 1998, 122, 389–395. [Google Scholar] [CrossRef]
  105. Kavallieratos, N.G.; Lykouressis, D.P.; Sarlis, G.P.; Stathas, G.J.; Segovia, A.S.; Athanassiou, C.G. The Aphidiinae (Hymenoptera: Ichneumonoidea: Braconidae) of Greece. Phytoparasitica 2001, 29, 306–340. [Google Scholar] [CrossRef]
  106. Poulin, R. Large-scale patterns of host use by parasites of freshwater fishes. Ecol. Lett. 1998, 1, 118–128. [Google Scholar] [CrossRef]
  107. Poulin, R. Relative infection levels and taxonomic distances among the host species used by a parasite: Insights into parasite specialization. Parasitology 2005, 130, 109–115. [Google Scholar] [CrossRef] [Green Version]
  108. Straub, C.S.; Ives, A.R.; Gratton, C. Evidence for a trade-off between host-range breadth and host-use efficiency in aphid parasitoids. Am. Nat. 2011, 177, 389–395. [Google Scholar] [CrossRef] [Green Version]
  109. Colles, A.; Liow, L.H.; Prinzing, A. Are specialists at risk under environmental change? Neoecological, paleoecological and phylogenetic approaches. Ecol. Lett. 2009, 12, 849–863. [Google Scholar] [CrossRef] [Green Version]
  110. Raymond, L.; Plantegenest, M.; Gagić, V.; Navasse, Y.; Lavandero, B. Aphid parasitoid generalism: Development, assessment, and implications for biocontrol. J. Pest Sci. 2016, 89, 7–20. [Google Scholar] [CrossRef]
  111. Van Baaren, J.; Héterier, V.; Hance, T.; Krespi, L.; Cortesero, A.M.; Poinsot, D.; Le Ralec, A.; Outreman, Y. Playing the hare or the tortoise in parasitoids: Could different oviposition strategies have an influence in host partitioning in two Aphidius species? Ethol. Ecol. Evol. 2004, 16, 231–242. [Google Scholar] [CrossRef]
  112. van Baaren, J.; Le Lann, C.; Pichenot, J.; Pierre, J.S.; Krespi, L.; Outreman, Y. How could host discrimination abilities influence the structure of a parasitoid community? Bull. Entomol. Res. 2009, 99, 299–306. [Google Scholar] [CrossRef] [PubMed]
  113. Battaglia, D.; Poppy, G.; Powell, W.; Romano, A.; Tranfaglia, A.; Pennacchio, F. Physical and chemical cues influencing the oviposition behaviour of Aphidius ervi. Entomol. Exp. Appl. 2000, 94, 219–227. [Google Scholar] [CrossRef]
  114. Weinbrenner, M.; Völkl, W. Oviposition behaviour of the aphid parasitoid, Aphidius ervi: Are wet aphids recognized as host? Entomol. Exp. Appl. 2002, 103, 51–59. [Google Scholar] [CrossRef]
  115. Michaud, J.P. The ecological significance of aphid cornicles and their secretions. Annu. Rev. Entomol. 2022, 67, 65–81. [Google Scholar] [CrossRef]
  116. Outreman, Y.; Le Ralec, A.; Plantegenest, M.; Chaubet, B.; Pierre, J.S. Superparasitism limitation in an aphid parasitoid: Cornicle secretion avoidance and host discrimination ability. J. Insect Physiol. 2001, 47, 339–348. [Google Scholar] [CrossRef]
  117. Poulin, R. “Adaptive” changes in the behaviour of parasitized animals: A critical review. Int. J. Parasitol. 1995, 25, 1371–1383. [Google Scholar] [CrossRef]
  118. Le Ralec, A.; Anselme, C.; Outreman, Y.; Poirié, M.; Van Baaren, J.; Le Lann, C.; Jacques, J.M. Evolutionary ecology of the interactions between aphids and their parasitoids. C. R. Biol. 2010, 333, 554–565. [Google Scholar] [CrossRef]
  119. Godfray, H.C.J. Parasitoids: Behavioural and Evolutionary Ecology; Princeton University Press: Princeton, Chichester, UK, 1994. [Google Scholar]
  120. Brodeur, J.; Rosenheim, J.A. Intraguild interactions in aphid parasitoids. Entomol. Exp. Appl. 2000, 97, 93–108. [Google Scholar] [CrossRef] [Green Version]
  121. Desneux, N.; Barta, R.J.; Hoelmer, K.A.; Hopper, K.R.; Heimpel, G.E. Multifaceted determinants of host specificity in an aphid parasitoid. Oecologia 2009, 160, 387–398. [Google Scholar] [CrossRef]
  122. Sullivan, D.J. Insect hyperparasitism. Annu. Rev. Entomol. 1987, 32, 49–70. [Google Scholar] [CrossRef]
  123. Sullivan, D.J.; Völkl, W. Hyperparasitism: Multitrophic ecology and behavior. Annu. Rev. Entomol. 1999, 44, 291–315. [Google Scholar] [CrossRef] [PubMed]
  124. Lohaus, K.; Vidal, S.; Thies, C. Farming practices change food web structures in cereal aphid-parasitoid-hyperparasitoid communities. Oecologia 2013, 171, 249–259. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  125. Ferrer-Suay, M.; Staverløkk, A.; Selfa, J.; Pujade-Villar, J.; Naik, S.; Ekrem, T. Nuclear and mitochondrial markers suggest new species boundaries in Alloxysta (Hymenoptera: Cynipoidea: Figitidae). Arthropod Syst. Phylogeny 2018, 76, 463–473. [Google Scholar]
  126. Rosenheim, J.A. Higher-order predators and the regulation of insect herbivore populations. Annu. Rev. Entomol. 1998, 43, 421–447. [Google Scholar] [CrossRef] [Green Version]
  127. Boivin, G.; Brodeur, J. Intra-and interspecific interactions among parasitoids: Mechanisms, outcomes and biological control. In Trophic and Guild in Biological Interactions Control; Brodeur, J., Boivin, G., Eds.; Springer: Dordrecht, The Netherlands, 2006; pp. 123–144. [Google Scholar]
  128. Holt, R.D.; Hochberg, M.E. The coexistence of competing parasites. Part II—Hyperparasitism and food chain dynamics. J. Theor. Biol. 1998, 193, 485–495. [Google Scholar] [CrossRef]
