Where does grasshopper come from heat

Where does grasshopper come from heat

All grasshoppers begin their lives as eggs. Yet eggs represent the least known stage of the grasshopper life cycle. They are laid in the soil of the habitat and develop hidden from the view of humans. Eggs of a few species, however, have been studied in both field and laboratory (Fig. 9).

Figure 9. One intact and one broken egg pod, exposing the eggs of the migratory grasshopper, Melanoplus sanguinipes (Fabricius).

Incubation of eggs begins immediately after females deposit them in the soil. The embryo, at first a tiny disc of cells laying on the ventral side of the yolk surface and at the posterior end of the eggs (Fig. 10), grows rapidly, receiving nourishment from the nutrient stores in the yolk.

From left to right: Stage 1 (5%) Stage 3 (10%) Stage 7 (20%) Stage 10 (30%) Stage 12 (40%) Stage 19 (50%)

Figure 10. Selected stages in the development of a grasshopper embryo (Melaoplus sanguinipes) held at a constant temperature of 30 C. Left two figures show whole egg; other figures show embryos removed from egg. (Illustrations adapted from Riegert, 1961; stages idetified and designated for embryos of Aulocara elliotti by Saralee Visscher, 1966).

In seven days the embryo of the migratory grasshopper, Melanoplus sanguinipes , held at an incubation temperature of 30½C, reaches Stage 19. In this stage the embryos of many rangeland species such as Aulocara elliotti and Camnula pellucida cease growth and begin a diapause . The embryo of the migratory grasshopper, however, continues to develop and at Stage 20 actively moves from the ventral to the dorsal surface and revolves 180½ on its long axis (see Figure 10, Stage 20). After 15 days the embryo has grown to Stage 24, having achieved 80 percent of its development. It then ceases growth and enters diapause. The embryo of the twostriped grasshopper, and probably others also, enter diapause at this stage. Exposed to favorable incubation temperatures, the eggs of a few rangeland species, such as Arphia conspersa and Xanthippus corallipes , develop completely and hatch during the same summer they are laid. The immediate cause of cessation of embryonic growth (diapause) in eggs of the majority of rangeland grasshoppers appears to be the shutdown of growth hormones. The embryos remain physiologically active as transfer of nutrient materials from the yolk into the embryonic fat body and other tissues continues. Cold temperatures of winter, however, slow or end this process and embryos enter a dormant period.

For eggs laid in temperate regions to reach their maximum development before diapause, they must receive sufficient heat, usually measured as day-degrees of heat accumulated in the soil at egg depth. Eggs deposited late in the season or during a cold summer may not receive this amount of heat, especially in northern areas such as the Canadian provinces of Alberta, Manitoba, and Saskatchewan. Eggs that do not reach their potential stage of development have reduced hatchability the following spring and thus do not contribute as much to the maintenance of a population.

During winter, low ground temperatures eventually break egg diapause. As soon as the ground warms above threshold soil temperatures of 50 to 55½F in spring, the embryos are ready to continue their development. Research has shown that for the few species studied, eggs need 400 day-degrees by fall to attain maximum embryonic growth and another 150 day-degrees in spring to initiate hatching. For completion of embryonic growth from start to finish, eggs require totals of 500 to 600 day-degrees.

In spring the emergence of hatching grasshoppers may be readily observed. All embryos of a single pod usually wriggle out one after another within several minutes. Once out, they immediately shed an embryonic membrane called the serosa. An individual hatchling, lying on its side or back and squirming, takes only a few minutes to free itself (Fig. 11). During this time the hatchlings are susceptible to predation by ants. After the shedding of the membrane the young grasshoppers stand upright and are able to jump away and escape attacking predators. In spring, young grasshoppers have available green and nutritious host plants. The majority of individuals in grasslands are grass feeders, but individuals of some species are mixed feeders, eating both grasses and forbs. Others are strictly forb feeders.

