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Review

Cladosporium—Insect Relationships

by
Rosario Nicoletti
1,2,*,
Elia Russo
2 and
Andrea Becchimanzi
2,3
1
Council for Agricultural Research and Economics, Research Center for Olive, Fruit and Citrus Crops, 81100 Caserta, Italy
2
Department of Agricultural Sciences, University of Naples Federico II, 80055 Portici, Italy
3
BAT Center—Interuniversity Center for Studies on Bioinspired Agro-Environmental Technology, University of Naples Federico II, 80055 Portici, Italy
*
Author to whom correspondence should be addressed.
J. Fungi 2024, 10(1), 78; https://doi.org/10.3390/jof10010078
Submission received: 14 December 2023 / Revised: 10 January 2024 / Accepted: 17 January 2024 / Published: 19 January 2024
(This article belongs to the Special Issue Exploring the Fascinating World of Fungal Symbioses)
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Abstract

:
The range of interactions between Cladosporium, a ubiquitous fungal genus, and insects, a class including about 60% of the animal species, is extremely diverse. The broad case history of antagonism and mutualism connecting Cladosporium and insects is reviewed in this paper based on the examination of the available literature. Certain strains establish direct interactions with pests or beneficial insects or indirectly influence them through their endophytic development in plants. Entomopathogenicity is often connected to the production of toxic secondary metabolites, although there is a case where these compounds have been reported to favor pollinator attraction, suggesting an important role in angiosperm reproduction. Other relationships include mycophagy, which, on the other hand, may reflect an ecological advantage for these extremely adaptable fungi using insects as carriers for spreading in the environment. Several Cladosporium species colonize insect structures, such as galleries of ambrosia beetles, leaf rolls of attelabid weevils and galls formed by cecidomyid midges, playing a still uncertain symbiotic role. Finally, the occurrence of Cladosporium in the gut of several insect species has intriguing implications for pest management, also considering that some strains have proven to be able to degrade insecticides. These interactions especially deserve further investigation to understand the impact of these fungi on pest control measures and strategies to preserve beneficial insects.

1. Introduction

Fungi in the genus Cladosporium (Dothideomycetes, Cladosporiaceae) are ubiquitous and reported from any terrestrial and marine substrate, including all kinds of living organisms [1]. This is linked to their profuse sporulation, which allows the spread of conidia through atmospheric agents over long distances. Thus, the mere isolation of these fungi from plants and animals does not necessarily imply a symbiotic association. However, generality is not a rule, and in many cases, this pervasiveness subtends either occasional or more systematic biotic relationships that influence the fitness of the associated organisms in diverse ways [2,3,4].
The range of interactions is particularly broad in the case of insects, with a various case history of antagonism and mutualism described so far. The available information on the relationships between Cladosporium and insects is examined in this paper, with the aim of shedding light on their biological/ecological assumptions, as well as on the ways and circumstances by which they may affect pest control strategies.

2. Taxonomic Aspects and Occurrence

Classification of fungi of the genus Cladosporium is problematic because of the infrequency of the teleomorphic stage and the absence of outstanding morphological differences in the conidial structures. Indeed, culturing and microscope observations only allowed a poorly rigorous separation of taxa, some of which are assumed to represent collective species, or ‘species complexes’ (s.c.); namely, C. cladosporioides, C. herbarum and C. sphaerospermum. The introduction of biomolecular tools in fungal taxonomy has enabled mycologists to resolve these aggregates with the separation of new entities so that as many as 169 species were recognized in the fundamental revision by Bensch et al. [1]. However, this number is continuously increasing following the finding and classification of novel isolates from all sorts of substrates and environmental contexts.
In this paper, we consider the findings of Cladosporium according to the updated nomenclature as far as possible. In fact, studies reporting species identification based on the sequencing of DNA markers are still a minority, and most of them only consider rDNA-ITS sequences, which have proved to be insufficient for a correct classification [1]. In this respect, the approximate phylogenetic reconstruction provided in some reports (e.g., reference [5]) significantly illustrates how identifications based on ITS only are merely tentative.
Data on the occurrence of Cladosporium species in association with insects resulting from the examination of the available literature are summarized in Table 1. In total, 303 entries, corresponding to the finding of these fungi on 171 insect species belonging to 16 orders, are included in this long list. On the fungus side, the species was not identified for about 46% of the entries (Cladosporium sp.), confirming classification to be quite a problematic aspect within this genus. When identification was accomplished, a total of 32 Cladosporium species were recognized. Among them, the three classic species mentioned above stand out in numerical terms; particularly, there are 56 reports for C. cladosporioides from 55 insect taxa belonging to 11 orders. This is not surprising considering that this is a polymorphic s.c. in which the existence of additional cryptic species has been conjectured [6]. Hence, at least in part, these identifications should be revised with reference to the updated nomenclature and the use of valid DNA markers. Indeed, among the reported identifications at the species level, as little as 11 (underlined in Table 1) are reliably based on the DNA markers officially considered in Cladosporium taxonomy [1,2]; for the time being, this limited insight does not allow us to advance hypotheses on any definite relationships between Cladosporium and insect species. The prevalence of findings concerning Hemiptera and Coleoptera is evident (Figure 1), in clear connection with their higher agricultural relevance.
In addition to C. fulvum, which has been reclassified as Fulvia fulva since a long time [171], the species C. chlorocephalum, cited in references [18,98,172,173], has not been included since it is now synonymized with Graphiopsis chlorocephala following a taxonomic revision [174]. Moreover, a few cases were disregarded because the reported identification of the insect-associated fungi was clearly wrong and misleading. As an example, an isolate from the gut of the locust Oxya chinensis (Orthoptera, Acrididae), claimed to be C. oxysporum based on a 5.8S rDNA sequence [175], rather belongs to a species of Curvularia; this emendation unequivocally results from both a blast of the sequence in the GenBank database and a visual examination of the conidia shown in the image provided in the published paper.
With reference to the geographical distribution, there are reports from as many as 54 countries in all continents, without any implication in terms of associations possibly depending on local conditions. Especially for small-sized species, the isolations have been mainly carried out from the whole body of the insects (Figure 2); however, there are many reports considering the gut and mouth parts in relation to the possible ingestion connected to feeding habits. In this respect, isolations from frass have been considered only when this material was obtained in the laboratory without any possible interference of environmental contamination (as in references [9,21,127,136]).
The various findings of Cladosporium in shelters, tunnels or other structures inhabited by insects have not been exhaustively considered, since in many instances they may derive from environmental contamination rather than direct interaction with the insects. This is the case of reports of C. cladosporioides as the most frequent fungus found in tunnels dug by larvae of the longicorn Rosalia alpina (Coleoptera, Cerambycidae) in wych elm (Ulmus glabra) trees at a Polish conservation site [176] and in mines of lepidopteran leafminers on black locusts (Robinia pseudacacia) [139]. Reports on the occurrence of Cladosporium in food supplies of social insects (e.g., bee bread [177]) and as undesired contaminants in laboratory assays or rearing diets of insects (e.g., reference [178]) have also been disregarded.
It must be pointed out that most entries of Cladosporium sp. in Table 1 refer to multiple concomitant isolations or detections in the insect samples, which may imply a broad species assortment. Particularly, this connotes the many metagenomic-based studies exploring the diversity of the mycobiome of insects in specific ecological contexts. In this regard, an investigation on fungi associated with the olive fruit fly (Bactrocera oleae: Diptera, Tephritidae) in Calabria, Southern Italy, disclosed a striking dominance of Cladosporium, matching about 80% of OTU sequences. More in detail, this set was dominated by members of the C. cladosporioides s.c., while the C. herbarum s.c., C. velox and a couple of unidentified Cladosporium spp. were much less frequent [107,179]. In a similar study concerning the congeneric Queensland fruit fly (Bactrocera tryoni), Cladosporium were again among the dominant fungi in the gut mycobiome, with a higher frequency in females [108]. More recently, a systematic investigation has been carried out in the olive ecosystems of Tunisia to study the microbiome associated with this crop. However, the taxonomic remarks are reported as collective data considering the occurrence of fungi in soil and in association with insect pests, which does not allow us to infer specific associations with the latter [180]. Likewise, in an investigation concerning plant-feeding true bugs (Heteroptera: Cimicomorpha and Pentatomomorpha) carried out in China, data were summarized with reference to the family, not allowing us to infer the associations of Cladosporium with every single bug species [181].
Another broad-scale investigation carried out in a Canadian aspen forest showed that Cladosporium were the most frequent fungi associated with arthropods; again, C. cladosporioides was the most common species, accounting for 77% of the isolates ascribed to this genus, followed by C. sphaerospermum at about 20%, while C. herbarum and C. orchidis were occasional. Unfortunately, no details on the insect species representing the isolation sources were provided in this study [182].