  129. Cusumano, A.; Harvey, J.A.; Bourne, M.E.; Poelman, E.H.; de Boer, J.G. Exploiting chemical ecology to manage hyperparasitoids in biological control of arthropod pests. Pest Manag. Sci. 2020, 76, 432–443. [Google Scholar] [CrossRef] [Green Version]
  130. Forrest, J.R. Complex responses of insect phenology to climate change. Curr. Opin. Insect Sci. 2016, 17, 49–54. [Google Scholar] [CrossRef]
  131. Gómez-Marco, F.; Urbaneja, A.; Jaques, J.A.; Rugman-Jones, P.F.; Stouthamer, R.; Tena, A. Untangling the aphid-parasitoid food web in citrus: Can hyperparasitoids disrupt biological control? Biol. Control 2015, 81, 111–121. [Google Scholar] [CrossRef]
  132. Neuville, S.; Le Ralec, A.; Outreman, Y.; Jaloux, B. The delay in arrival of the parasitoid Diaeretiella rapae influences the efficiency of cabbage aphid biological control. BioControl 2016, 61, 115–126. [Google Scholar] [CrossRef]
  133. Oliver, K.M.; Noge, K.; Huang, E.M.; Campos, J.M.; Becerra, J.X.; Hunter, M.S. Parasitic wasp responses to symbiont-based defense in aphids. BMC Biol. 2012, 10, 11. [Google Scholar] [CrossRef] [PubMed]
  134. Nakashima, Y.; Higashimura, Y.; Mizutani, K. Host discrimination and ovicide by aphid hyperparasitoids Asaphes suspensus (Hymenoptera: Pteromalidae) and Dendrocerus carpenteri (Hymenoptera: Megaspilidae). Appl. Entomol. Zool. 2016, 51, 609–614. [Google Scholar] [CrossRef]
  135. Liang, Q.F.; Liu, T.X. Interspecific host discrimination and intrinsic competition between Aphelinus asychis and Aphidius gifuensis in Myzus persicae. Entomol. Exp. Appl. 2017, 163, 265–271. [Google Scholar] [CrossRef]
  136. Greenstone, M.H. Molecular methods for assessing insect parasitism. Bull. Entomol. Res. 2006, 96, 1–13. [Google Scholar] [CrossRef] [PubMed]
  137. Traugott, M.; Kamenova, S.; Ruess, L.; Seeber, J.; Plantegenest, M. Empirically characterising trophic networks: What emerging DNA-based methods, stable isotope and fatty acid analyses can offer. Adv. Ecol. Res. 2013, 49, 177–224. [Google Scholar]
  138. Rand, T.; van Veen, F.J.F.; Tscharntke, T. Landscape complexity differentially benefits generalized fourth, over specialized third, trophic level natural enemies. Ecography 2012, 35, 97–104. [Google Scholar] [CrossRef]
  139. Gagić, V.; Tscharntke, T.; Dormann, C.F.; Gruber, B.; Wilstermann, A.; Thies, C. Food web structure and biocontrol in a four-trophic level system across a landscape complexity gradient. Proc. Royal Soc. B 2011, 278, 2946–2953. [Google Scholar] [CrossRef]
  140. Gariepy, T.; Kuhlmann, U.; Gillott, C.; Erlandson, M. A large-scale comparison of conventional and molecular methods for the evaluation of host-parasitoid associations in non-target risk-assessment studies. J. Appl. Ecol. 2008, 45, 708–715. [Google Scholar] [CrossRef]
  141. Hrcek, J.; Godfray, H.C. What do molecular methods bring to host-parasitoid food webs? Trends Parasitol. 2015, 31, 30–35. [Google Scholar] [CrossRef]
  142. Macfadyen, S.; Gibson, R.; Raso, L.; Sint, D.; Traugott, M.; Memmott, J. Parasitoid control of aphids in organic and conventional farming systems. Agric. Ecosyst. Environ. 2009, 133, 14–18. [Google Scholar] [CrossRef]
  143. Yang, F.; Yao, Z.W.; Zhu, Y.L.; Wu, Y.K.; Liu, L.T.; Liu, B.; Desneux, N.; Lu, Y.H. A molecular detection approach for assessing wheat aphid-parasitoid food webs in northern China. Entomol. Gen. 2020, 40, 273–284. [Google Scholar] [CrossRef]
  144. Zhu, Y.C.; Greenstone, M.H. Polymerase chain reaction techniques for distinguishing three species and two strains of Aphelinus (Hymenoptera: Aphelinidae) from Diuraphis noxia and Schizaphis graminum (Homoptera: Aphididae). Ann. Entomol. Soc. Am. 1999, 92, 71–79. [Google Scholar] [CrossRef]
  145. Rennstam Rubbmark, O.; Sint, D.; Cupic, S.; Traugott, M. When to use next generation sequencing or diagnostic PCR in diet analyses. Mol. Ecol. Resour. 2019, 19, 388–399. [Google Scholar] [CrossRef] [PubMed]
  146. González-Chang, M.; Wratten, S.D.; Lefort, M.C.; Boyer, S. Food webs and biological control: A review of molecular tools used to reveal trophic interactions in agricultural systems. Food Webs 2016, 9, 4–11. [Google Scholar] [CrossRef]
  147. Wirta, H.K.; Hebert, P.D.N.; Kaartinen, R.; Prosser, S.W.; Varkonyi, G.; Roslin, T. Complementary molecular information changes our perception of food web structure. Proc. Natl. Acad. Sci. USA 2014, 111, 1885–1890. [Google Scholar] [CrossRef] [Green Version]
  148. Lefort, M.C.; Wratten, S.; Cusumano, A.; Varennes, Y.D.; Boyer, S. Disentangling higher trophic level interactions in the cabbage aphid food web using high-throughput DNA sequencing. Metabarcoding Metagenom. 2017, 1, e13709. [Google Scholar] [CrossRef]
  149. Evans, D.M.; Kitson, J.N.J.; Lunt, H.D.; Straw, A.N.; Pocock, J.O.M. Merging DNA metabarcoding and ecological network analysis to understand and build resilient terrestrial ecosystems. Funct. Ecol. 2016, 30, 1904–1916. [Google Scholar] [CrossRef] [Green Version]
  150. Evans, D.M.; Kitson, J.J.N. Molecular ecology as a tool for understanding pollination and other plant-insect interactions. Curr. Opin. Insect Sci. 2020, 38, 26–33. [Google Scholar] [CrossRef]