Figure 11. The lifecycle of the bigheaded grasshopper, Alucara ellliotti (Thomas). During summer in bare spots of grassland the female deposits at intervals batches of eggs. As soon as the eggs are laid, they begin embryonic development and reach an advanced stage in which they enter diapause and pass the winter. In spring the eggs complete embryonic devlopment and hatch. The young grasshopper sheds a serosal skin, the exoskeleton hardens, and the nymph begins to feed and grow. After molting five times and developing through five instars in 30-40 days, it becomes an adult grasshopper with functional wings. The adult female matures groups of six to eight eggs at a time and deposits them in the soil at intervwls of three to four days for the duration of her short life.

As insects grow and develop, they molt at intervals, changing structures and their form. This process is called metamorphosis . A number of insects undergo gradual (simple) metamorphosis, such as grasshoppers. With this type of metamorphosis the insect that hatches looks like the adult except for its smaller size, lack of wings, fewer antennal segments, and rudimentary genitalia (Fig. 11). Other insects with gradual metamorphosis include the true bugs, aphids, leafhoppers, crickets, and cockroaches. The majority of insects undergo complete (complex) metamorphosis, as the eggs hatch into wormlike larvae adapted for feeding and have a vastly different appearance from that of the adult insect. Before full-grown larvae can become adult insects they must enter into the pupal stage. In this stage they develop and grow the adult structures. Common examples of insects that undergo complete metamorphosis are beetles, butterflies, bees, wasps, and flies.

For young grasshoppers to continue their growth and development and reach the adult stage, they must periodically molt or shed their outer skin (Fig. 11). Depending on species and sex, they molt four to six times during their nymphal or immature life. The insect between molts is referred to as an instar; a species with five molts thus has five instars. After shedding the serosal skin, the newly hatched nymph is the first instar. After each molt the instar increases by one so that the nymph consecutively becomes a second, third, fourth, and fifth instar. When the fifth instar molts, the grasshopper becomes an adult or an imago.

The new adult has fully functional wings but is not yet ready to reproduce. The female has a preoviposition period of one to two weeks during which she increases in weight and matures the first batch of eggs. Having mated with a male of her species, the female digs a small hole in the soil with her ovipositor and deposits the first group of eggs. Once egg laying begins, the female continues to deposit eggs regularly for the rest of her short life. Depending on the species, production may range from three pods per week to one pod every one to two weeks. The species that lay fewer eggs per pod oviposit more often than those that lay more eggs per pod.

The egg pods of grasshoppers vary not only in the number of eggs they contain but also in their size, shape, and structure. Based on structure, four types have been recognized. In type I a stout pod forms from frothy glue and soil surrounding the eggs; froth is lacking between the eggs. In type II a weaker pod is formed from frothy glue between and surrounding the eggs. In type III frothy glue is present between the eggs but does not completely surround them. In type IV only a small amount of froth is secreted on the last eggs of a clutch, and most of the eggs lie loosely in the soil. Grasshopper eggs themselves vary in size, color, and shell sculpturing. Depending on the species eggs range from 4 to 9 mm long and may be white, yellow, olive, tan, brownish red, or dark brown. Eggs of certain species are two-toned brown and tan.

Events in the life cycle of an individual species of grasshopper — hatching, nymphal development, and adulthood — occur over extended periods. The eggs may hatch over a period of three to four weeks. Nymphs may be present in the habitat eight to ten weeks and adults nine to 11 weeks. Because of the overlapping of stages and instars, raw field data obtained by sampling populations do not answer several important questions. For example, how many eggs hatched? How many individuals molted successfully to the next instar? What was the average duration of each instar? How many became adults? What was the average length of life and the average fecundity of adult females? To obtain answers to these questions, detailed sampling data must be treated mathematically.

Laboratory data may also be used in studying grasshopper life histories. Table 4 provides information on the life history of the migratory grasshopper, Melanoplus sanguinipes , reared at a constant temperature of 86½F and 30-35% relative humidity and fed a nutritious diet of dry feed, green wheat, and dandelion leaves. The entire nymphal period averages 25 days for males and 30 days for females. Each instar takes four to five days to complete development except for the last instar, which takes seven days. Adult longevity of males averages 51 days and females, 52 days. Longevity of adults in the field is no doubt briefer because of the natural predators and parasites cutting short the lives of their prey.

TABLE 4. Life history of the migratory grasshopper, Melanoplus sanguinipes, reared in the laboratory at a constant temperature of 86.5 F.


