Will moth larvae pupate in response to food deprivation

Will moth larvae pupate in response to food deprivation

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Context: Trying to exterminate a clothes moth infestation.

Supposing you have a room with clothes moths, or rather, with clothes moth eggs and larvae.

Suppose then, that you remove all of the soft furnishings, all of the clothes, all of the carpet, and you thoroughly clean and vaccuum to remove as much dust and hair as possible.

Would one predict that the remaining larvae would then immediately pupate into moths, in response to this food deprivation?

I know that lots of plants will suddenly flower or fruit if they perceive that they are at risk of dying, in order to reproduce before they die. I believe the moths are essentially just a reproductive phase of the moth, so I'd be unsurprised if they did pupate.

On the other hand, I imagine pupation takes a lot of energy and food reserves (not to mention needing material for the cocoon), so I can imagine that they might be unable to do this?

Which way do it go?

Will moth larvae pupate in response to food deprivation

Not officially an expert, but I have been watching clothes moths for years and haven't noticed any small ones, so dont expect early pupation. I would expect small pupae to give small moths (like with houseflies). I do see the hormone traps reduce population, but only with lots of them (2 per small room).


Natural Enemies

Moths are fecund invertebrates that produce 100–10,000 times more ova than will ultimately survive. A panoply of biotic and abiotic mortality factors comes into play during every life stage (and larval instar) to limit population growth. Biotic factors include an enormous range of pathogens, parasites, and predators. Population numbers of many pest species, especially in forests and other stable communities, often are controlled by viruses, bacteria, and fungi. Viruses are often highly specific, and one wonders if there might be nearly as many lepidopteran viruses as there are moths and butterflies. There are comparatively few recorded species of bacteria and fungi that attack insects, but the taxonomy of insect pathogens, especially those that cannot be easily cultured, is understudied although some are highly specific, others have broad host ranges. Bacillus thuringiensis, the most widely used pathogen for the control of Lepidoptera in gardens, agriculture, and forests, has a host range that includes hundreds (thousands?) of species across many superfamilies. Fungal pathogens, mostly in the Entomophthorales and Fungi Imperfecti, often require special conditions such as a period of high humidity before appreciable infection will occur. A species of special note is Entomophaga maimaiga. This fungal pathogen was introduced into North America from Japan in 1910 and 1911 as a biological control agent to stem the ever-increasing populations of the European gypsy moth (Lymantria dispar). The introduction was considered unsuccessful until 1989 when the fungus began showing up in caterpillar cadavers throughout the Northeast. It has since spread throughout much of the North American range of the gypsy moth and continues to be a principal mortality agent for the gypsy moth caterpillar. A diverse assemblage of nematodes and nematomorphans also attacks Lepidoptera, especially the larvae, but their importance is generally regarded as modest among nonsoil dwellers. Microsporidians affect both fertility and longevity in many moths.

Wolbachia are known from across the Obtectomera and Heterocera, but infections likely occur across the order, given that some 25–75% of all insect species are known to be hosts to these endosymbiotic rickettsia-like bacteria ( Kozek and Rao, 2007) . In many species the cytoplasmic infections are passed vertically from mother to offspring (in the egg), whereas in others transmission occurs through sexual contact. The consequences of infection are both varied and complex, and adequate description is beyond the scope of this treatment. Known effects within the order include male-killing (through selective killing of male embryos) feminization, whereby genetic males are rendered into functional females and egg–sperm cytoplasmic incompatibility (between sexual partners/mating strains). The nature of the infections is taxon-dependent, and ranges from entirely pathogenic and lethal (e.g., in those where males are killed) to symbiotic (e.g., some Wolbachia-infected insect lineages are reported to confer resistance to an array of RNA viruses and some pesticides). In Phyllonorycter blandcardella (Gracillariidae) ( Figure 4 ) – a leaf miner of apple – Wolbachia infections maintain green tissues about the larval mine, even while the remainder of the leaf is yellowing, allowing larvae to complete their development before the host leaf senesces ( Kaiser et al., 2010 ).

Lepidoptera are primary hosts for a multitude of parasitoids. Three of the largest insect families on this planet owe much of their evolutionary success to the Lepidoptera: the Braconidae (>10,000 species) (Hymenoptera), Ichneumonidae (>10,000 species) (Hymenoptera), and Tachinidae (>20,000 species) (Diptera). Another 20 or so families of insect parasitoids attack Lepidoptera, all but two of which (Diptera: Bombyliidae and Sarcophagidae) are parasitic hymenopteran wasps, mostly in Chalcidoidea and Proctotrupoidea. All four life stages are exploited by parasitoids, although most attack larvae. Typically only one or a few instars are attacked by a given wasp species. Most Hymenoptera are endoparasitoids that grow slowly initially, consuming only blood (hemolymph), their own nutritive cells released into the host, or nonvital host cells and tissues, then finish off the host in a burst of feeding and growth, often triggered by the host larva's size or hormonal state. In some cases this may be months after the wasp eggs were laid within the body of the caterpillar (see Wagner et al., 2011 for a brief synopsis). Some of the smallest insects, trichogrammatid and mymarid hymenopterans, with body lengths of only 0.2 mm, are egg parasites. A great majority of insect parasitoids are specialists that utilize but one or a few related species. Large, long-lived caterpillars support rich guilds of parasitoids: across eastern North America, 29 fly and wasp parasitoids are recorded from Hyalophora cecropia, at least three of which are believed to be specialists on this saturniid and its congeners. Large caterpillars may form the base of an entire food web, as evidently there are six hymenopteran hyperparasitoids that parasitize the parasitoids of Hyalophora, and one wasp hyperparasitoid that principally attacks the hyperparasitoids. Askew and others have noted that leaf miners and gall-forming caterpillars have an exceptionally rich parasitoid fauna. Phyllonorycter apparella (Gracillariidae) has a wingspan of only 7 mm, yet larvae and pupae of this leaf miner are known to host 20 species of hymenopteran parasitoids and hyperparasitoids.