3. Entomopathogenicity

Conventionally, Cladosporium species are not considered full-right representatives of the guild of entomopathogens, which is generally restricted to specialized fungi such as Beauveria, Metarhizium and Lecanicillium/Akanthomyces [183,184,185]. However, like other fungi that are widely associated with crops such as Trichoderma and Talaromyces [186,187], the evidence is increasing that Cladosporium may also infect insects and cause epizootics in pest populations or promote plant defense reactions.
Direct observations of the parasitic aptitude of insects are limited and essentially concern the case of C. cladosporioides on the sugarcane white wooly aphid (Ceratovacuna lanigera: Hemiptera, Aphididae); both light and electron microscopy at the host–parasite interface showed that nymphs and adults of the aphid were completely overgrown by the fungal mycelium, which penetrated and disrupted their powdery waxy coating [5].
However, circumstantial evidence of entomopathogenic aptitude in Cladosporium derives from several studies reporting on mortality induced by conidial suspensions administered at various concentrations and exposure times. In this regard, the available data concerning strains that proved to be effective against various targeted pests in experimental trials are summarized in Table 2.
Alternatively, the anti-insectan effect can be assessed through the addition of the fungi or their products to the laboratory diet. In this respect, when incorporated in the feed of larvae of the tobacco budworm (Chloridea virescens: Lepidoptera, Noctuidae), an isolate of C. cladosporioides was found to reduce larval and pupal weights by 56% and 7%, respectively; moreover, in preference tests, the caterpillars showed a marked tendency to avoid feed amended with the fungus [195]. Development of another noctuid moth, the tobacco cutworm (Spodoptera litura), was significantly prolonged when larvae were fed on a diet amended with ethyl acetate extract of C. uredinicola at concentrations of 1.25–2.00 μL g−1; moreover, at 2.00 μL g−1, a significantly higher number of adults emerged showing morphological deformities. At higher concentrations, significant reductions in adult emergence, longevity and reproductive potential were recorded. Finally, the toxicity of the ethyl acetate extract was further evidenced by a reduction in feed utilization by the larvae [196].
The ethyl acetate and methylene chloride extracts of a strain of C. cladosporioides were effective against nymphs and adults of the cotton aphid (Aphis gossypii: Hemiptera, Aphididae) [13,197]. Aphicidal effect was also displayed by formulations based on emulsions of culture filtrates of an endophytic strain of C. oxysporum endowed with proteolytic activity, which were more active than conidial suspensions against the black bean aphid (Aphis fabae: Hemiptera, Aphididae) [190]. In a subsequent experiment, formulations based on culture filtrates of this strain and two more endophytic isolates of C. echinulatum and Cladosporium sp. showed activity against the green peach aphid (Myzus persicae: Hemiptera, Aphididae), which increased at increasing concentrations. A significant reduction in the number of colonizing aphids and a relative increase in the number of winged adults were recorded. Moreover, the pretreatment of plants negatively influenced embryonic development, thus affecting fertility [198]. In the same study, consistent chitinolytic activity was determined in the culture filtrates of Cladosporium sp.; indeed, chitinases are considered a main factor in the bioactivity of fungal culture filtrates, as also documented for other strains of Cladosporium spp. [193], C. cladosporioides [27,48], C. tenuissimum and C. xanthocromaticum [48].
Even more, the anti-insectan effects of culture filtrates may depend on the presence of toxic compounds (Figure 3). Fungi in the genus Cladosporium are known as prolific producers of bioactive secondary metabolites [199], some of which have been detected as possible determinants of detrimental effects on insects. This is the case of bassianolide, a cycloligomer depsipeptide identified as a product of a strain related to the C. cladosporioides s.c. [200]. The alkaloid 3-(4β-hydroxy-6-pyranonyl)-5-isopropylpyrrolidin-2-one was identified in the ethyl acetate extracts of another strain of C. cladosporioides displaying aphicidal activity [13]. Another alkaloid, hydroxyquinoline, was identified as the potentially active product in the extracts of a strain of C. subuliforme [167]. The novel compound citreoviridin A was extracted from an isolate of C. herbarum from a marine sponge and found to inhibit the growth of larvae of the cotton leafworm (Spodoptera littoralis: Lepidoptera, Noctuidae) [201]. Chlorogenic acid, purified from an endophytic isolate of C. velox, displayed insecticidal activity by inducing significant mortality in the larvae of S. litura or adversely prolonging their developmental period. This phenolic compound, previously known to cause gut toxicity in lepidopterans [202], was characterized as an α-glucosidase inhibitor, performing a non-competitive type of inhibition in vitro; it also inhibited the activity of α-glycosidases in the gut of the larvae [203,204].
The importance of secondary metabolites for entomopathogenic aptitude in Cladosporium has been further affirmed after a study carried out on strains associated with the Chinese white wax scale (Ericerus pela: Hemiptera, Coccidae). This insect is known to be infected by Cladosporium spp. related to C. sphaerospermum and C. langeronii, which kill the scales after dramatically altering their microbiome [34]. However, the scales were later found to also harbor a non-infective Cladosporium. Genome sequencing showed that the non-infective strain is related to C. cladosporioides and has a larger genome size than a pathogenic one, which is more related to C. sphaerospermum. Particularly, the former has specific genes involved in nutrition pathways that are absent in the pathogen. Conversely, the latter possesses genes participating in the biosynthetic pathways of mycotoxins, such as asperfuranone, emericellamide and fumagillin. These genes were not found in the nonpathogenic strain, which, on the other hand, presented genes associated with reduced virulence [3].