Where does grasshopper come from heat

Texas Cooperative Extension, Texas A&M University, College Station, Texas

Grasshoppers: Frequently Asked Questions, 2003

Dr. Allen Knutson,
Professor & Extension Entomologist, Texas Cooperative Extension

hy are grasshoppers so bad this year, again? Consecutive years of hot, dry summers and warm, dry autumns favor grasshopper survival and reproduction. Warm, dry fall weather allows grasshoppers more time to feed and lay eggs. The large numbers of grasshoppers present last fall left many eggs in the soil which hatched this spring. Also, rains in the spring when eggs are hatching drown young hoppers. Thus, dry weather in the spring favors their survival.

Grasshopper, Brachystala magna

Where do grasshoppers come from?
Grasshopper eggs are deposited in the soil l/2 – 2 inches deep in weedy areas, fencerows, ditches and hay fields. The eggs hatch in the spring and early summer. Eggs of different grasshopper species hatch out at different times, so young grasshoppers can be seen throughout the spring and early summer. Young grasshoppers, called nymphs, feed for about six weeks. Once nymphs reach the adult stage, they can fly. As weedy plants are consumed or dry in the summer heat, adult grasshoppers can fly from weedy areas and pastures to more succulent crops and landscapes.

When will grasshopper numbers decrease this season?
Although grasshoppers complete only one generation a year, eggs hatch over a long period of time. Development from egg to adult requires about 40-60 days. Also, eggs of different species hatch at different times so small grashoppers can be found throughout the growing season. Grasshopper can persist until late fall when old adults begin to die or when a killing frost occurs.

What can be done to reduce their numbers?
Weed control. Eliminating weeds will starve young hoppers and later discourage adults from laying eggs in the area. Destroying weeds infested with large numbers of grasshoppers can force the hungry grasshoppers to move to nearby crops or landscapes. Control the grasshoppers in the weedy area first with insecticides or be ready to protect nearby crops if they become infested. Grasshoppers deposit their eggs in undisturbed soil, as in fallow fields, road banks, and fence rows. Shallow tillage of the soil in late summer may be of some benefit in discouraging egg lay.

Are insecticides effective?
Grasshoppers are susceptible to many insecticides. However, insecticides typically do not persist more than a few days and grasshoppers may soon re-invade the treated area. The length of control will depend on the residual activity of the insecticides and the frequency of retreatment. Controlling grasshoppers over a large area will reduce the numbers present which can re-infest a treated area. Dimilin 2L provides long residual of young hoppers but is not effective against adults.

When should insecticides be applied?
Monitor grasshopper infestations and treat threatening infestations while grasshopper are still small and before they move into crops and landscapes. Immature grasshoppers (without wings) are more susceptible to insecticides than adults.

What about insecticide baits for grasshopper control?
Sevin 5 Bait is a ready-to-use bait which can be applied to many crop and non-crop sites, including around ornamentals and many fruit and vegetable crops. For those wanting to make their own grashopper bait, the labels for Sevin XLR and Sevin 4-Oil ULV provide directions for mixing these products with cereal grains to make a 2% to 10% carbaryl bait. The bait is labeled for use in rangeland, wasteland, ditch banks and roadsides. The label further states the bait is for use “only by government personnel or persons under their direct supervision (e.g. USDA, state and local extension personnel, etc.)”

Are biological control products such as Nolo Bait, Grashopper Attack, and others effective?
These products contain spores of a protozoan called Nosema locustae, formulated in a bait. Grasshoppers consuming the bait become infected by the Nosema organism. Some immature grasshoppers die while adults often survive but females lay fewer eggs. Nosema baits act too slowly and kill too few grasshoppers to be much value when the need for control is immediate.

Some insecticides for controlling grasshoppers in the home landscape at present (2003) include:

  • Cyfluthrin. The active ingredient in Bayer Advanced Home and Garden Spray
  • Bifenthrin. Active ingredient in Ortho Ready-to-Use Houseplant and Garden Insect Killer
  • Permethrin. Active ingredient in Spectracide and other products.
  • Acephate. Active ingredient in Orthene (at present, but being phased out).

Note: Tempo (cyfluthrin) and Demon (cypermethrin) are labeled for use by Professional Pest Control Operators (2003) for insect control in lawns and landscapes.