Invertebrate predators of major importance include mites, spiders, predaceous stink bugs, beetles, robber flies, ants, and wasps. Many of the physical, chemical, and behavioral defenses of caterpillars are known to be effective in thwarting the pillages of ants ( Figure 5 ). No doubt, these defenses are of special importance in tropical ecosystems where ants make up much of the insect biomass. Mammals, birds, reptiles, and amphibians are the principal vertebrate predators. Birds are such an important selection pressure for externally feeding caterpillars that there is a growing consensus that they have shaped not only what many caterpillars look like, but also what, when, and how they feed ( Heinrich, 1993 ). Their influence on adult color patterns and behavior also seems to be pre-eminent. Bats, too, have been an exceptionally important evolutionary force in the diversification of the order. Hearing organs, sensitive to the frequencies emitted by bats, have evolved multiple times within one of these groups, the Arctiinae (tiger moths), some chemically protected taxa have evolved the ability to signal back to bats. On detecting ultrasound frequencies, these moths begin clicking, presumably advertising their whereabouts (and toxicity). In addition, there is growing evidence that some can even thwart bats by jamming their echolocation sensory systems. Corcoran et al. (2009) have shown that Bertholida tiger moths employ a sound-producing tymbal located below the hindwing to generate ultrasonic frequencies that are sufficiently disruptive to cause bats to veer off course and abort their normal attack repertoire.


Metamorphosis is a type of molting through which an immature insect transforms into its adult form [1, 2]. Under optimal conditions for larval growth, holometabolous insects generally undergo a species-specific number of larval molts and eventually metamorphose into adults via the pupal stage. Last instar larvae must decide when to initiate the processes leading to metamorphosis however, our knowledge on how they determine the timing of metamorphosis is limited to a few species of insects [3].

The timing of and cue for larval decision toward metamorphic molting have been intensively studied in the tobacco hornworm Manduca sexta (Lepidoptera: Sphingidae) [3, 4]. The key signal for initiating metamorphosis is secretion of prothoracicotropic hormone (PTTH) from the brain, which stimulates secretion of ecdysone from the prothoracic glands. Last instar larvae of M. sexta cease feeding in response to a small surge of ecdysteroid, and subsequently pupate via wandering and prepupal stages. In the last instar, however, removal of hemolymph juvenile hormone (JH) is a prerequisite for the onset of metamorphosis in M. sexta because JH inhibits the secretion of PTTH from the brain [5, 6]. Processes to remove JH, i.e., cessation of JH synthesis and clearing of hemolymph JH by JH esterase, are initiated at the time when the last instar larvae of M. sexta attain a critical weight (CW) [5, 7]. JH is cleared from the hemolymph within an approximately constant time after the attainment of CW, which drives larvae to pupate. Since CW is attained in the middle of the feeding phase, food deprivation after a certain period of feeding does not affect the timing of pupation: the starved larvae become small pupae at the predetermined time point (Fig 1A). This mechanism to determine the onset of metamorphosis is referred to as critical weight-mediated pupation. Critical weight-mediated pupation has also been found in another moth, Trichoplusia ni (Lepidoptera: Noctuidae) [8], and the fruit fly, Drosophila melanogaster (Diptera: Drosophilidae) [9, 10], although critical weight-mediated pupation in D. melanogaster may not be identical to that in M. sexta [11].

A) Critical weight-mediated pupation (CWmP). B) Typical starvation-induced pupation (SiP). Modified from Nijhout (2008) [15].

Another mechanism to determine the onset of metamorphosis, starvation-induced pupation (SiP), has been found in Coleoptera (Fig 1B). Shafiei et al. [12] found that starvation of a larva of the dung beetle Onthophagus taurus (Coleoptera: Scarabaeidae) during the feeding phase in the last instar induces precocious pupation, provided that the larva has attained a minimum viable weight. The time between the start of starvation and the occurrence of pupation was constant regardless of the larval body weight upon starvation or the length of prior feeding, indicating that in O. taurus larvae, the decision to pupate is made a constant time after the start of starvation regardless of its timing in the feeding phase (Fig 1B). SiP is considered as a bailout mechanism that operates when an animal’s dietary environment deteriorates [3]. SiP has also been reported for two other beetles: fungus beetle Dacne picta (Erotylidae) [13] and blister beetle Epicauta gorhami (Meloidae) [14].

As described above, the mechanism to determine the timing of pupation appears to differ with the lineage of insects however, the insect species studied to date are very limited in number and taxonomy [15]. The yellow-spotted longicorn beetle Psacothea hilaris (Coleoptera: Cerambycidae) is a pest of mulberry and fig trees [16]. The native range of P. hilaris is eastern Asia, but they have recently invaded Europe [16–18]. Their larvae infest branches of live host trees, and the beetles damage leaves. Adults of P. hilaris in the field show a large variation in body size, which affects mating behavior [19]. Two types of P. hilaris that differ in morphology, ecology, and physiology, i.e., west-Japan type and east-Japan type, are known to inhabit western and eastern parts of the main islands of Japan, respectively [20]. The control of larval development in P. hilaris has been intensively studied using the west-Japan type [21–23]. Under long-day conditions in the laboratory, west-Japan type P. hilaris larvae feeding on an artificial diet ad libitum pupate from the 4th or 5th instar [21, 23]. When the larvae are starved upon ecdysis into 5th instar, they precociously develop into pupae that are smaller than the counterpart raised under continuously-fed conditions [22]. A similar starvation-induced earlier pupation also occurs in 4th instar larvae [23, 24]. These findings suggest the presence of a SiP mechanism in P. hilaris.