3.1. Interactions with Biocontrol Agents

Reports on the occurrence of an association with predatory and parasitoid insects introduce the question of whether the insecticidal properties of Cladosporium may also affect the performances of biocontrol agents employed in crop protection. Indeed, this association can be more than merely occasional, considering that Cladosporium were the most abundant fungi detected in the gut of the multicolored Asian lady beetle (Harmonia axyridis: Coleoptera, Coccinellidae) feeding on the pea aphid (Acyrthosiphon pisum: Hemiptera, Aphididae) [129]. Concerning this issue, a previously mentioned strain of Cladosporium sp. from H. armigera was found not to induce significant harmful effects on a panel of beneficial predatory insects, including the red and blue beetle (Dicranolaius bellulus: Coleoptera, Melyridae), the transverse ladybird (Coccinella transversalis: Coleoptera, Coccinellidae), the green lacewing (Mallada signatus: Neuroptera, Chrysopidae) and the damsel bug (Nabis kinbergii: Hemiptera, Nabidae) [130]. Conversely, laboratory assays carried out in Egypt showed that treatment with C. uredinicola affected the biocontrol of the silverleaf whitefly (Bemisia tabaci: Hemiptera, Aleyrodidae) by the eleven-spotted ladybird (Coccinella undecimpunctata: Coleoptera, Coccinellidae) and the parasitoid Eretmocerus mundus (Hymenoptera, Aphelinidae) in various ways. In fact, all larval stages of the coccinellid were sensitive to the fungus and tended to avoid feeding on the infected whiteflies. As for the parasitoid, although mortality of the exposed individuals was low, most females avoided laying eggs on treated nymphs; nevertheless, the combined use of C. uredinicola and E. mundus was found to synergistically increase the suppression of nymphs [205].
Olfactory experiments carried out in the laboratory indicated that the parasitoid wasp Lysiphlebus fabarum (Hymenoptera, Braconidae) can detect cues from aphids (A. fabae) infected by a pathogenic strain of Cladosporium sp. and avoid them; hence, the employment of this strain in the field could not affect the performance of the parasitoid, implying compatibility between these, and possibly more, biological control agents of aphids [157].

3.2. Plant-Mediated Interactions

In addition to arising after direct contact or ingestion of conidia, the entomopathogenic effects of Cladosporium can also be exerted in planta, as promoted by strains able to develop endophytically. Indeed, it is known that endophytic fungi may improve plant resistance to biotic adversities through various mechanisms, including general effects on fitness and growth promotion eventually exerted in synergistic relationships with other components of the plant microbiome [206,207]. The belief is gaining ground that these valuable properties could be exploited for improving yields while reducing the input of chemicals in crop management [208,209].
Cauliflower plants artificially infected with an endophytic strain of C. uredinicola did not show any disease symptoms, and the vigor of endophyte-infected plants also did not differ from untreated plants. Interestingly, larvae of S. litura feeding on leaves from treated plants were sluggish and underwent significantly higher mortality than the control. Most of the larvae died at the time of molting to the last instar, while the survivors took a significantly longer time to pupate and further suffered significantly higher mortality at the pupal stage. In the end, fewer adults emerged from larvae on endophyte-supplemented plants; some adults exhibited morphological deformities, such as crumpled and unequal wings, and survived for a very short time. Inhibitory effects were also observed on the reproductive potential and the hatchability of eggs. The life span of females that emerged from larvae fed on plants hosting C. uredinicola reduced significantly, while male longevity remained unaffected [210]. All these effects were assumed to depend on physiological changes induced by the endophyte. In fact, further studies disclosed cytotoxic effects on hemocytes of S. litura fed on endophyte-supplemented cauliflower plants, which showed changes in shape, extensive vacuolization and necrosis. Moreover, these abnormalities increased along with the feeding duration and ultimately resulted in adverse consequences on the fitness and survival of the insect [211].
However, it is quite intuitive to consider that plant-mediated relationships should be examined case by case, as the outcome of the interaction is not necessarily unfavorable to the insects. When inoculated in perennial thistle (Cirsium arvense), where it is known to develop endophytically, C. cladosporioides increased feeding of the thistle tortoise beetle (Cassida rubiginosa: Coleoptera, Chrysomelidae), while it had no effect on the cabbage moth (Mamestra brassicae: Lepidoptera, Noctuidae). Nevertheless, dual infection with C. cladosporioides and Trichoderma viride greatly reduced beetle feeding [212]. These findings indicate that the promoting effects of C. cladosporioides, as well as of other endophytes, depend on both the degree of specialization of the herbivore and the species assortment in the plant microbiome, which in turn may induce chemical changes in the host. Undoubtedly, these fungi deserve higher attention in the study of insect–plant interactions, considering that their endophytic occurrence could remarkably influence insect growth and even pest population dynamics.