  151. Gariepy, T.D.; Kuhlmann, U.; Gillott, C.; Erlandson, M. Parasitoids, predators and PCR: The use of diagnostic molecular markers in biological control of arthropods. J. Appl. Entomol. 2007, 131, 225–240. [Google Scholar] [CrossRef]
  152. Pocock, M.J.O.; Evans, D.M.; Memmott, J. The robustness and restoration of a network of ecological networks. Science 2012, 335, 973–977. [Google Scholar] [CrossRef] [Green Version]
  153. Bukovinszky, T.; van Veen, F.J.F.; Jongema, Y.; Dicke, M. Direct and indirect effects of resource quality on food web structure. Science 2008, 319, 804–807. [Google Scholar] [CrossRef] [PubMed]
  154. Tylianakis, J.M.; Tscharntke, T.; Lewis, O.T. Habitat modification alters the structure of tropical host-parasitoid food webs. Nature 2007, 445, 202–205. [Google Scholar] [CrossRef] [PubMed]
  155. Morris, R.J.; Lewis, O.T.; Godfray, H.C.J. Experimental evidence for apparent competition in a tropical forest food web. Nature 2004, 428, 310–313. [Google Scholar] [CrossRef] [PubMed]
  156. van Veen, F.J.F.; Morris, R.J.; Godfray, H.C.J. Apparent competition, quantitative food webs, and the structure of phytophagous insect communities. Annu. Rev. Entomol. 2006, 51, 187–208. [Google Scholar] [CrossRef]
  157. Derocles, S.A.P.; Le Ralec, A.; Besson, M.M.; Maret, M.; Walton, A.; Evans, D.M.; Plantegenest, M. Molecular analysis reveals high compartmentalization in aphid-primary parasitoid networks and low parasitoid sharing between crop and noncrop habitats. Mol. Ecol. 2014, 23, 3900–3911. [Google Scholar] [CrossRef]
  158. Alhmedi, A.; Haubruge, E.; D’Hoedt, S.; Francis, F. Quantitative food webs of herbivore and related beneficial community in non-crop and crop habitats. Biol. Control 2011, 58, 103–112. [Google Scholar] [CrossRef] [Green Version]
  159. Andrade, T.O.; Krespi, L.; Bonnardot, V.; van Baaren, J.; Outreman, Y. Impact of change in winter strategy of one parasitoid species on the diversity and function of a guild of parasitoids. Oecologia 2016, 180, 877–888. [Google Scholar] [CrossRef]
  160. Traugott, M.; Bell, J.R.; Raso, L.; Sint, D.; Symondson, W.O.C. Generalist predators disrupt parasitoid aphid control by direct and coincidental intraguild predation. Bull. Entomol. Res. 2012, 102, 239–247. [Google Scholar] [CrossRef]
  161. Frago, E.; Godfray, H.C.J. Avoidance of intraguild predation leads to a long-term positive trait-mediated indirect effect in an insect community. Oecologia 2014, 174, 943–952. [Google Scholar] [CrossRef]
  162. Tougeron, K.; Le Lann, C.; Brodeur, J.; van Baaren, J. Are aphid parasitoids from mild winter climates losing their winter diapause? Oecologia 2017, 183, 619–629. [Google Scholar] [CrossRef]
  163. Tougeron, K.; Damien, M.; Le Lann, C.; Brodeur, J.; van Baaren, J. Rapid responses of winter aphid-parasitoid communities to climate warming. Front. Ecol. Evol. 2018, 6, 173. [Google Scholar] [CrossRef] [Green Version]
  164. Derocles, S.A.P.; Lunt, D.H.; Berthe, S.C.F.; Nichols, P.C.; Moss, E.D.; Evans, D.M. Climate warming alters the structure of farmland tritrophic ecological networks and reduces crop yield. Mol. Ecol. 2018, 27, 4931–4946. [Google Scholar] [CrossRef] [PubMed]
  165. Ortiz-Martinez, S.A.; Lavandero, B. The effect of landscape context on the biological control of Sitobion avenae: Temporal partitioning response of natural enemy guilds. J. Pest Sci. 2018, 91, 41–53. [Google Scholar] [CrossRef]
  166. Hawro, V.; Ceryngier, P.; Tscharntke, T.; Thies, C.; Gagić, V.; Bengtsson, J.; Bommarco, R.; Winqvist, C.; Weisser, W.W.; Clement, L.W.; et al. Landscape complexity is not a major trigger of species richness and food web structure of European cereal aphid parasitoids. BioControl 2015, 60, 451–461. [Google Scholar] [CrossRef] [Green Version]
  167. Jeavons, E.; van Baaren, J.; Le Ralec, A.; Buchard, C.; Duval, F.; Llopis, S.; Postic, E.; Le Lann, C. Third and fourth trophic level composition shift in an aphid-parasitoid-hyperparasitoid food web limits aphid control in an intercropping system. J. Appl. Ecol. 2021, 59, 300–313. [Google Scholar] [CrossRef]
  168. Damien, M.; Le Lann, C.; Desneux, N.; Alford, L.; Al Hassan, D.; Georges, R.; Van Baaren, J. Flowering cover crops in winter increase pest control but not trophic link diversity. Agric. Ecosyst. Environ. 2017, 247, 418–425. [Google Scholar] [CrossRef] [Green Version]
  169. Kampfraath, A.A.; Giesen, D.; van Gestel, C.A.M.; Le Lann, C. Pesticide stress on plants negatively affects parasitoid fitness through a bypass of their phytophage hosts. Ecotoxicology 2017, 26, 383–395. [Google Scholar] [CrossRef]
  170. Ghising, K.; Harmon, J.P.; Beauzay, P.B.; Prischmann-Voldseth, D.A.; Helms, T.C.; Ode, P.J.; Knodel, J.J. Impact of Rag1 aphid resistant soybeans on Binodoxys communis (Hymenoptera: Braconidae), a parasitoid of soybean aphid (Hemiptera: Aphididae). Environ. Entomol. 2012, 41, 282–288. [Google Scholar] [CrossRef] [Green Version]
  171. Hodge, S.; Ward, J.L.; Galster, A.M.; Beale, M.H.; Powell, G. The effects of a plant defence priming compound, beta-aminobutyric acid, on multitrophic interactions with an insect herbivore and a hymenopterous parasitoid. BioControl 2011, 56, 699–711. [Google Scholar] [CrossRef]