What insecticides can be used in ornamental production?
Several products may be used including those containing bifenthrin, lambda-cyhalothrin, diazinon, dimethoate, malathion, acephate and carbaryl (2003).

For more information, see: Extension bulletin L-5201, Grasshoppers and Their Control.


Keeping Insects

Caring for a praying mantis, butterflies, stick insects and beetles


Whether or not you want to keep grasshoppers as pets or as food insects for your reptile, mantis or other pet, this page is the place for you. Here you can find how to take care of grasshoppers and locusts, with a special focus on the “common” pet grasshopper species Locusta migratoria and Schistocerca gregaria. You will also learn how to breed them.

A grasshopper nymph. You can see it is not yet adult because of the short underdeveloped wings.

Grasshoppers fall in the order of Orthoptera, in the suborder Caelifera. Grasshoppers are sometimes called locusts. There are many species, the most famous ones are the desert grasshoppers that also occur in the bible as one of the plagues. Both species Locusta migratoria and Schistocerca gregaria are desert grasshoppers and are in the family Acrididae. Both species reach a size of around 7 cm in length.

Development and life cycle of grasshoppers

A female grasshopper lays eggs in small egg clusters, usually in the ground but she can also deposit them in plant material. Young grasshoppers are called nymphs and already look like miniature versions of the parents. Nymphs lack wings, only adult grasshoppers have fully functional wings. Grasshopper nymphs grow fast and shed their skin (molt) around 8 times in the process. At the last molt both males and females grow long wings that pass the abdomen.

It takes around 2 – 3 weeks for eggs to hatch. In around 4 weeks the nymphs reach adulthood. Around two weeks after that they begin to mate and produce eggs. If the temperature is low, development and breeding will slow down too. A grasshopper will die of old age when it has been an adult for around 5 months.

Schistocerca gregaria nymph

Housing your grasshoppers

Housing grasshoppers is easy. You need a container that is big enough, has some ventilation and can be closed properly to prevent escape. Grasshoppers can chew through fabric gauze, so net cages or cages with a fabric cover are not suitable for grasshoppers. A fauna box, a glass terrarium or a plastic terrarium with metal mesh for ventilation will do. If you keep the grasshoppers as pets, a glass terrarium with a mesh lid will look good. If you keep the grasshoppers to feed them to reptiles or praying mantises you a plastic container is more practical, as it is lightweight and cheaper. Make sure the container is big enough for all grasshoppers. Twelve adult grasshoppers need a cage of around 50 x 50 x 30 cm at least. Bigger is always better.

Fill the bottom of the container with dry sand, dry oatmeal flakes or dry coconut fibers. Place some dry twigs or branches in the enclosure to provide extra surface for the grasshoppers to sit on. The food of the grasshoppers, grass and/or leaves, will also serve as decoration and perching areas. Make sure light reaches into the container, either by a light bulb (see next section about Temperature) or by natural light. Direct sunlight shining into the enclosure could heat it up too much, make sure to prevent overheating.

Schistocerca gregaria nymph seen from top

Temperature and humidity

The grasshopper species Locusta migratoria and Schistocerca gregaria are desert species. They need a dry and warm environment to thrive. A too humid environment will result in infections and death of the grasshoppers. A too cold environment will slow down development and make breeding grasshoppers impossible.

Keep the temperature during the day between 25 and 35 degrees Celsius. By night you can allow the temperature to drop to 15 degrees Celsius. The best way to heat the enclosure of grasshoppers is with a regular light bulb. It is also possible to heat the terrarium with a species heat bulb, found in reptile-specialized pet shops. You can heat the enclosure with a heat mat too. More information about heating any insect enclosure can be read at our page Temperature.

Keep the humidity low by placing dry bedding in the enclosure (dry coconut fiber, oatmeal flakes or dry sand) and not spraying with water. Grasshoppers do need moisture to survive, but can get this from their food. Lightly spray fresh food with water before feeding it to your grasshoppers. If you feel like the enclosure is getting moist, for example when you have an enclosure with little ventilation or if the enclosure is placed in a room with high air humidity, then you can better skip the spraying of the food. The locusts will get all their moisture from the fresh plant material that you give them.