The primary aim of the present study was to investigate whether SiP occurs in P. hilaris. We observed the occurrence of pupation in response to food deprivation at various timings in the 5th-instar larvae of P. hilaris. Unexpectedly, P. hilaris was found to exhibit an atypical SiP system: SiP occurred only in larvae starved in the late feeding phase of the instar. Based on our findings, we discuss conservative and nonconservative aspects of the SiP systems in Coleoptera, and also discuss the mechanism to determine the timing of pupation under continuously-fed conditions.

Figure 1.

Battus philenor with ventral surface visible. High quality figures are available online.

In Arizona, the iridescent warning coloration of B. philenor on the ventral hind wings varies in ways that may be attributed to differences in larval diet. Rutowski et al. (2010) reported significant differences in the iridescent coloration between lab-reared individuals fed ad libitum and field-caught individuals. Therefore, variation in larval food availability could be a source of adult color variation, including warning color variation. As in the B. philenor populations previously studied, populations in Arizona are likely to experience food deprivation as larvae because their hostplant, Aristolochia watsonii Wooton and Sandley (Aristolochiales: Aristolochiaceae), is a small plant that larvae often completely denude of leaves before completing development (personal observation). Therefore, to evaluate the role of food restriction on warning coloration and to determine if observed natural variation in iridescent signals is due to food deprivation, the amount of food to which B. philenor larvae had access was varied among three different treatments. The adult coloration was compared among treatment groups. The effects of food deprivation were evaluated for three different color patches: the iridescent blue field and the orange spots of the ventral hind wing surface, which contribute to the warning signal, and the iridescent blue on the male dorsal hind wing. The male dorsal hind wing is a signal used by females, likely to assess either male quality or species identity (Rutowski and Rajyaguru 2013).

If food deprivation is a source of variation in the iridescent signals of B. philenor, it is predicted that food deprivation will cause increased brightness, shorter wavelength hues, and higher chroma for the ventral surface iridescence and higher intensities in the dorsal iridescence. These expectations are based on the difference between animals reared in the lab and those from the field reported in Rutowski et al. (2010). Additionally, if there are differences in the ventral surface iridescence and the orange spots between treatments, even if this variation does not match that found in Rutowski et al. (2010), it will indicate that food restriction can be a significant source of intraspecific variation in warning signals.


Measures of body condition

Compared to those with unrestricted access to food during development, food-restricted larvae developed into significantly smaller adults with smaller fat reserves. This result was expected based on previous studies of larval food limitation in butterflies (e.g., Bauerfeind and Fischer 2005 Boggs and Freeman 2005).The size of the adults produced from Day 3 treatment larvae was within the range of adult sizes observed in the field ( Rutowski et al. 2010), indicating that the level of food deprivation induced in the Day 3 treatment was likely within the range of food limitation that this species experiences in nature. The effect of larval food deprivation on the level of sequestered aristolochic acids is currently under investigation.

Warning coloration

Food restriction produced significant variation in both the orange and blue components of the ventral warning coloration of B. philenor. The hue of the ventral blue iridescence shifted to shorter wavelengths with increased food deprivation. The chroma of the orange spots decreased with increased food deprivation.

These results suggest proximate links between coloration, the structures and chemicals that produce color, and diet quantity, but these linkages are not clear at the moment (see Kemp et al. 2006). For B. philenor, this is true for both the orange spots and the blue patches, but there are some possible connections that could be tested. The diffuse reflection of the orange spots indicates that the pigments played a major role in shaping the reflectance spectrum by absorbing short wavelengths, which allows longer wavelengths to be reflected from the wing surface ( Rutowski et al. 2005). The specific pigments involved are not known but are likely to be ommachromes or papilochromes synthesized de novo by the butterflies from the amino acid tryptophan ( Nijhout 1991). The chroma of the orange spots should be positively related to the quantity of pigment in the scale, as more pigment means greater absorption of short wavelength light. During development, diet-restricted individuals whose orange is less chromatic may deposit less pigment in their scales due to a lower availability of tryptophan. On the other hand, the iridescent blue is likely a product of thin film interference, and the higher hue of diet-restricted individuals suggests a thicker film ( Land 1972). If true, it is not clear how the film would be thicker in diet-restricted individuals who presumably experience restrictions in the materials needed to build these cuticular films. Again, questions about the potential proximate connections between diet and color phenotype remain untested but warrant investigation.

An experiment with captive Curve-billed Thrashers showed that the blue iridescence and the orange spots of the ventral hind wing were used by avian predators to recognize B. philenor as distasteful, and each component alone elicited a rejection response ( Pegram et al. 2013). It is unknown whether the variation in the hue of the iridescent patches induced by food restriction would alter the effectiveness of the aposematic coloration of B. philenor. Although both reptiles and invertebrates (e.g., spiders and dragonflies) have been observed preying on B. philenor ( Rausher 1979b, 1980), insectivorous birds are likely to be their most common predators in Arizona (Pe-gram, Han, and Rutowski, unpublished data). Visual models indicated that birds should be able to distinguish the spectral differences observed in adult coloration due to treatment. However, even though avian predators may be able to discriminate these colors, predators may generalize a learned warning signal to similar colors ( Ham et al. 2006 Ruxton et al. 2008 Svádová et al. 2009) or the differences may not be detectable in complex and changing conditions of lighting and background ( Lindstedt et al. 2011). Either way, the color shifts caused by food deprivation may not decrease signal effectiveness. Signal effectiveness could also be influenced if the observed responses altered conspicuousness ( Gittleman and Harvey 1980 Gamberale-Stille and Tullberg 1999 Lindström et al. 1999 Riipi et al. 2001 Lindstedt et al. 2008). From these results, it is concluded that food deprivation did contribute to intraspecific variation in warning coloration, but determining if this variation correlates with signal effectiveness will require further study.