  172. Karatolos, N.; Hatcher, P.E. The effect of acetylsalicylic acid and oxalic acid on Myzus persicae and Aphidius colemani. Entomol. Exp. Appl. 2009, 130, 98–105. [Google Scholar] [CrossRef]
  173. Kos, M.; Houshyani, B.; Achhami, B.B.; Wietsma, R.; Gols, R.; Weldegergis, B.T.; Kabouw, P.; Bouwmeester, H.J.; Vet, L.E.M.; Dicke, M.; et al. Herbivore-mediated effects of glucosinolates on different natural enemies of a specialist aphid. J. Chem. Ecol. 2012, 38, 100–115. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  174. Von Burg, S.; van Veen, F.J.F.; Alvarez-Alfageme, F.; Romeis, J. Aphid-parasitoid community structure on genetically modified wheat. Biol. Lett. 2011, 7, 387–391. [Google Scholar] [CrossRef] [PubMed]
  175. Cowgill, S.E.; Danks, C.; Atkinson, H.J. Multitrophic interactions involving genetically modified potatoes, nontarget aphids, natural enemies and hyperparasitoids. Mol. Ecol. 2004, 13, 639–647. [Google Scholar] [CrossRef] [PubMed]
  176. Omacini, M.; Chaneton, E.J.; Ghersa, C.M.; Muller, C.B. Symbiotic fungal endophytes control insect host-parasite interaction webs. Nature 2001, 409, 78–81. [Google Scholar] [CrossRef] [PubMed]
  177. Omacini, M.; Eggers, T.; Bonkowski, M.; Gange, A.C.; Jones, T.H. Leaf endophytes affect mycorrhizal status and growth of co-infected and neighbouring plants. Funct. Ecol. 2006, 20, 226–232. [Google Scholar] [CrossRef]
  178. Härri, S.A.; Krauss, J.; Müller, C.B. Trophic cascades initiated by fungal plant endosymbionts impair reproductive performance of parasitoids in the second generation. Oecologia 2008, 157, 399–407. [Google Scholar] [CrossRef] [Green Version]
  179. Härri, S.A.; Krauss, J.; Müller, C.B. Extended larval development time for aphid parasitoids in the presence of plant endosymbionts. Ecol. Entomol. 2009, 34, 20–25. [Google Scholar] [CrossRef] [Green Version]
  180. Bultman, T.L.; Aguilera, A.; Sullivan, T. Influence of fungal isolates infecting tall fescue on multitrophic interactions. Fungal Ecol. 2012, 5, 372–378. [Google Scholar] [CrossRef]
  181. Hempel, S.; Stein, C.; Unsicker, S.B.; Renker, C.; Auge, H.; Weisser, W.W.; Buscot, F. Specific bottom-up effects of arbuscular mycorrhizal fungi across a plant-herbivore-parasitoid system. Oecologia 2009, 160, 267–277. [Google Scholar] [CrossRef] [Green Version]
  182. Albittar, L.; Ismail, M.; Lohaus, G.; Ameline, A.; Visser, B.; Bragard, C.; Hance, T. Bottom-up regulation of a tritrophic system by Beet yellows virus infection: Consequences for aphid-parasitoid foraging behaviour and development. Oecologia 2019, 191, 113–125. [Google Scholar] [CrossRef]
  183. Calvo, D.; Fereres, A. The performance of an aphid parasitoid is negatively affected by the presence of a circulative plant virus. BioControl 2011, 56, 747–757. [Google Scholar] [CrossRef]
  184. Colfer, R.G.; Rosenheim, J.A. Predation on immature parasitoids and its impact on aphid suppression. Oecologia 2001, 126, 292–304. [Google Scholar] [CrossRef]
  185. Costamagna, A.C.; Landis, D.A.; Difonzo, C.D. Suppression of soybean aphid by generalist predators results in a trophic cascade in soybeans. Ecol. Appl. 2007, 17, 441–451. [Google Scholar] [CrossRef] [Green Version]
  186. Sanders, D.; Van Veen, F.J.F. The impact of an ant-aphid mutualism on the functional composition of the secondary parasitoid community. Ecol. Entomol. 2010, 35, 704–710. [Google Scholar] [CrossRef]
  187. Oliver, K.M.; Russell, J.A.; Moran, N.A.; Hunter, M.S. Facultative bacterial symbionts in aphids confer resistance to parasitic wasps. Proc. Natl. Acad. Sci. USA 2003, 100, 1803–1807. [Google Scholar] [CrossRef] [Green Version]
  188. Moran, N.A.; Russell, J.A.; Koga, R.; Fukatsu, T. Evolutionary relationships of three new species of Enterobacteriaceae living as symbionts of aphids and other insects. Appl. Environ. Microbiol. 2005, 71, 3302–3310. [Google Scholar] [CrossRef] [Green Version]
  189. Vorburger, C.; Sandrock, C.; Gouskov, A.; Castañeda, L.E.; Ferrari, J. Genotypic variation and the role of defensive endosymbionts in an all-parthenogenetic host-parasitoid interaction. Evolution 2009, 63, 1439–1450. [Google Scholar] [CrossRef] [Green Version]
  190. Vorburger, C.; Gehrer, L.; Rodriguez, P. A strain of the bacterial symbiont Regiella insecticola protects aphids against parasitoids. Biol. Lett. 2010, 6, 109–111. [Google Scholar] [CrossRef] [Green Version]
  191. Brady, C.M.; White, J.A. Cowpea aphid (Aphis craccivora) associated with different host plants has different facultative endosymbionts. Ecol. Entomol. 2013, 38, 433–437. [Google Scholar] [CrossRef]
  192. Wu, T.; Monnin, D.; Lee, R.A.R.; Henry, L.M. Local adaptation to hosts and parasitoids shape Hamiltonella defensa genotypes across aphid species. Proc. R. Soc. B 2022, 289, 20221269. [Google Scholar] [CrossRef]
  193. Leybourne, D.J.; Bos, J.I.B.; Valentine, T.A.; Karley, A.J. The price of protection: A defensive endosymbiont impairs nymph growth in the bird cherry-oat aphid, Rhopalosiphum padi. Insect Sci. 2020, 27, 68–85. [Google Scholar] [CrossRef] [Green Version]
  194. Scarborough, C.L.; Ferrari, J.; Godfray, H.C.J. Aphid protected from pathogen by endosymbiont. Science 2005, 310, 1781. [Google Scholar] [CrossRef]