4. Other Ecological Relationships

Mycophagy somehow represents the reverse condition of entomopathogenicity, in which insects perform a suppressive role on Cladosporium by feeding on the mycelium. However, this relationship may still imply an ecological advantage for the fungus, deriving from the use of insects as carriers for its propagation [213]. Springtails (Collembola) are especially known for feeding on soil fungi, including Cladosporium [214,215], and C. cladosporioides has been used as feed to preserve laboratory stocks of the species Hypogastrura tullbergi (Poduromorpha, Hypogastruridae) and Proisotoma minuta (Entomobryomorpha, Isotomidae) [216]. This species was also found to support the development of the minute brown scavenger beetle (Dienerella argus: Coleoptera, Latridiidae) [217]; moreover, it is part of the diet of the sap beetle Brachypeplus glaber (Coleoptera, Nitidulidae), as demonstrated by gut content analyses and observations of adult and larval feeding [19].
Several insect pests of stored grains belonging to different orders and families have been reported to feed and even reproduce on Cladosporium grown in axenic cultures. More in detail, a strain of C. cladosporioides was found to support the development of the reticulate-winged booklouse (Lepinotus reticulatus: Psocoptera, Atropidae) and a series of Coleoptera, including the narrownecked grain beetle (Anthicus floralis: Anthicidae), the sigmoid fungus beetle (Cryptophagus varus: Cryptophagidae), the lesser grain borer (Rhyzopertha dominica: Bostrichidae), the wheat weevil (Sitophilus granarius: Curculionidae), the bean weevil (Acanthoscelides obtectus: Bruchidae), the larger black flour beetle (Cynaeus angustus: Tenebrionidae), the square-nosed fungus beetle (Lathridius minutus) and Microgramma arga (Lathrididae) [218]. One of them, C. angustus, was even attracted by fresh-milled corn flour amended with Cladosporium [219]. Moreover, the ability to develop in vitro on pure cultures of an unidentified Cladosporium strain was reported for the foreign grain beetle (Ahasversus advena: Coleoptera, Sylvanidae), which females also demonstrated to prefer Cladosporium in oviposition tests [220].
Some contrast is evident in reports concerning the interaction of C. cladosporioides with R. dominica. In fact, after the above supporting effect [218], induction of mortality has recently been documented in this borer insect [56]. This contradiction could be easily explained considering that different strains of the same species may have different biological properties, particularly with reference to secondary metabolite production. Moreover, after the recent adjustments in Cladosporium taxonomy, it is quite possible that, actually, these isolates belong to different species, which further underlines the importance of a correct identification. Rather than being a mere hypothesis, this inference was verified in the case of the Cladosporium associates of the fruit fly Drosophila suzukii (Diptera, Drosophilidae) in raspberries; in fact, it transpired that isolates preliminarily identified as C. cladosporioides belonged to at least two more species (C. anthropophilum and C. pseudocladosporioides) after a more accurate taxonomic identification based on biomolecular markers [9].
The ambrosia beetles (Coleoptera, Curculionidae, Scolytinae) are known to spread fungi that develop in the galleries they dig in the host trees [221]. Although experimental evidence leans for selected fungi, such as species of Fusarium, Geosmithia, Penicillium and Raffaelea, to be more systematically involved in these mutualistic relationships [36,52,222,223], Cladosporium is often isolated both from the insects and their galleries [52,94,133,134]. Particularly, isolation and identification based on biomolecular methods demonstrated the association of C. perangustum with Xylosandrus compactus after female beetles were found carrying the fungus on their body [77]; previously, Cladosporium had been regarded as the principal food source of this species [224].
Likewise, the coffee berry borer (Hypothenemus hampei: Coleoptera, Curculionidae) was found to be associated with C. oxysporum and an unidentified Cladosporium sp. [76]. Another unidentified Cladosporium sp. was found to systematically occur in leaf rolls inhabited by larvae of the weevil Euops lespedezae (Coleoptera, Attelabidae) on the leafy bush clover (Lespedeza cyrtobotrya); however, this fungus was not found in mycangia of the weevil females, indicating that it is not a vertically transmitted symbiont unlike other leaf roll-associated fungi [225]. Cladosporium spp. were also dominant in leaf rolls of another Attelabidae, Heterapoderopsis bicallosicollis, on the Chinese tallow tree (Triadica sebifera); this is considered not to be a specific association, as the authors hypothesize that these and other cellulolytic fungi, providing nutritional support to the larvae, colonize the leaf rolls once they fall to the soil [226].
In addition to the above findings, not surprisingly, our fungi have been reported to colonize galls formed on plants by insects of various taxonomic assortment. This is the case of aphids of the genus Pemphigus (Hemiptera, Pemphigidae), commonly associated with poplars (Populus spp.), in which galls C. cladosporioides and other unidentified Cladosporium spp. can be found among other fungi [227]. Moreover, C. sphaerospermum was found to frequently occur in galls of the eastern spruce adelgid (Adelges abietis: Hemiptera, Adelgidae), even if no circumstantial relationship could be documented [228]. Also very common is the occurrence of Cladosporium in galls formed on a multitude of plants by Asphondylia spp. and other midges in the Cecidomyidae (Diptera) [103,104,105,132,229]. Again, this association does not seem to entail any functional symbiotic relationships, considering that on Greek savory (Micromeria graeca), these fungi were also found in ungalled flower buds. Moreover, the isolates did not belong to a single species, which could have implied an ecological specialization; rather, they were representative of a wide taxonomic assortment, including two novel species [16].
Connected with the feeding aptitude is the isolation of Cladosporium spp. from the gut of larvae of aquatic shredders of the genera Phylloicus (Trichoptera, Calamoceratidae) and Stenochironomus (Diptera, Chironomidae); these strains displayed cellulolytic and xylanolytic properties in laboratory assays, supporting the hypothesis that they might improve the digestibility of leaves by the insects [63,73,150]. Likewise, Cladosporium spp. were reported as prevalent in the gut of males of the Asian tiger mosquito (Aedes albopictus: Diptera, Culicidae) and are considered to play a role in the assimilation of fructose, which is a relevant component in their nectarivorous diet [86]. As part of an extensive array of microorganisms that play a crucial role in the digestion of feed, in the absorption of nutrients and in the protection against pathogens, the occurrence of Cladosporium in the digestive tract has been documented in many unrelated insect species, both after direct isolation [3,15,19,24,26,47,50,51,58,59,78,79,100,101,118,125,127] and as a result of studies based on biochemical (e.g., denaturing gradient gel electrophoresis) and metagenomic analyses [3,40,47,88,90,101,107,108,109,117,118,125,133,138,141,144,146,151,154,161,230]. In addition to descriptive aims, these studies have addressed various aspects that more or less influence the gut microbiome species assortment, such as age, instar, gender, caste, diet, pesticides, antibiotics and various environmental factors. Increasing evidence indicates that disturbances in the gut microbiota can compromise the host’s health and that its diversity has a far-reaching impact on the insect’s fitness. This is also closely related to the issue of pesticide resistance; in fact, the capacity to degrade chlorpyrifos and endosulfan has been documented by strains of C. cladosporioides [231,232,233]. Indeed, a better understanding of the role played by these fungi and the interacting microbial communities in the insects’ gut is essential in view of increasing opportunities for their possible exploitation [230,234,235].
The helper role of Cladosporium towards pests can be reciprocated in the cases where insects act as vectors of plant pathogenic strains. In this respect, D. suzukii was found to act as a carrier of Cladosporium spp., enabling these fungi to proliferate in raspberries [9,127]. The dark-winged fungus gnat Bradysia impatiens (=B. agrestis: Diptera, Sciaridae) has also been reported to spread Cladosporium conidia [78]. Clearly, the possible epidemiological role of some insects as occasional vectors of plant pathogenic species must be taken into careful consideration, even if the impact of Cladosporium spp. as plant pathogens is generally lower than other insect-associated fungi such as Fusarium [236].
A particular kind of interaction with Cladosporium has been documented for two wasps (Dolichovespula sylvestris and Paravespula vulgaris: Hymenoptera, Vespidae) that act as pollinators for the orchid Epipactis helleborine. In fact, the wasps were found to spread conidia of Cladosporium while visiting the flowers; these fungi are presumed to produce ethanol contained in the nectar, which in turn is thought to attract the wasps [126]. This finding could be indicative of a more widespread and relevant role of Cladosporium in the interaction with pollen vectors for angiosperms.
Cypripedium fargesii is a nectarless endangered orchid that requires cross-pollination to produce the maximum number of viable embryos. The flat-footed fly (Agathomyia sp.: Diptera, Platypezidae) is used to visit flowers of this orchid, and individuals examined after entering or leaving the labellum sac were found to carry Cladosporium conidia on their legs and mouthparts, suggesting mycophagy. Quite intriguingly, the upper surface of foliage presents blackish hairy spots that mimic the mold spots of Cladosporium, thereby serving as visual lures. Moreover, the floral scent composition includes some volatile compounds, such as 3-methyl-1-butanol, 2-ethyl-1-hexanol and 1-hexanol, which have also been detected in Cladosporium cultures [89]. Based on these remarks, it can be inferred that the endophytic adaptation of Cladosporium has led this fungus to play a fundamental intermediary role in the reproduction of these plants, which deserves to be further investigated.