Response of iridescent coloration and comparison to natural coloration

The hue of the ventral and dorsal iridescent patches shifted to shorter (bluer) wavelengths with increased food deprivation. This was the opposite direction of what was predicted based on the results of Rutowski et al. (2010). Rutowski et al. (2010) also found that lab-reared individuals had more intense ventral iridescence than did field-caught individuals, where no effect of rearing conditions on ventral iridescence brightness was found in our study. Therefore, differences in the lab and field individuals previously observed were not likely due to increased food deprivation in the field-caught B. philenor. The difference between the dorsal and ventral surfaces in male chroma observed in our study was expected based on Rutowski et al. (2010), but the lack of treatment effects and differences between the sexes was inconsistent with their results. Differences between male and female ventral hue were observed in the previous study, but not in our study. From these differences, it can be concluded that the differences observed between lab and field individuals in Rutowski et al. (2010) were not likely caused by food deprivation in field individuals.

However, differences in the results of these studies could be caused by at least two other factors. First, there were differences between the studies in seasons in which observations occurred (spring ( Rutowski et al. 2010) vs. autumn (our study)). Second, the larvae of the field-caught individuals could have undergone food deprivation throughout the larval stage, while the larvae in our experiment only underwent food restriction in the last larval instar.

Also, the hue and chroma of an individual’s dorsal iridescence were correlated with the hue and chroma of its ventral wing surface, which suggests a coupling of the iridescent surfaces. This is consistent with the findings of Rutowski et al. (2010), but the causes of this coupling are not understood at this time.

Because the dorsal coloration may serve as a signal of male quality ( Rutowski and Rajya guru 2013), we expected to see heightened condition dependence over a naturally selected signal (e.g., Andersson 1986 Cotton et al. 2004a, b Kemp 2008). However, there were no significant surface by treatment interactions to suggest there is heightened condition dependence of the iridescence on the dorsal surface.

Spores, please! Gypsy moth larvae love poplar leaves infected by fungi

A gypsy moth caterpillar (Lymantria dispar) relishing the spores of Melampsora larici-populina a rust fungus that has spread on a poplar leaf. The new study shows that the insect is not only herbivorous, but also fungivorous, that is, likes to feed on nutrient-rich fungi. Credit: Franziska Eberl, Max Planck Institute for Chemical Ecology

Black poplar leaves infected by fungi are especially susceptible to attack by gypsy moth caterpillars. A research team at the Max Planck Institute for Chemical Ecology in Jena, Germany, has now further investigated this observation. The scientists found that the young larvae of this herbivore upgrade their diet with fungal food: Caterpillars that fed on leaves covered with fungal spores grew faster and pupated a few days earlier than those feeding only on leaf tissue. The higher concentrations of important nutrients in fungi, such as amino acids, nitrogen and vitamins, are probably the reason for their better performance. The results shed new light on the co-evolution of plants and insects, in which fungi and other microorganisms play a much greater role than previously assumed.

Gypsy moth caterpillars are known as feeding generalists this means they accept a large variety of deciduous trees species and shrubs as their food plants. Outbreaks of this species have been documented every now and then also in German forest ecosystems.

Sybille Unsicker and her research team are investigating how poplars defend themselves against herbivores, including the gypsy moth. The scientists had observed that these trees downregulate their defense against the voracious insect when they are simultaneously being attacked by fungi. "We noticed that caterpillars are attracted by the odor of fungus-infested poplars, so we wondered why this is so: Would the caterpillars prefer to feed on infested leaves as well? Would this provide an advantage? And if so, what kind of chemicals are responsible for this?" first author Franziska Eberl asks, describing the basic questions of the study.

Feeding experiments in which the gypsy moth larvae were offered a choice of leaves with or without fungal infection revealed the clear preference of the caterpillars for leaves infected with fungi. In the early larval stage, they even consumed the fungal spores on the leaf surface before feeding on leaf tissue. "Whether rust fungi or mildew, young caterpillars selectively fed on the spores and preferred to feed on infected leaves," explains Franziska Eberl.

Chemical analyses showed that mannitol, a substance that is also used as an artificial sweetener in human food, is primarily responsible for this preference. Eberl also monitored larval fitness, which is shown by how well larvae develop—a measurement that depends largely on their diet. "Larvae that consume fungus-infected leaves develop faster and also pupate earlier. This gives them an advantage over their siblings who feed on healthy leaves. Important nutrients, such as amino acids, nitrogen and B vitamins, are likely responsible for increased growth, because their concentration is higher infected leaves," said the researcher.

The role of microorganisms puts the co-evolution of plants and insects in a new light

The observation that an insect classified as an herbivore is actually a fungivore—at least in its early larval stage—was a real surprise for the research team. "Our results suggest that microorganisms living on plants might have a more important role in the co-evolution of plants and insects than previously thought," says Sybille Unsicker, head of the study. "In the black poplar trees from our study, fungal infestation occurs every year. It is therefore indeed imaginable that herbivorous insects have been able to adapt to the additional fungal resource. Especially with regard to the longevity of trees, the evolutionary adaptation to a diet consisting of leaves and fungi seems plausible for such insects."

Further investigations are needed to clarify how widespread fungivory is in other herbivorous insect species and what influence the combination of plant and fungal food has on the immune system of insects. It is possible that this food niche also has an effect on the insects' own defense against their enemies, such as parasitoid wasps. The role of microorganisms in the interactions between plants and insects has long been underestimated, even overlooked. This study is an important step to make up for that neglect.


Financial support to the first author through the University of Manitoba Graduate Fellowship (UMGF) and Manitoba Graduate Scholarship (MGS) is appreciated. We thank Judy Johnson (USDA ARS, Parlier, CA) and Gerhard Gries (Simon Fraser University) for their generosity in providing Indianmeal moth populations. We also thank Frank Arthur (USDA, ARS, Center for Grain and Animal Health Research, Manhattan, KS) Neil Holliday, Steve Whyard (University of Manitoba) and Desiree Vanderwel (University of Winnipeg) for their suggestions during this study.