  195. Łukasik, P.; van Asch, M.; Guo, H.F.; Ferrari, J.; Godfray, H.C.J. Unrelated facultative endosymbionts protect aphids against a fungal pathogen. Ecol. Lett. 2013, 16, 214–218. [Google Scholar] [CrossRef]
  196. Parker, B.J.; Hrček, J.; McLean, A.H.C.; Godfray, H.C.J. Genotype specificity among hosts, pathogens, and beneficial microbes influences the strength of symbiont-mediated protection. Evolution 2017, 71, 1222–1231. [Google Scholar] [CrossRef] [Green Version]
  197. Guo, J.; Hatt, S.; He, K.; Chen, J.; Francis, F.; Wang, Z. Nine facultative endosymbionts in aphids. A review. J. Asia Pac. Entomol. 2017, 20, 794–801. [Google Scholar] [CrossRef]
  198. Zytynska, S.E.; Weisser, W.W. The natural occurrence of secondary bacterial symbionts in aphids. Ecol. Entomol. 2016, 41, 13–26. [Google Scholar] [CrossRef] [Green Version]
  199. Łukasik, P.; Dawid, M.A.; Ferrari, J.; Godfray, H.C.J. The diversity and fitness effects of infection with facultative endosymbionts in the grain aphid, Sitobion avenae. Oecologia 2013, 173, 985–996. [Google Scholar] [CrossRef]
  200. Alkhedir, H.; Karlovsky, P.; Mashaly, A.M.A.; Vidal, S. Phylogenetic relationships of the symbiotic bacteria in the aphid Sitobion avenae (Hemiptera: Aphididae). Environ. Entomol. 2015, 44, 1358–1366. [Google Scholar] [CrossRef]
  201. Fakhour, S.; Ambroise, J.; Renoz, F.; Foray, V.; Gala, J.L.; Hance, T. A large-scale field study of bacterial communities in cereal aphid populations across Morocco. FEMS Microbiol. Ecol. 2018, 94, fiy003. [Google Scholar] [CrossRef] [Green Version]
  202. Liu, X.D.; Lei, H.X.; Chen, F.F. Infection pattern and negative effects of a facultative endosymbiont on its insect host are environment-dependent. Sci. Rep. 2019, 9, 4013. [Google Scholar] [CrossRef] [Green Version]
  203. Zepeda-Paulo, F.; Lavandero, B. Effect of the genotypic variation of an aphid host on the endosymbiont associations in natural host populations. Insects 2021, 12, 217. [Google Scholar] [CrossRef]
  204. Li, T.; Xiao, J.H.; Xu, Z.H.; Murphy, R.W.; Huang, D.W. Cellular tropism, population dynamics, host range and taxonomic status of an aphid secondary symbiont, SMLS (Sitobion miscanthi L Type symbiont). PLoS ONE 2011, 6, e21944. [Google Scholar] [CrossRef]
  205. Li, T.; Xiao, J.H.; Wu, Y.Q.; Huang, D.W. Diversity of bacterial symbionts in populations of Sitobion miscanthi (Hemiptera: Aphididae) in China. Environ. Entomol. 2014, 43, 605–611. [Google Scholar] [CrossRef]
  206. Li, T.; Wu, X.J.; Jiang, Y.L.; Zhang, L.; Duan, Y.; Miao, J.; Gong, Z.J.; Wu, Y.Q. The genetic diversity of SMLS (Sitobion miscanthi L type symbiont) and its effect on the fitness, mitochondrial DNA diversity and Buchnera aphidicola dynamic of wheat aphid, Sitobion miscanthi (Hemiptera: Aphididae). Mol. Ecol. 2016, 25, 3142–3151. [Google Scholar] [CrossRef]
  207. Guo, J.; Liu, X.; Poncelet, N.; He, K.; Francis, F.; Wang, Z. Detection and geographic distribution of seven facultative endosymbionts in two Rhopalosiphum aphid species. MicrobiologyOpen 2019, 8, e00817. [Google Scholar] [CrossRef] [Green Version]
  208. Csorba, A.B.; Fora, C.G.; Bálint, J.; Felföldi, T.; Szabó, A.; Máthé, I.; Loxdale, H.D.; Kentelky, E.; Nyárádi, I.I.; Balog, A. Endosymbiotic bacterial diversity of corn leaf aphid, Rhopalosiphum maidis Fitch (Hemiptera: Aphididae) associated with maize management systems. Microorganisms 2022, 10, 939. [Google Scholar] [CrossRef]
  209. Telesnicki, M.C.; Ghersa, C.M.; Martinez-Ghersa, M.A.; Arneodo, J.D. Molecular identification of the secondary endosymbiont Hamiltonella defensa in the rose-grain aphid Metopolophium dirhodum. Rev. Argent. Microbiol. 2012, 44, 255–258. [Google Scholar]
  210. Sepúlveda, D.A.; Zepeda-Paulo, F.; Ramírez, C.C.; Lavandero, B.; Figueroa, C.C. Diversity, frequency, and geographic distribution of facultative bacterial endosymbionts in introduced aphid pests. Insect Sci. 2017, 24, 511–521. [Google Scholar] [CrossRef]
  211. Oliver, K.M.; Higashi, C.H.V. Variations on a protective theme: Hamiltonella defensa infections in aphids variably impact parasitoid success. Curr. Opin. Insect. Sci. 2019, 32, 1–7. [Google Scholar] [CrossRef]
  212. Oliver, K.M.; Moran, N.A.; Hunter, M.S. Variation in resistance to parasitism in aphids is due to symbionts not host genotype. Proc. Natl. Acad. Sci. USA 2005, 102, 12795–12800. [Google Scholar] [CrossRef] [Green Version]
  213. Schmid, M.; Sieber, R.; Zimmermann, Y.S.; Vorburger, C. Development, specificity and sublethal effects of symbiont-conferred resistance to parasitoids in aphids. Funct. Ecol. 2012, 26, 207–215. [Google Scholar] [CrossRef] [Green Version]
  214. Cayetano, L.; Vorburger, C. Symbiont-conferred protection against hymenopteran parasitoids in aphids: How general is it? Ecol. Entomol. 2015, 40, 85–93. [Google Scholar] [CrossRef] [Green Version]
  215. Asplen, M.K.; Bano, N.; Brady, C.M.; Desneux, N.; Hopper, K.R.; Malouines, C.; Oliver, K.M.; White, J.A.; Heimpel, G.E. Specialisation of bacterial endosymbionts that protect aphids from parasitoids. Ecol. Entomol. 2014, 39, 736–739. [Google Scholar] [CrossRef]
  216. McLean, A.H.C.; Godfray, H.C.J. Evidence for specificity in symbiont-conferred protection against parasitoids. Proc. Royal Soc. B 2015, 282, 20150977. [Google Scholar] [CrossRef]