5. Conclusions

Within the diverse ecological relationships connecting insects and fungi [237], evidence from the available experimental data is mostly indicative of an antagonistic role of Cladosporium against insects deserving higher attention for practical exploitation in pest control. In this respect, the reported infectious aptitude of C. cladosporioides against spider mites [238,239] and the known mycoparasitic ability against several plant pathogens, such as rusts [240], candidate these fungi as multipurpose biocontrol agents. Clearly, this opportunity requires an assessment of whether these useful roles can be simultaneously performed by single selected strains. The recent observation that phylogenetically diverse species of Cladosporium display concurrent effects supports the conjecture that these functional traits are rooted in ancestry in this genus, providing a favorable indication in this respect [241].
A final aspect connected with the Cladosporium–insect association to be taken into careful consideration refers to the recent trend of introducing insects into the human diet [242,243]. Indeed, the high frequency with which these fungi can be found infecting or coating insects should be particularly monitored in the rearing conditions, which can be conducive to their spread and the ensuing possible mycotoxin contamination [199,244,245,246].

Author Contributions

Conceptualization, R.N.; resources, E.R. and A.B.; writing—original draft preparation, R.N.; writing—review and editing, R.N., E.R. and A.B. All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding.

Conflicts of Interest

The authors declare no conflicts of interest.