Size compensation in moth larvae: attention to larval instars

Environmental perturbations such as starvation and poor diet often prevent animals from attaining their optimal sizes. When the perturbation has a transient character, compensatory responses are expected in terms of faster growth or a prolonged developmental period. In the case of insect larvae, details of such responses are insufficiently known at the proximate level. Attention to responses at the level of particular larval instars should promote an understanding of insect developmental plasticity also in a more general context. To provide an instar-specific analysis of compensatory growth, larvae of the moth Orgyia antiqua (L.) are reared on inferior diet during one larval instar. Responses in growth parameters are recorded in the course of the manipulated instars, as well as at the level of the entire larval period. The negative relationship between development time and size in response to the inferior food quality, typical of the entire larval periods, is also observed within the manipulated instars taken separately. The manipulated larvae remain smaller than the larvae of the control group (significant in males only), even by the end of the subsequent instar during which all individuals are provided with superior host. In males, close to full size compensation by the time of pupation is achieved only by means of adding an extra larval instar. The inability of larvae to fully compensate during one and even two instars is considered as an indication of the presence of constraints on the within-instar growth pattern. An alternative, adaptational explanation for the incomplete compensation could be based on the cost of prolonged development period. Given the ecological context of the species' life history, such an explanation appears less likely.


The greater wax moth was described for the first time in a colony of Apis cerana (eastern or Asiatic honeybee), that is, wild honeybees found in southern and eastern Asia. Its systematic position is presented in Fig. 1. Being a cosmopolitan species and pest of bee colonies, the greater wax moth has spread to almost all continents (except Antarctica), usually covering most or all of their areas (Kwadha et al. 2017). Its occurrence basically coincides with the beekeeping economy in individual countries, as this pest can be found in beehives or stored waxes causing a phenomenon called galleriosis (Fig. 2). According to the latest data, the greater wax moth has so far been confirmed in 27 countries in Africa, 9 in Asia, 5 in North America, 3 in Latin America, Australia and New Zealand and in 33 countries in Europe and almost all of the larger islands associated with them. It is expected that the species will continue to spread to unmanaged areas, which may be associated with changing climatic conditions (Kwadha et al. 2017 G. mellonella is a typical holometabolous insect, that is, it undergoes four developmental stages in its life cycle, namely, the egg, larva, pupa and adult (Smith 1965 Fasasi and Malaka 2006 Swamy 2008 Ellis, Graham and Mortensen 2013 Hosamani et al. 2017 Kwadha et al. 2017 Desai et al. 2019). Below, with the description of its developmental stages, we provide information about the general biology of each stage, including behaviour and characteristic morphological features.

Systematics of G. mellonella and imago (photograph: M. Kucharczyk).

Systematics of G. mellonella and imago (photograph: M. Kucharczyk).

Abandoned beehive inhabited by G. mellonella: pupal cocoons (p) found outside beehive (A) waxes affected by galleriosis (indicated by the arrow in (B)) and magnification thereof (C): eggs (e) and silk (s) on the wax (photograph: G. K. Wagner).

Abandoned beehive inhabited by G. mellonella: pupal cocoons (p) found outside beehive (A) waxes affected by galleriosis (indicated by the arrow in (B)) and magnification thereof (C): eggs (e) and silk (s) on the wax (photograph: G. K. Wagner).

Eggs, glued together, are laid in batches of 50 to 150 (Kwadha et al. 2017) or, as reported by Desai et al. ( 2019), even from 175 to 355. They are oval, white when laid and cream or pale pink when older. Reticulate and very rough, they are composed of interconnected polygons (squares, pentagons, hexagons and heptagons). The micropylar area is surrounded by concentrically arranged elements of the microstructure, reminiscent of rounded flower petals (Ellis, Graham and Mortensen 2013). The egg dimensions given by different authors are similar: length from 0.44 to 0.47 mm and width from 0.29 to 0.39 mm (Swamy 2008 Ellis, Graham and Mortensen 2013 Hosamani et al. 2017 Kwadha et al. 2017 Desai et al. 2019). About 4 days before eclosing, the larva is visible as a dark ring. Twelve hours before hatching, the fully formed larva is clearly visible through the thin chorion (Paddock 1918).


Larvae most often hatch in the morning, between 08.30 and 11.00 h (Hosamani et al. 2017 Desai et al. 2019). Depending on the research carried out, egg survival ranges from

84 to 100% (Pastagia and Patel 2007 Swamy 2008 Hosamani et al. 2017 Desai et al. 2019). Shortly after hatching, larvae move from the cracks and crevices to the honeycomb, where they begin to feed and build protective silken tubes, destroying the honeycomb structure in the process. The directional movement and feeding are probably stimulated chemically. This was confirmed by Paddock ( 1918) and Nielsen and Brister ( 1979), who observed that G. mellonella larvae isolated from honeycombs always went back towards their food source. Feeding larvae usually expand their ever-widening tubes towards the central part of the honeycomb, where they tend to accumulate. In the absence of food, cannibalism may occur (Nielsen and Brister 1979 Williams 1997).