  217. Gimmi, E.; Vorburger, C. Strong genotype-by-genotype interactions between aphid-defensive symbionts and parasitoids persist across different biotic environments. J. Evol. Biol. 2021, 34, 1944–1953. [Google Scholar] [CrossRef]
  218. Dion, E.; Zélé, F.; Simon, J.C.; Outreman, Y. Rapid evolution of parasitoids when faced with the symbiont-mediated resistance of their hosts. J. Evol. Biol. 2011, 24, 741–750. [Google Scholar] [CrossRef]
  219. Rouchet, R.; Vorburger, C. Experimental evolution of parasitoid infectivity on symbiont-protected hosts leads to the emergence of genotype-specificity. Evolution 2014, 68, 1607–1616. [Google Scholar] [CrossRef] [Green Version]
  220. Dennis, A.B.; Patel, V.; Oliver, K.M.; Vorburger, C. Parasitoid gene expression changes after adaptation to symbiont-protected hosts. Evolution 2017, 71, 2599–2617. [Google Scholar] [CrossRef] [Green Version]
  221. Zepeda-Paulo, F.; Villegas, C.; Lavandero, B. Host genotype-endosymbiont associations and their relationship with aphid parasitism at the field level. Ecol. Entomol. 2017, 42, 86–95. [Google Scholar] [CrossRef]
  222. Li, S.; Liu, D.; Zhang, R.; Zhai, Y.; Huang, X.; Wang, D.; Shi, X. Effects of a presumably protective endosymbiont on life-history characters and their plasticity for its host aphid on three plants. Ecol. Evol. 2018, 8, 13004–13013. [Google Scholar] [CrossRef]
  223. Martinez, A.J.; Weldon, S.R.; Oliver, K.M. Effects of parasitism on aphid nutritional and protective symbioses. Mol. Ecol. 2014, 23, 1594–1607. [Google Scholar] [CrossRef]
  224. McLean, A.H.C.; Hrček, J.A.N.; Parker, B.J.; Godfray, H.C.J. Cascading effects of herbivore protective symbionts on hyperparasitoids. Ecol. Entomol. 2017, 42, 601–609. [Google Scholar] [CrossRef] [Green Version]
  225. Enders, L.S.; Hefley, T.J.; Girvin, J.J.; Whitworth, R.J.; Smith, C.M. Spatiotemporal distribution and environmental drivers of Barley yellow dwarf virus and vector abundance in Kansas. Phytopathology 2018, 108, 1196–1205. [Google Scholar] [CrossRef] [PubMed]
  226. Thierry, H.; Monteil, C.; Parry, H.; Vialatte, A. Simulating seasonal drivers of aphid dynamics to explore agronomic scenarios. Ecosphere 2021, 12, e03533. [Google Scholar] [CrossRef]
  227. Klueken, A.M.; Hau, B.; Ulber, B.; Poehling, H.M. Forecasting migration of cereal aphids (Hemiptera: Aphididae) in autumn and spring. J. Appl. Entomol. 2009, 133, 328–344. [Google Scholar] [CrossRef]
  228. Moreno-Delafuente, A.; Viñuela, E.; Fereres, A.; Medina, P.; Trębicki, P. Simultaneous increase in CO2 and temperature alters wheat growth and aphid performance differently depending on virus infection. Insects 2020, 11, 459. [Google Scholar] [CrossRef]
  229. Tamburini, G.; van Gils, S.; Kos, M.; van der Putten, W.; Marini, L. Drought and soil fertility modify fertilization effects on aphid performance in wheat. Basic Appl. Ecol. 2018, 30, 23–31. [Google Scholar] [CrossRef]
  230. Aqueel, M.A.; Leather, S.R. Effect of nitrogen fertilizer on the growth and survival of Rhopalosiphum padi (L.) and Sitobion avenae (F.) (Homoptera: Aphididae) on different wheat cultivars. Crop Prot. 2011, 30, 216–221. [Google Scholar] [CrossRef]
  231. Hambäck, P.A.; Vogt, M.; Tscharntke, T.; Thies, C.; Englund, G. Top-down and bottom-up effects on the spatiotemporal dynamics of cereal aphids: Testing scaling theory for local density. Oikos 2007, 116, 1995–2006. [Google Scholar] [CrossRef]
  232. Borer, E.T.; Seabloom, E.W.; Mitchell, C.E.; Power, A.G. Local context drives infection of grasses by vector-borne generalist viruses. Ecol. Lett. 2010, 13, 810–818. [Google Scholar] [CrossRef]
  233. Faheem, M.; Sajjad, A.; Shafique, R.M. Balanced use of fertilizers can reduce aphid infestation and improve yield in wheat crop. Asian J. Agric. Biol. 2015, 3, 50–55. [Google Scholar]
  234. Gu, S.; Zalucki, M.P.; Men, X.; Li, J.; Hou, R.; Zhang, Q.; Ge, F.; Ouyang, F. Organic fertilizer amendment promotes wheat resistance to herbivory and biocontrol services via bottom-up effects in agroecosystems. J. Pest Sci. 2022, 95, 339–350. [Google Scholar] [CrossRef]
  235. Aqueel, M.A.A.; Collins, C.M.; Raza, A.M.R.; Ahmad, S.; Tariq, M.; Leather, S.R. Effect of plant nutrition on aphid size, prey consumption, and life history characteristics of green lacewing. Insect Sci. 2013, 21, 74–82. [Google Scholar] [CrossRef] [PubMed]
  236. Hondelmann, P.; Poehling, H.M. Diapause and overwintering of the hoverfly Episyrphus balteatus. Entomol. Exp. Appl. 2007, 124, 189–200. [Google Scholar] [CrossRef]
  237. Zhao, Z.H.; Hui, C.; Hardev, S.; Ouyang, F.; Dong, Z.; Ge, F. Responses of cereal aphids and their parasitic wasps to landscape complexity. J. Econ. Entomol. 2014, 107, 630–637. [Google Scholar] [CrossRef] [Green Version]
  238. Zhao, Z.H.; Hui, C.; He, D.H.; Li, B.L. Effects of agricultural intensification on ability of natural enemies to control aphids. Sci. Rep. 2015, 5, 7. [Google Scholar] [CrossRef] [Green Version]
  239. Aqueel, M.A.; Raza, A.B.M.; Balal, R.M.; Shahid, M.A.; Mustafa, I.; Javaid, M.M.; Leather, S.R. Tritrophic interactions between parasitoids and cereal aphids are mediated by nitrogen fertilizer. Insect Sci. 2015, 22, 813–820. [Google Scholar] [CrossRef]