References

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Figure 1. Findings of insect-associated Cladosporium spp. grouped by insect orders.
Figure 1. Findings of insect-associated Cladosporium spp. grouped by insect orders.
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Figure 2. Isolation of Cladosporium from the pine tortoise scale (Toumeyella parvicornis).
Figure 2. Isolation of Cladosporium from the pine tortoise scale (Toumeyella parvicornis).
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Figure 3. Chemical structure of Cladosporium secondary metabolites displaying anti-insectan effects.
Figure 3. Chemical structure of Cladosporium secondary metabolites displaying anti-insectan effects.
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Table 1. Occurrence of Cladosporium in association with insect species.
Table 1. Occurrence of Cladosporium in association with insect species.
Cladosporium SpeciesInsect Species *LocationReference
C. aggregatocicatricatumDryocosmus kuriphilusMonti Cimini (Italy)[7]
Xylosandrus compactusCirceo Promontory (Italy)[8]
C. anthropophilumDrosophila suzukiiMaryland (USA)[9]
C. aphidisunidentified aphidPiracicaba (São Paulo, Brazil)
Berlin; Brandenburg (Germany)
Parma (Italy)
Wilmington (Delaware, USA)		[1]
Aphis gossypiiPuerto Rico
Aphis sp.Berlin (Germany)
Aphis symphytiKlosterneuburg (Austria)
Brevicoryne brassicaeAssiut (Egypt)[10]
Rhopalosiphum maidisHonolulu (Hawaii, USA)[1]
unidentified scaleEssen (Germany)
C. austrohemisphaericumX. compactusCirceo Promontory (Italy)[8]
C. chasmanthicolaSpodoptera frugiperdaKansas (USA)[11]
C. cladosporioidesAleurothrixus aepimBrazil[12]
Aphis craccivoraEgypt[13]
Apion ulicisChristchurch area (New Zealand)[14]
Apis melliferaAssiut Governorate (Egypt)[15]
Asphondylia micromeriae
Asphondylia nepetae		Isle of Vivara (Italy)
Averno; Matera (Italy)		[16]
Atta capiguara
Atta laevigata		Botucatu (São Paulo, Brazil)[17]
Bemisia spp.Dakahlia Governorate (Egypt)[18]
Brachypeplus glaberFlorida (USA)[19]
B. brassicaeDakahlia Governorate (Egypt)
Assiut (Egypt)		[20]
[10]
Ceratovacuna lanigeraAnakapalle (Andhra Pradesh, India)[5]
Chitaura brachypteraDumoga-Bone (Sulawesi, Indonesia)[21]
Chrysomphalus aonidumQualubia (Egypt)[22]
Chrysomya megacephalaLudhiana (Punjab, India)[23]
Chrysoperla rufilabrisMonroe; Oktibbeha (Mississippi, USA)[24]
unidentified lignicolous ColeopteraÅs (Norway)[25]
Coptotermes formosanusNew Orleans (Louisiana, USA)[26]
Culex pipiensQena Governorate (Egypt)[27]
Culex quinquefasciatusBasrah (Iraq)[28]
Cydia ulicetanaChristchurch area (New Zealand)[14]
Desoria albellaWarren Woods (Michigan, USA)[29]
Diaphorina citriFlorida (USA)[30]
unidentified DipteraCoimbra (Portugal)[31]
D. suzukiiMaryland (USA)
Geneva (New York, USA)		[9]
[32]
Dytiscus marginalisEast Anatolia (Turkey)[33]
Epiphyas postvittanaChristchurch area (New Zealand)[14]
Ericerus pelaKunming (China)[34]
Euphalerus clitoriaePernambuco (Brazil)[35]
Euwallacea interjectusHiroshima Prefecture (Japan)[36]
Galleria mellonellaNorthern China
Beijing (China)		[37]
[38]
Gromphadorhina portentosaColumbus (Ohio, USA)[39]
unspecified grasshoppersUlu-Endau (Malaysia)[21]
Hermetia illucensGiessen (Germany)[40]
Heteraphorura subtenuisAlberta (Canada)[41]
Homalodisca vitripennisSouthern California (USA)[42]
Hydrophilus piceusEast Anatolia (Turkey)[33]
unidentified dead insectThailand[43]
Ips sexdentatusNortheastern Ukraine[44]
Kermes sp.Guangdong (China)[45]
Lycorma delicatulaBerks County (Pennsylvania, USA)[46]
Megaplatypus mutatusBragado (Argentina)[47]
Mesambria maculipesDumoga-Bone (Sulawesi, Indonesia)[21]
Myzus persicaeSalah El-Din Governorate (Iraq)[48]
Nilaparvata lugensGazipur (Bangladesh)[49]
Odontotermes formosanusJinhua (China)[50]
Pantala flavescensJinhua (China)[51]
Pityogenes bidentatusBabimost; Mielec; Opole (Poland)[52]
Prays oleaeMirandela-Bragança region (Portugal)[53]
Pterostichus melanariusAssiut Governorate (Egypt)[15]
unidentified PyrrhocoridaeLebanon[54]
Rhynchophorus ferrugineusAssiut Governorate (Egypt)[15]
Scolytogenes birosimensisCentral and Western Japan[55]
Sericothrips staphylinusChristchurch area (New Zealand)[14]
Sitophilus oryzaeMultan (Pakistan)[56]
unidentified ThysanuraCoimbra (Portugal)[31]
Tomicus piniperdaMościska Forest (Poland)[57]
Triatoma brasiliensis
Triatoma pseudomaculata		Rio de Janeiro (Brazil)[58]
Triatoma infestansMendoza; Salta; Santa Fe (Argentina)[59]
Troglophilus neglectusTolmin (Slovenia)[60]
C. cucumerinumunidentified Diptera
unidentified Thysanura		Coimbra (Portugal)[31]
C. cycadicolaTribolium castaneumIksan (South Korea)[61]
C. delicatulumThrips sp.Hamedan (Iran)[62]
C. dominicanumX. compactusCirceo Promontory (Italy)[8]
C. endophyticumStenochironomus sp.Adolpho Ducke Reserve (Amazonas); Lajeado State Park (Tocantins, Brazil)[63]
C. exasperatumNasutitermes octopilisNouragues Nature Reserve (French Guiana)[64]
C. exileAphis sp.Somesara (Iran)[62]
C. halotoleransG. mellonellaNapoli area (Italy)[65]
Stenochironomus sp.Lajeado State Park (Tocantins, Brazil)[63]
Thitarodes xiaojinensisXiaojin (China)[66]
T. castaneumSangju (South Korea)[61]
C. herbarumAleurodicus cocoisRecife (Brazil)[67]
A. gossypiiLatvia[68]
A. ulicisChristchurch area (New Zealand)[14]
A. melliferaAssiut Governorate (Egypt)[15]
B. brassicaeAssiut (Egypt)[10]
C. ulicetanaChristchurch area (New Zealand)[14]
D. marginalisEast Anatolia (Turkey)[33]
E. postvittanaChristchurch area (New Zealand)[14]
H. subtenuisAlberta (Canada)[41]
H. piceusEast Anatolia (Turkey)[33]
Musca domesticaSeropedica (Rio de Janeiro, Brazil)[69]
Gauteng Province (South Africa)[70]
P. bidentatusBabimost (Poland)[52]
Pseudopsis subulataMontmorency Forest (Canada)[71]
S. staphylinusChristchurch area (New Zealand)[14]
Tenebrio molitorAarhus (Denmark)[72]
T. piniperdaMościska Forest (Poland)[57]
T. brasiliensis
T. pseudomaculata		Rio de Janeiro (Brazil)[58]
C. iranicumunidentified scaleIran[1]
C. kenpeggiiStenochironomus sp.Adolpho Ducke Reserve (Amazonas); Lajeado State Park (Tocantins, Brazil)[63]
Triplectides sp.Lajeado State Park (Tocantins, Brazil)[73]
C. langeroniiD. kuriphilusMonti Cimini (Italy)[7]
E. pelaKunming (China)[34]
C. macrocarpumH. subtenuisAlberta (Canada)[41]
C. oxysporumAnoplolepis custodiens
Aonidiella aurantii		Eastern Transvaal (South Africa)[74]
A. gossypiiMataffin (South Africa)[75]
A. melliferaAssiut Governorate (Egypt)[15]
C. lanigeraAnakapalle (Andhra Pradesh, India)[5]
C. aonidumEastern Transvaal (South Africa)[74]
Hypothenemus hampeiEl Tizal (Mexico)[76]
unidentified MuscidaeEastern Transvaal (South Africa)[74]
Planococcus citriMataffin (South Africa)[75]
P. oleaeMirandela–Bragança region (Portugal)[53]
Pseudococcus longispinusEastern Transvaal (South Africa)[74]
P. melanariusAssiut Governorate (Egypt)[15]
Pulvinaria aethiopica
Toxoptera citricidus
Trioza erytreae		Eastern Transvaal (South Africa)[74]
C. perangustumX. compactusGrottammare (Italy)[77]
C. pini-ponderosaeunidentified DipteraCoimbra (Portugal)[31]
C. pseudocladosporioidesEulophid parasitoid Averno; Napoli (Italy)[16]
D. suzukiiMaryland (USA)[9]
C. ramotenellumA. nepetaeNapoli (Italy)[16]
C. sphaerospermumBradysia impatiensSouth Korea[78]
B. brassicaeAssiut (Egypt)[10]
D. kuriphilusMonti Cimini (Italy)[7]
E. pelaKunming (China)[34]
unidentified grasshoppersUlu-Endau (Malaysia)[21]
H. vitripennisSouthern California (USA)[42]
H. subtenuisAlberta (Canada)[41]
Imbrasia belinaFrancistown (Botswana)[79]
M. maculipesDumoga-Bone (Sulawesi, Indonesia)[21]
Macrotermes barneyiGuangdong (China)[80]
Onychiurus pseudofimetariusAuckland (New Zealand)[1]
P. melanarius
R. ferrugineus		Assiut Governorate (Egypt)[15]
S. birosimensisCentral and Western Japan[55]
T. piniperdaMościska Forest (Poland)[57]
T. infestansMendoza; Santa Fe (Argentina)[59]
T. castaneumGoseong; Sangju (South Korea)[61]
Cladosporium sp.Acmaedora flavolineataTurkey[81]
Acronicta majorHangzhou (China)[82]
Adelges piceaeGaspé Peninsula (Canada)[83]
Adelges tsugaeNortheastern USA[84]
Aedes albopictusNice; Portes-lès-Valence; Saint Priest (France)
Mananjary; Toamasina; Tsimbazaza (Madagascar)
Bình Dương; Hồ Chí Minh; Vũng Tàu (Vietnam)
Villeurbanne (France)
Lubbock (Texas, USA)
[85]