In natural conditions, G. mellonella larvae feed on honeycombs, which contain a significant amount of beeswax, some honey, exuviae of bee larvae and pollen residues. From such food, they obtain a large amount of energy but relatively little protein (Kwadha et al. 2017). If the amount of dietary protein falls below a certain level, the larvae cease spinning silk (Jindra and Sehnal 1989), probably due to the lack of essential amino acids for silk protein synthesis (Shaik, Mishra and Sehnal 2017). The protein content also affects the rate of larval development. Their growth is fast on old honeycombs, which contain bee maggots and pollen, but very slow on white or new honeycombs. The positive dietary effect of bee pollen on the growth rate of G. mellonella larvae and the fertility of females developing from them was confirmed by Mohamed et al. ( 2014). The rapid growth of foraging larvae leads to complete destruction of the honeycombs within a week of colonisation (Hosamani et al. 2017). Larvae can also develop on an artificial diet consisting of cereal products, milk powder, yeast, honey and glycerol (Desai et al. 2019). A close relationship between larval diet quality and resistance against pathogens has been demonstrated: if there is a deficiency of nutrients, larvae become susceptible to Candida albicans Berhout infection (Banville, Browne and Kavanagh 2012).

Feeding greater wax moth larvae spin protective silken tubes, within which they are not detected by bees (Shaik, Mishra and Sehnal 2017). However, host workers have been repeatedly observed removing dead larvae (presumably killed) of this pest (G. K. Wagner, oral information). This fact undoubtedly undermines the 100% effectiveness of these silken structures to protect their owners. The composition of the silk from which the protective tubes are spun is similar to that in pupal cocoons. The core of the silk filament consists of heavy and light chain fibroins and the P25 chaperonin, whereas the filament coating is composed of sericins (Fedič, Žurovec and Sehnal 2002 Shaik, Mishra and Sehnal 2017). A feeding pause has been observed before each larval moult. Old cuticles are shed separately from the head capsule and the rest of the body. The average optimal larval development temperature for this moth is 29–33°C (Warren and Huddleston 1962 Nielsen and Brister 1979 Williams 1997). The average duration of each consecutive larval instar L1-L7 is 4.08, 5.72, 5.28, 6.96, 6.76, 7.64 and 8.40 days, respectively, giving a total duration of the larval stage of

45 days (Pastagia and Patel 2007 Swamy 2008 Hosamani et al. 2017 Rahman et al. 2017 Desai et al. 2019). The last two larval instars grow the most intensively (Ellis, Graham and Mortensen 2013).

Immediately after eclosing, the first larval instar (L1) is white, slim and very short (mean length 1.27 mm) (Hosamani et al. 2017). During further growth, it turns greyish white in colour, and from the third larval stage onwards its body begins to thicken conspicuously, becoming massive and stocky by the end of its development (Fasasi and Malaka 2006 Ellis, Graham and Mortensen 2013 Kwadha et al. 2017 Desai et al. 2019). Being very weakly sclerotised, most of the body surface of the first-instar larva is devoid of pigment, except for the head (the most strongly sclerotised part of the body). In later larval instars, the tergites of the pronotum and abdominal segment X as well as the protarsus and claws of the ventral prolegs, which gradually darken after each moult, taking different shades from light to dark brown, are also well sclerotised (Ellis, Graham and Mortensen 2013). In the fully coloured final stage larva, a bright ecdysial line is visible along the middle of the dorsal side (especially well marked on the prothorax) (Kwadha et al. 2017

The G. mellonella larva belongs to the polypod (eruciform or caterpillar-shaped) and peripneustic (nine pairs of spiracles) type. Its body consists of a head, a three-segmented thorax and an abdomen of 11 segments (Fig. 3). On the highly sclerotised head, there is a pair of short, two-segmented antennae, chewing mouthparts and four stemmata on each side – these are bright, oval and separated from each other (Ellis, Graham and Mortensen 2013). The presence of stemmata on the head of the G. mellonella larva is an important diagnostic character, which distinguishes the larva of this species from that of the lesser wax moth Achroia grisella, that is, another pyralid and apiary pest, which does not have this feature (Ellis, Graham and Mortensen 2013). The thorax bears three pairs of five-segmented thoracic legs (one pair per segment), each ending in a single hooked claw. There are prolegs on abdominal segments III-VI, which become visible 3 days after hatching (Desai et al. 2019). The terminal abdominal segment (XI) bears a pair of anal prolegs. There is one oval, brown and clearly visible spiracle on each side of the prothorax and on each side of abdominal segments I-VIII, a total of nine pairs (the peripneustic respiratory system) the last pair, on abdominal segment VIII, is the largest. The body bears rather thinly distributed, long, protruding, light brown, hair-like setae (Smith 1965, the present study).

Morphology of G. mellonella larvae. Dorsal (I), ventral (II) and lateral (III) view of a G. mellonella larva. A - sclerotised head with lateral stemmata, B - thorax, C - abdomen, D - antennae, E - chewing mouthparts, F - pair of thoracic legs, G - claw, H - pair of prolegs, I - anal prolegs, J - prothorax spiracle, K - abdominal spiracle, L - spiracle of abdominal segment VIII (the largest of all).

Morphology of G. mellonella larvae. Dorsal (I), ventral (II) and lateral (III) view of a G. mellonella larva. A - sclerotised head with lateral stemmata, B - thorax, C - abdomen, D - antennae, E - chewing mouthparts, F - pair of thoracic legs, G - claw, H - pair of prolegs, I - anal prolegs, J - prothorax spiracle, K - abdominal spiracle, L - spiracle of abdominal segment VIII (the largest of all).