  240. Winqvist, C.; Bengtsson, J.; Aavik, T.; Berendse, F.; Clement, L.W.; Eggers, S.; Fischer, C.; Flohre, A.; Geiger, F.; Liira, J.; et al. Mixed effects of organic farming and landscape complexity on farmland biodiversity and biological control potential across Europe. J. Appl. Ecol. 2011, 48, 570–579. [Google Scholar] [CrossRef]
  241. Duan, X.; Pan, S.; Fan, M.; Chu, B.; Ma, Z.; Gao, F.; Zhao, Z. Cultivar mixture enhances crop yield by decreasing aphids. Agronomy 2022, 12, 335. [Google Scholar] [CrossRef]
  242. Batyrshina, Z.S.; Cna’ani, A.; Rozenberg, T.; Seifan, M.; Tzin, V. The combined impacts of wheat spatial position and phenology on cereal aphid abundance. PeerJ 2020, 8, e9142. [Google Scholar] [CrossRef]
  243. Gilabert, A.; Gauffre, B.; Parisey, N.; Gallic, J.F.L.; Lhomme, P.; Bretagnolle, V.; Dedryver, C.A.; Baudry, J.; Plantegenest, M. Influence of the surrounding landscape on the colonization rate of cereal aphids and phytovirus transmission in autumn. J. Pest Sci. 2017, 90, 447–457. [Google Scholar] [CrossRef]
  244. Török, E.; Zieger, S.; Rosenthal, J.; Földesi, R.; Gallé, R.; Tscharntke, T.; Batáry, P. Organic farming supports lower pest infestation, but less natural enemies than flower strips. J. Appl. Ecol. 2021, 58, 2277–2286. [Google Scholar] [CrossRef]
  245. Bischoff, A.; Pollier, A.; Tricault, Y.; Plantegenest, M.; Chauvel, B.; Franck, P.; Gardarin, A. A multi-site experiment to test biocontrol effects of wildflower strips in different French climate zones. Basic Appl. Ecol. 2022, 62, 33–44. [Google Scholar] [CrossRef]
  246. Bettaibi, K.; Mezghani-Khémakhem, M.; Bouktila, D.; Charaabi, K.; Raboudi, F.; Makni, H.; Maknim, M. A novel method for molecular targeting of insecticide resistance in Rhopalosiphum padi L. (Homoptera: Aphididae). Int. J. Pest Manag. 2016, 62, 284–287. [Google Scholar] [CrossRef]
  247. Wang, K.; Bai, J.; Zhao, J.; Su, S.; Liu, L.; Han, Z.; Chen, M. Super-kdr mutation M918L and multiple cytochrome P450s associated with the resistance of Rhopalosiphum padi to pyrethroid. Pest Manag. Sci. 2020, 76, 2809–2817. [Google Scholar] [CrossRef]
  248. Walsh, L.E.; Schmidt, O.; Foster, S.P.; Varis, C.; Grant, J.; Malloch, G.L.; Gaffney, M.T. Evaluating the impact of pyrethroid insecticide resistance on reproductive fitness in Sitobion avenae. Ann. Appl. Biol. 2021, 180, 361–370. [Google Scholar] [CrossRef]
  249. Foster, S.P.; Paul, V.L.; Slater, R.; Warren, A.; Denholm, I.; Field, L.M.; Williamson, M.S. A mutation (L1014F) in the voltage-gated sodium channel of the grain aphid, Sitobion avenae, is associated with resistance to pyrethroid insecticides. Pest Manag. Sci. 2013, 70, 1249–1253. [Google Scholar] [CrossRef]
  250. Yahya, M.; Saeed, N.A.; Nadeem, S.; Hamed, M.; Shokat, S. Role of wheat varieties and insecticide applications against aphids for better wheat crop harvest. Pakistan J. Zool. 2017, 49, 2217–2225. [Google Scholar] [CrossRef] [Green Version]
  251. Mohammed, A.A.A.H.; Desneux, N.; Fan, Y.; Han, P.; Ali, A.; Song, D.; Gao, X.W. Impact of imidacloprid and natural enemies on cereal aphids: Integration or ecosystem service disruption? Entomol. Gen. 2018, 37, 47–61. [Google Scholar] [CrossRef] [Green Version]
  252. Janssen, A.; van Rijn, P.C.J. Pesticides do not significantly reduce arthropod pest densities in the presence of natural enemies. Ecol. Lett. 2021, 24, 2010–2024. [Google Scholar] [CrossRef]
  253. Chen, G.; Hinds, J.; Zobel, E.; Rosario-Lebron, A.; Hooks, C.R.R. Evaluation of prophylactic sprays on pest abundance, foliar damage and yield in winter wheat. Int. J. Pest Manag. 2015, 61, 161–170. [Google Scholar] [CrossRef]
  254. Zuo, Y.; Wang, K.; Zhang, M.; Peng, X.; Piñero, J.C.; Chen, M. Regional susceptibilities of Rhopalosiphum padi (Hemiptera: Aphididae) to ten insecticides. Fla. Entomol. 2016, 99, 269–275. [Google Scholar] [CrossRef]
  255. Cao, J.Y.; Xing, K.; Zhao, F. Complex delayed and transgenerational effects driven by the interaction of heat and insecticide in the maternal generation of the wheat aphid, Sitobion avenae. Pest Manag. Sci. 2021, 77, 4453–4461. [Google Scholar] [CrossRef] [PubMed]
  256. Roschewitz, I.; Hücker, M.; Tscharntke, T.; Thies, C. The influence of landscape context and farming practices on parasitism of cereal aphids. Agric. Ecosyst. Environ. 2005, 108, 218–227. [Google Scholar] [CrossRef]
  257. Caballero-López, B.; Bommarco, R.; Blanco-Moreno, J.M.; Sans, F.X.; Pujade-Villar, J.; Rundlöf, M.; Smith, H.G. Aphids and their natural enemies are differently affected by habitat features at local and landscape scales. Biol. Control 2012, 63, 222–229. [Google Scholar] [CrossRef]
  258. Jonsson, M.; Buckley, H.L.; Case, B.S.; Wratten, S.D.; Hale, R.J.; Didham, R.K. Agricultural intensification drives landscape–context effects on host–parasitoid interactions in agroecosystems. J. Appl. Ecol. 2012, 49, 706–714. [Google Scholar] [CrossRef]
Table 1. Cereal aphid parasitoids (Hymenoptera: Braconidae: Aphidiinae) in Europe.
Table 1. Cereal aphid parasitoids (Hymenoptera: Braconidae: Aphidiinae) in Europe.