[86]
[87]
Aedes aegyptiLubbock (Texas, USA)[87]
Aedes japonicusUrbana (Illinois, USA)[88]
Aedes triseriatus
Agathomyia sp.Yaoshan Mountain (China)[89]
Agrilus maliYining (China)[90]
Aleocharinae spp.Denmark[91]
Aleurocanthus spiniferusSouthern Anhui (China)[92]
A. aepimBahia (Brazil)[93]
Alniphagus aspericollisGreater Vancouver region (Canada)[94]
Anastrepha fraterculusCaxias do Sul (Rio Grande do Sul, Brazil)[95]
Anopheles coluzziiKoulikoro (Mali)[96]
Apertochrysa formosanusSugadaira-Kogen (Japan)[97]
A. craccivora
Aphis durantae
A. gossypii		Dakahlia Governorate (Egypt)[98]
Apis ceranaChiang Mai (Thailand) [99]
A. melliferaTucson (Arizona, USA)
Assiut Governorate (Egypt)
Grugliasco (Italy)		[100]
[15]
[101]
Halle (Germany)
Athens (Greece)
Lublin (Poland)
Chiang Mai (Thailand)
London (United Kingdom)		[102]

[99]

Asphondylia glabrigerminisMittagong; Melbourne area (Australia)[103]
A. micromeriaeAstroni Nature Reserve (Italy)[104]
Asphondylia serpylliLublin area (Poland)[105]
A. capiguara
A. laevigata		Botucatu (São Paulo, Brazil)[17,106]
Atomaria spp.Denmark[91]
Bactrocera oleaeCalabria (Italy)[107]
Bactrocera tryoniNew South Wales; Victoria (Australia)[108]
Bemisia tabaciDakahlia Governorate (Egypt)[98]
Bombus terrestrisBeijing (China)[109]
Bombyx moriHangzhou (China)[82]
B. impatiensSouth Korea[110]
Calliopum aeneumDenmark[91]
Callosobruchus maculatusColumbus; Miami County (Ohio); Erie County (Pennsylvania, USA)[111]
Carabidae spp.Uckermark (Germany)[112]
Carpophilus sp.Gualmatán (Colombia)[113]
Ceroplastes floridensisMansoura (Egypt)
Kiryat Tivon (Israel)		[22]
[114]
Ceroplastes rusciMansoura (Egypt)
Larnaca (Cyprus)		[115]
[114]
Ceroplastes sp.Malaga (Spain)[114]
Chalcophora detritaTurkey[116]
Coccus hesperidumRamat HaShofet (Israel)[114]
unspecified ColeopteraÅs (Norway)[25]
unidentified CollembolaWarren Woods (Michigan, USA)[29]
Colletes cuniculariusAve-et-Auffe (Belgium)[117]
Cortinicara gibbosaDenmark[91]
Crocothemis serviliaHefei (China)[118]
Ctenolepisma longicaudatumWien (Austria)[119]
C. pipiensSan Francisco; San Rafael (California, USA)[120]
C. quinquefasciatusNakhon Nayok (Thailand)
Mar del Plata (Argentina)		[121]
[122]
Cydia pomonellaAustria[123]
Cytilus sericeusOstrava (Czechia)[124]
Diabrotica sp.Gualmatán (Colombia)[113]
Diabrotica virgiferaDeutsch-Jahrndorf (Germany)[125]
Diaphania pyloalisHangzhou (China)[82]
unidentified DipteraWarren Woods (Michigan, USA)[29]
Dolichovespula sylvestrisHavreballe Forest (Denmark)[126]
D. suzukiiMaryland (USA)[127]
Drosophilidae spp.
Enicmus transversus		Denmark[91]
E. pelaKunming (China)[3]
Ferrisia virgataMadurai (India)[128]
H. vitripennisSouthern California (USA)[42]
Harmonia axyridisHubei (China)[129]
Helicoverpa armigeraNarrabri; New South Wales (Australia)[130]
H. hampeiChiapas (Mexico)
El Tizal (Mexico)		[131]
[76]
Illiciomyia yukawaiMie Prefecture (Japan)[132]
Ips acuminatusLibavá (Czechia)[133]
Ips cembrae
Ips duplicatus		Rouchovany (Czechia)
I. sexdentatusRouchovany (Czechia)
Northeastern Ukraine		[133]
[44]
Ips typographusRouchovany (Czechia)[133]
unidentified LepidopteraCoimbra (Portugal)[31]
Liparthrum colchicum Migliarino Natural Park (Italy)[134]
Loberus impressusIberia Parish (Louisiana, USA)[135]
Lonchodes brevipesSingapore[136]
Lonchoptera spp.Denmark[91]
Lutzomyia sp.Antioquia Department (Colombia)[137]
Lymantria dispar asiaticaHarbin (China)[138]
Macrosaccus robiniellaTransylvania (Romania)[139]
Marchalina hellenicaIschia (Italy)†
Matsumurasca onukiivarious regions in China[140]
Meiltaea cinxiaEckerö; Sund (Finland)[141]
Milviscutulus mangiferaeUpper Galilee (Israel)[114]
Minettia fasciata
Minettia longipennis
Minettia plumicornis		Denmark[91]
M. domesticaAhwaz (Iran)[142]
Gauteng Province (South Africa)[70]
Bruxelles (Belgium)
Butare (Rwanda)		[143]
Neobathyscia mancinii
Neobathyscia pasai		Damati Cave (Italy)
Tana delle Sponde Cave (Italy)		[144]
N. lugensHangzhou (China)[145]
Orchelimum vulgareHouston (Texas, USA)[146]
Orthoperus brunnipesDenmark[91]
Orthotomicus erosusItalian harbors, on imported wood[147]
Ostrinia nubilalisAndau (Germany)[125]
Paracoccus marginatusMadurai (India)[128]
Parasaissetia nigraKiryat Tivon (Israel)[114]
Paravespula vulgarisHavreballe Forest (Denmark)[126]
Parectopa robiniellaTransylvania (Romania)[139]
Phlebotomus papatasiTehran (Iran)[148]
Phlebotomus spp.Tunisia[149]
Phylloicus sp.Adolpho Ducke Reserve (Amazonas, Brazil)[150]
Pieris brassicaeBornem (Belgium)
Randwijk; Wageningen (Netherlands)		[151]
Planococcus ficusGholan Heights (Israel)[152]
Poecilocerus pictusChennai (India)[153]
Polygraphus polygraphusRouchovany (Czechia)[133]
Probergrothius angolensisNamibia[154]
Pseudatomoscelis seriatusCollege Station (Texas, USA)[155]
unidentified PsocopteraWarren Woods (Michigan, USA)[29]
Psylliodes attenuataDaqing; Harbin; Changchun; Qujing (China)[156]