Some authors report that there may be from 5 to as many as 10 larval instars in the development of G. mellonella. The smallest number (five) of larval stages has so far been reported by Fasasi and Malaka ( 2006), who explain that this quite unusual result is related to the type of food and other optimal conditions of their rearing programme, which required rapid development and thus a smaller number of moults. Other reports, however, including very recent ones, most frequently mention seven (L1-L7) larval instars in the development of this insect (Sehnal 1966 Anderson and Mignat 1970 Swamy 2008 Ellis, Graham and Mortensen 2013 Venkatesh Hosamani et al. 2017 Desai et al. 2019). This is confirmed by accurate measurements (in mm) of body length (l), body width (w) and head capsule width (wh) of L1-L7: l-1.27, 2.40, 4.80, 9.30, 15.50, 21.60 and 25.40, respectively w-0.25, 0.45, 1.26, 1.56, 2.65, 3.30 and 4.86, respectively wh-0.21, 0.32, 0.54, 1.15, 1.28, 1.55 and 2.30, respectively (Hosamani et al. 2017). In this context, the latest metric data regarding the length and width of the body of greater wax moth larvae, recently published by Desai et al. ( 2019), are worthy of attention: l-0.81, 2.10, 5.86, 8.76, 14.24, 19.58 and 23.88, respectively w-0.29, 0.44, 1.11, 1.99, 2.03, 2.54 and 3.55, respectively. These figures differ conspicuously from those given 2 years before. This may have been caused by the different type of artificial food that was used for breeding: a mixture of wheat flour, corn flour, wheat bran, powdered milk, yeast, honey and glycerol. At the larval stage, there are still no external structural features enabling the sex of the future adult form to be determined (Kwadha et al. 2017).


When fully grown, last instar larvae stop feeding and they move vigorously in search of suitable, safe places where they can attach the cocoon and pupate. In active beehives, these are mainly spaces beyond honeycombs (e.g. the outer surfaces of bee frames or the inner surfaces of the hive's walls). In abandoned hives, by contrast, pupal cocoons have been found anywhere within them (G. K. Wagner, oral information). The wooden parts of the hive are often the sites where cocoons are constructed. Fully grown larvae excavate species-characteristic boat-shaped depressions in the wood, which can weaken the entire structure of the affected parts of the hive (Paddock 1918 Ellis, Graham and Mortensen 2013). Having found and excavated a suitable site, the larvae begin to spin a silken pupal cocoon, which they then attach to the eroded cavities. Cocoon construction takes on average 2.25 days, although this depends on the abiotic conditions of the environment (Paddock 1918). The cocoon protects first the larva and then the pupa against worker bees and possible parasites and probably also stabilises the abiotic conditions during pupal development (Jindra and Sehnal 1989 Shaik, Mishra and Sehnal 2017). The outer layer of the cocoon soon becomes hard while the interior remains soft (Ellis, Graham and Mortensen 2013). In the front of the cocoon, the larva makes an exit hole for the future adult. Just before pupation, however, this opening is closed off with a thin layer of silk (Paddock 1918 Desai et al. 2019). Having constructed the cocoon, the slightly shrunken larva becomes inactive a few hours before pupation, passing through a short-lived developmental stage known as a prepupa. As in all Lepidoptera, however, the G. mellonella prepupa is not considered to be a distinct developmental stage because it is not separated from the last larval instar by a moult (Chapman 1998).

The entire developmental phase of the greater wax moth, in which the larva builds a cocoon and then pupates, has been defined as the preparatory period (Hosamani et al. 2017 Desai et al. 2019). During the pupal stage, as in other holometabolous insects, histolysis and phagocytosis of the larval structures take place first, followed by the histogenesis of the imaginary structures that arise from so-called imaginary disks. These are made from embryonic cells that can divide quickly. The whole process is controlled by hormones (Chapman 1998).

Data on the external morphology of the G. mellonella pupa are given in relatively few reports (Paddock 1918 Smith 1965 Swamy 2008 Hosamani et al. 2017 Kwadha et al. 2017 Desai et al. 2019). Most often, these refer only to the general appearance of this developmental stage (e.g. colour, sexual dimorphism) and its dimensions. To date, only Smith ( 1965) has given a detailed account of the external structure of this pupa.

The pupa of the greater wax moth is obtect (i.e. it represents a type in which all the appendages are cemented to the body by means of a special secretion). The colour of the pupa changes with age from white (just after pupation) through yellow and brown to dark brown 4 days later. The body is moderately elongate,

3.1–3.5-fold as long as wide in the widest place. The eyes are large and well visible. The antennae are long, slightly arched in the front, usually extending to the edge of the second pair of wings (hind wings). The pretarsus of the hind legs protrudes slightly beyond the edge of the hind wings (Smith 1965). There are two pairs of short, protruding setae on the parietals, resembling tiny horns. There are two to seven pairs of short setae on body segments. Segments II-VII are each equipped with a pair of active spiracles located on the sides of the body. The ventral side of abdominal segments VIII and IX exhibits well-marked sexual dimorphism: female – the sclerite of segment VIII is separated and segment IX has a single copulatory aperture male – the sclerite of segment VIII is uniform and segment IX has a pair of rounded knobs representing the phallomeres and gonopore between them (Desai et al. 2019). The dimensions of the G. mellonella pupa given in the literature are: length: 11.9–20 mm width: 3.2–7 mm (Paddock 1918 Smith 1965 Swamy 2008 Ellis, Graham and Mortensen 2013 Hosamani et al. 2017 Kwadha et al. 2017 Desai et al. 2019). The respective average dimensions of the female pupa are significantly larger than those of the male pupa: length–15.83 and 11.86 mm width–4.17 and 3.17 mm (Desai et al. 2019).

Depending on the temperature and humidity, the pupal stage in G. mellonella lasts from 8 (at 28°C, 65% RH - relative humidity) to

50 days (from 2.5°C to 24°C, 44% to 100% RH) (Pastagia and Patel 2007 Swamy 2008 Hosamani et al. 2017 Kumar and Khan 2018 Desai et al. 2019).

Emergence of adults

The eclosion of adults from cocoons has been observed at night and late in the evening. As they leave the cocoons, they push out the silk lids covering the cocoon exit holes (Swamy 2008). Once free of the cocoons, the adults remain inactive until their wings are fully extended and hardened. At first, the moths are creamy white (teneral forms), later darkening to a grey colour (Nielsen and Brister 1979 Swamy 2008 Desai et al. 2019). It has frequently been observed that the imagines of G. mellonella prefer dark places, run around in an agitated manner if illuminated and try to hide in various unlit corners of the hive (G. K. Wagner, oral information).