Aphidiinae SpeciesAphid HostsReferences
Aclitus obscuripennis FörsterAnoecia sp.[9]
Adialytus ambiguus (Haliday)Sipha maydis Passerini, Sipha elegans del Guercio[10,43,44]
Aphidius avenae HalidayMetopolophium dirhodum (Walker), Rhopalosiphum padi (L.), Schizaphis graminum (Rondani), Sitobion avenae (F.), Sitobion fragariae (Walker)[9,10,44]
Aphidius colemani ViereckR. padi, S. avenae[9,10,44]
Aphidius ervi HalidayDiuraphis noxia (Kurdjumov), M. dirhodum,
S. graminum, Rhopalosiphum maidis (Fitch),
R. padi, S. avenae, S. fragariae
Aphidius matricariae (Haliday)D. noxia, R. maidis, R. padi, S. graminum, S. avenae, S. fragariae[9,34,44,45]
Aphidius rhopalosiphi De StefaniD. noxia, M. dirhodum, Metopolophium festucae (Theobald), R. maidis, R. padi, S. graminum,
S. avenae, S. fragariae
Aphidius uzbekistanicus LuzhetzkiM. dirhodum, M. festucae, S. avenae, S. fragariae,
S. graminum
Binodoxys angelicae (Haliday)S. avenae[9]
Diaeretiella rapae (M’Intosh)D. noxia, R. maidis, R. padi, S. avenae, S. graminum[9,10,34,44,45]
Ephedrus plagiator (Nees)Anoecia sp., M. dirhodum, R. maidis, R. padi,
S. graminum, S. avenae, S. fragariae
Lipolexis gracilis FörsterR. padi[9]
Lipolexis labialis Tomanović and KocićAnoecia corni (F.)[46]
Lysiphlebus dissolutus (Nees)A. corni[9]
Lysiphlebus fabarum (Marshall)R. maidis, S. avenae[9,10,44]
Lysiphlebus testaceipes (Cresson)R. maidis, S. graminum, R. padi, S. avenae[10,34,44]
Monoctonus caricis (Haliday)R. padi, Sitobion sp.[47]
Paralipsis brachycaudi Tomanović and StarýTetraneura ulmi (L.)[48]
Paralipsis enervis (Nees)Geoica utricularia (Passerini)[48]
Praon abjectum (Haliday)R. padi[9]
Praon gallicum StarýM. dirhodum, R. padi, S. graminum, S. avenae[9,10,44]
Praon necans MackauerR. padi[9]
Praon volucre (Haliday)M. dirhodum, R. padi, S. graminum, S. avenae,
S. fragariae
Toxares deltiger (Haliday)M. dirhodum, S. avenae[9,31]
Trioxys auctus (Haliday)R. padi[9,49]
Trioxys sunnysidensis Fulbright and PikeR. padi[50]
Table 2. Literature review of hyperparasitoids of cereal aphid parasitoid in European agroecosystems. Kend denotes koinobiont endohyperparasitoids. Iect denotes idiobiont ectohyperparasitoids.
Table 2. Literature review of hyperparasitoids of cereal aphid parasitoid in European agroecosystems. Kend denotes koinobiont endohyperparasitoids. Iect denotes idiobiont ectohyperparasitoids.
FamilyHyperparasitoid SpeciesAphids Hosting Parasitized AphidiinesBiologyReferences
EncyrtidaeSyrphophagus aphidivorus (Mayr)M. dirhodum, R. padi, Rhopalosiphum sp., S. graminum,
S. avenae
Syrphophagus mamitus (Walker) Kend[49]
FigitidaeAlloxysta arcuata (Kieffer)R. padiKend[52]
Alloxysta brachyptera (Hartig)S. avenaeKend[51]
Alloxysta brevis (Thomson)M. dirhodum, R. maidis, R. padi,
S. avenae, Sipha spp.
Alloxysta castanea (Hartig)S. avenae, Hyalopterus pruni (Geoffroy), Hyalopterus sp.Kend[53,52]
Alloxysta fracticornis (Thomson)S. avenaeKend[52]
Alloxysta fulviceps (Curtis) Kend[37]
Alloxysta kovilovica Ferrer-Suay and Pujade-VillarS. avenaeKend[52]
Alloxysta leunisii (Hartig) Kend[49]
Alloxysta macrophadna (Hartig) Kend[49]
Alloxysta mullensis (Cameron)Sipha sp., S. avenaeKend[53,52]
Alloxysta pedestris (Curtis) Kend[37]
Alloxysta victrix (Westwood)M. dirhodum, R. padi, Rhopalosiphum sp., S. graminum,
S. avenae, Sitobion sp.
Phaenoglyphis villosa (Hartig)M. dirhodum, R. padi,
Rhopalosiphum sp., S. avenae,
H. pruni, Hyalopterus sp.
MegaspillidaeDendrocerus aphidum (Rondani)S. avenaeIect[51,55]
Dendrocerus carpenteri (Curtis)M. dirhodum, R. padi, Rhopalosiphum sp., S. graminum, Sipha sp., S. avenaeIect[12,51,55]
Dendrocerus laticeps (Hedicke) Iect[49]
Dendrocerus rectangularis (Kieffer) Iect[49]
Dendrocerus serricornis (Boheman)S. avenaeIect[55]
PteromalidaeAsaphes suspensus (Nees)M. dirhodum, R. padi, Rhopalosiphum sp., S. graminum,
S. avenae
Asaphes vulgaris WalkerM. dirhodum, Rhopalosiphum sp.,
S. avenae
Coruna clavata WalkerS. avenaeIect[37,49]
Pachyneuron aphidis (Bouché)S. avenae, M. dirhodumIect[12,37]
Pachyneuron concolor (Förster)M. dirhodum, S. avenaeIect[12]
Pachyneuron formosum WalkerS. avenaeIect[37,49]
Pachyneuron muscarum (L.)R. maidis, S. avenaeIect[37,51]
Pachyneuron solitarium (Hartig)S. avenaeIect[37]
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Tomanović, Ž.; Kavallieratos, N.G.; Ye, Z.; Nika, E.P.; Petrović, A.; Vollhardt, I.M.G.; Vorburger, C. Cereal Aphid Parasitoids in Europe (Hymenoptera: Braconidae: Aphidiinae): Taxonomy, Biodiversity, and Ecology. Insects 2022, 13, 1142.

AMA Style

Tomanović Ž, Kavallieratos NG, Ye Z, Nika EP, Petrović A, Vollhardt IMG, Vorburger C. Cereal Aphid Parasitoids in Europe (Hymenoptera: Braconidae: Aphidiinae): Taxonomy, Biodiversity, and Ecology. Insects. 2022; 13(12):1142.

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Tomanović, Željko, Nickolas G. Kavallieratos, Zhengpei Ye, Erifili P. Nika, Andjeljko Petrović, Ines M. G. Vollhardt, and Christoph Vorburger. 2022. "Cereal Aphid Parasitoids in Europe (Hymenoptera: Braconidae: Aphidiinae): Taxonomy, Biodiversity, and Ecology" Insects 13, no. 12: 1142.

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