P. oleaeMirandela–Bragança region (Portugal)[53]
P. melanariusAssiut Governorate (Egypt)[15]
Pulvinaria aurantiiNorthern Iran[157]
Pulvinaria psidiiMansoura (Egypt)[22]
Pulvinaria tenuivalvataDakahlia Governorate (Egypt)[158]
R. ferrugineusAssiut Governorate (Egypt)[15]
Saissetia sp.Larnaca (Cyprus)[114]
Sapromyza quadripunctataDenmark[91]
Scolypopa australisNelson (New Zealand)[159]
S. birosimensisCentral and Western Japan[55]
S. frugiperdaGuangzhou (China)
Kansas (USA)		[160]
[12]
Stenochironomus sp.Adolpho Ducke Reserve (Amazonas); Lajeado State Park (Tocantins, Brazil)[63,150]
Stephostethus lardarius
Stilbus testaceus		Denmark[91]
Taeniothrips inconsequensNortheastern USA[84]
T. molitorAarhus (Denmark)
Shenyang (China)		[72]
[161]
Thaumastocoris peregrinusSoutheast Uruguay[162]
unidentified ThysanuraCoimbra (Portugal)[31]
Toumeyella parvicornisCampania (Italy)[163]
Trialeurodes riciniDakahlia Governorate (Egypt)[98]
T. brasiliensis
T. pseudomaculata		Rio de Janeiro (Brazil)[58]
T. castaneumSouth Korea[61]
Triplectides sp.Lajeado State Park (Tocantins, Brazil)[73]
Xyleborinus saxeseniiItalian harbors, on imported wood
Northeastern Italy
South Florida (USA)		[147]
[164]
[165]
Xyleborus bispinatus
Xyleborus volvulus		South Florida (USA)[165]
X. compactusCentral Florida (USA)
Migliarino Natural Park (Italy)		[166]
[134]
Xylosandrus crassiusculusSouth Florida (USA)[165]
Xylosandrus germanusNortheastern Italy[164]
C. subtilissimumA. capiguaraBotucatu (São Paulo, Brazil)[17]
C. subuliformeD. citriGuilin (China)[167]
unidentified StreblidaeFurna do Morcego (Pernambuco, Brazil)[168]
Triplectides sp.Lajeado State Park (Tocantins, Brazil)[73]
C. tenuissimumM. mutatusBragado (Argentina)[47]
M. persicaeSalah El-Din Governorate (Iraq)[48]
unidentified ThysanuraCoimbra (Portugal)[31]
Trachymela sloaneiGuangdong (China)[169]
C. uredinicolaA. gossypiiDakahlia Governorate (Egypt)[170]
Bemisia spp.Dakahlia Governorate (Egypt)[18]
B. tabaciDakahlia Governorate (Egypt)[170]
C. veloxB. oleaeCalabria (Italy)[107]
T. castaneumMungyeong; Sangju (South Korea)[61]
C. verrucocladosporioidesTriplectides sp.Lajeado State Park (Tocantins, Brazil)[73]
C. xanthocromaticumM. persicaeSalah El-Din Governorate (Iraq)[48]
* Colors are indicative of the order to which the species belong, as follows: Blattodea, Coleoptera, Collembola (Entomobryomorpha, Poduromorpha), Diptera, Hemiptera, Hymenoptera, Lepidoptera, Neuroptera, Odonata, Orthoptera, Phasmatodea, Psocoptera, Thysanoptera, Thysanura, Trichoptera. Identification of the strains isolated from the underlined sources is based on complete set of DNA markers. † Original isolation recently obtained by E.R.
Table 2. Reported effectiveness of conidial suspensions of Cladosporium in inducing mortality on insect pests.
Table 2. Reported effectiveness of conidial suspensions of Cladosporium in inducing mortality on insect pests.
Cladosporium SpeciesSourceInsect TargetsCountryReference
C. cladosporioidesBemisia sp.Bemisia sp.Egypt[18]
Brevicoryne brassicaeB. brassicaeEgypt[20]
Culex quinquefasciatusC. quinquefasciatusIraq[28]
endophyticDuponchelia fovealisBrazil[188]
Kermes sp.Hemiberlesia pitysophilaChina[45]
Lycorma delicatulaTenebrio molitorUSA[46]
Myzus persicaeM. persicaeIraq[48]
Nilaparvata lugensBemisia tabaciBangladesh[49]
Pulvinaria aurantiiAphis fabaeIran[157]
Sitophilus oryzaeRhyzopertha dominica
Sitophilus zeamais
Trogoderma granarium		Pakistan[56]
soilMetopolophium dirhodumEgypt[189]
C. oxysporumendophyticA. fabaeAlgeria[190]
endophyticChilo partellusIndia[191]
Planococcus citriPseudococcus longispinus
Pulvinaria aethiopica
Toxoptera citricida
Trioza erytreae		South Africa[75]
unknownAphis craccivoraIndia[192]
C. sphaerospermumendophyticD. fovealisBrazil[188]
Cladosporium sp.Helicoverpa armigeraAphis gossypii
B. tabaci
H. armigera		Australia[130]
Spodoptera frugiperdaS. frugiperdaChina[169]
Cladosporium spp.several species of sap-sucking HemipteraA. craccivora
A. gossypii
B. tabaci		Egypt[98,193]
C. subuliformeDiaphorina citriD. citriChina[167]
C. tenuissimumM. persicaeM. persicaeIraq[48]
Trachymela sloaneiS. frugiperdaChina[194]
C. uredinicolaA. gossypii
B. tabaci		A. gossypii
B. tabaci		Egypt[170]
Bemisia sp.Bemisia sp.Egypt[18]
C. xanthocromaticumM. persicaeM. persicaeIraq[48]
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Nicoletti, R.; Russo, E.; Becchimanzi, A. Cladosporium—Insect Relationships. J. Fungi 2024, 10, 78. https://doi.org/10.3390/jof10010078

AMA Style

Nicoletti R, Russo E, Becchimanzi A. Cladosporium—Insect Relationships. Journal of Fungi. 2024; 10(1):78. https://doi.org/10.3390/jof10010078

Chicago/Turabian Style

Nicoletti, Rosario, Elia Russo, and Andrea Becchimanzi. 2024. "Cladosporium—Insect Relationships" Journal of Fungi 10, no. 1: 78. https://doi.org/10.3390/jof10010078

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