Adults and mating

Adults are incapable of consuming food because their mouthparts are degenerate hence, they do not live very long, from

7 to 30 days, depending on ambient conditions (Paddock 1918 El-Sawaf 1950 Opoosun and Odebiyi 2009 Hosamani et al. 2017 Kumar and Khan 2018). As reported by El-Sawaf ( 1950), males live longer (21–30 days) than females (8–15 days), which have three phases in their lifetimes: pre-oviposition (1.60 ± 0.50 days), oviposition (6.12 ± 1.09 days) and post-oviposition (2.00 ± 0.87 days) (Desai et al. 2019).

Unlike most moths, G. mellonella adults have a unique mating behaviour. Males lure females with a two-component pheromone (n-nonanal + n-undecanal) and in addition emit short pulses of sound at a frequency of 75 kHz, which can play a significant role in the selection of reproductive pairs (Finn and Payne 1977 Greenfield 1981). They generate this acoustic signal using structures found on the wings (Spangler, 1985, 1986). Females react to the sound by fanning their wings (Spangler 1988), although they are unable to locate its source. Sex pheromones, which are released by males in response to female wing movements, help in this, ultimately attracting their partners before mating (Leyrer and Monroe 1973 Spangler, 1985, 1986, 1987, 1988 Jones et al. 2002). Males begin to produce sound impulses after sunset, when the light intensity is near that inside the honey beehive and they are close to or in contact with other wax moths. Interestingly, the sound is never produced in the presence of its natural hosts (i.e. honeybee workers Spangler 1986). The exact mechanism of acoustic signal production in males of the greater wax moth was described by Spangler ( 1986). According to Nielsen and Brister ( 1977), copulation can take place on trees adjoining the apiary, after which only the females return to the hives.

Oviposition and fertility

Egg laying begins within a relatively short time after adults appear and mate (Paddock 1918). Nielsen and Brister ( 1977) observed oviposition

24 h after the appearance of imagines, which continued for 4 consecutive nights. Females usually enter the hive at night, when the bees are already inactive. Attempts by G. mellonella females to get into the hives before evening have also been observed, but then they were attacked by aggressive host workers (Nielsen and Brister 1977). In the hive, the moths seek out various cracks and crevices on honeycombs or other parts of the hive (Charriere and Imdorf 1999), as far as possible from any light source. Having found a suitable place in the hive, the female stretches her abdomen to the maximum, extending the tip as deep as possible. The strategy described above minimises the detection of eggs by bees or possible parasites and increases the survival of the larvae hatched from them (Williams 1997 Ellis, Graham and Mortensen 2013 Kwadha et al. 2017). Hosamani et al. ( 2017) reports that oviposition usually takes place at night, between 19.00 and 03.00 h.

The overall fertility of G. mellonella females can differ widely: this is probably related to the abiotic and biotic conditions (including infections) in which they breed (Mohamed et al. 2014). The number of eggs laid by one female is usually from 500 to 1800 with

60 eggs per day (El-Sawaf 1950 Warren and Huddleston 1962 Hosamani et al. 2017). Much smaller total numbers of eggs (i.e. from 107 to 297) were laid by single females in laboratory conditions (26.7°C, 93.0% RH) (Fasasi and Malaka 2006). Interesting data in this respect were obtained by Mohamed et al. ( 2014), who demonstrated a close relationship between various types of natural food and the fertility and duration of the oviposition period in G. mellonella females in constant breeding conditions (30°C, 50% RH). The lowest (392 eggs, 5.2 days) and highest (1308 eggs, 8.4 days) fertility and oviposition periods were obtained for females reared on an empty new wax comb and an old wax comb with pollen, respectively. As it turned out, however, the type of diet had only a minimal impact on the length of the embryonic development period, which ranged from

10–11 days, depending on the type of food (Mohamed et al. 2014 Kumar and Khan 2018.)

Depending on the temperature, humidity and food resources, the overall developmental period from the oviposition to the appearance of adults ranges from

93 days (2.5–24°C, 44–100% RH, food shortage) (Kumar and Khan 2018). Because this moth usually lives in a fairly stable microenvironment (e.g. hive, warehouse) as regards prevailing abiotic conditions, it can periodically produce from four to six generations per year (Kwadha et al. 2017). Their number and longevity depend on environmental conditions, the most important of which appear to be the temperature and type of food (Mohamed et al. 2014 Kumar and Khan 2018).

Endosome Signaling Part A

Sylvain Loubéry , Marcos González-Gaitán , in Methods in Enzymology , 2014

2.2.3 The day of the experiment

Pupae dissection has been very precisely described by Jauffred and Bellaiche (2012) . Proceed as such, with the only exception that 500 μl Clone 8 medium has to be used as dissection medium (instead of PBS). Once the notum is detached from the animal body, transfer it to 1 ml Clone 8 medium in a silanized hourglass dish to wash away fat bodies. Always use pipette tips that have been cut at the tip while transferring a notum from dish to dish, and pipette gently, so that the tissue sticks neither to the pipette tip nor to glass also take care to aspirate as little medium as possible when doing this, so as not to dilute the receiving solutions.

In a second silanized hourglass dish, mix either: 295 μl Clone 8 medium and 5 μl of the anti-Delta-Zenon mix, or 87.5 μl Clone 8 medium and 12.5 μl of the anti-Notch-Zenon mix. Transfer the notum to this dish and homogenize the solution. Pulse for 5 min.

Wash three times in 1 ml Clone 8 medium, then transfer the notum to a drop of 500 μl Clone 8 medium in a glass-bottom Fluorodish culture dish (World Precision Instruments). The tissue is now ready to be imaged.

Watch the video: Complete Life Cycle of the Indian Meal Moth (August 2022).