Locusta migratoria nymph

Food and feeding grasshoppers

Locusta migratoria and Schistocerca gregaria eat only plant material. The best food and easiest food you can give them is fresh grass. Even better is fresh reed, reedgrass or canary grass (Phalaris arundinacea) if available. Fresh wheat leaves, corn leaves and other vegetable plants may also be eaten. But actually, many plant species will be eaten by grasshoppers. You can try to feed them any kind of grass-like species; if they eat the plants, it means that this is a suitable plant species. Generally grasshoppers will refuse to eat any poisonous plants. Be very aware of insecticide, if any plant has been spraying with insecticide it will be deadly to your grasshoppers.

Place the food plants inside the enclosure of the grasshoppers. They will start to eat from it instantly. At some point the plant will be too dry to eat, then the grasshoppers should be fed again with fresh plants. Once in a while you should clear out all old dry plant material from the enclosure.

Schistocerca gregaria nymph

Breeding grasshoppers

It is very easy to breed grasshoppers as long as you keep them in the right circumstances. They really need high temperature, low humidity and plenty of fresh food. If you have males and females, breeding will occur naturally. You don’t need to move the eggs or nymphs to a different container. If you like, you can of course. The nymphs are very small, so you might want to keep them in a different container than the parents to cater to their needs, prevent escaping and keep a better eye on them.

If you really keep grasshoppers well, you will be awarded with a lot of young grasshoppers. Make sure this is actually what you want! If you don’t want to breed with the grasshoppers, remove the bedding with the eggs or collect the eggs and place them in the freezer. This will kill the eggs before they start to develop. Never release grasshoppers into nature. They can cause plagues, disrupt nature and compete with native grasshopper species.

Locusta migratoria nymph

Buying grasshoppers

It’s actually very easy so buy grasshoppers. You can buy them in reptile-oriented pet shops. They are sold there as feeder animals. They are sold in different sizes, from small nymphs, to bigger nymphs and to adults. If you want to keep them for fun, you can better buy the nymphs. If you want to breed them, it’s faster to just buy the adults.

Make sure the grasshoppers look healthy when you buy them. There should not be any dead grasshoppers in the container.

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What triggers colour change? Effects of background colour and temperature on the development of an alpine grasshopper

J. Pablo Valverde

Department of Evolutionary Biology, Bielefeld University, Morgenbreede 45, 33615, Bielefeld, Germany

Holger Schielzeth

Department of Evolutionary Biology, Bielefeld University, Morgenbreede 45, 33615, Bielefeld, Germany


Colour polymorphisms are a fascinating facet of many natural populations of plants and animals, and the selective processes that maintain such variation are as relevant as the processes which promote their development. Orthoptera, the insect group that encompasses grasshoppers and bush crickets, includes a particularly large number of species that are colour polymorphic with a marked green-brown polymorphism being particularly widespread. Colour polymorphism has been associated with the need for crypsis and background matching and background-dependent homochromy has been described in a few species. However, when and how different environmental conditions influence variation in colour remains poorly understood. Here we test for effects of background colour and ambient temperature on the occurrence of colour morph switches (green to brown or brown to green) and developmental darkening in the alpine dwelling club-legged grasshopper Gomphocerus sibiricus.

We monitored individually housed nymphae across three of their four developmental stages and into the first week after final ecdysis. Our data show an absence of colour morph switches in G. sibiricus, without a single switch observed in our sample. Furthermore, we test for an effect of temperature on colouration by manipulating radiant heat, a limiting factor in alpine habitats. Radiant heat had a significant effect on developmental darkening: individuals under low radiant heat tended to darken, while individuals under high radiant heat tended to lighten within nymphal stages. Young imagoes darkened under either condition.


Our results indicate a plastic response to a variable temperature and indicate that melanin, a multipurpose pigment responsible for dark colouration and presumed to be costly, seems to be strategically allocated according to the current environmental conditions. Unlike other orthopterans, the species is apparently unable to switch colour morphs (green/brown) during development, suggesting that colour morphs are determined genetically (or very early during development) and that other processes have to contribute to crypsis and homochromy in this species.


Colour polymorphism has fascinated biologists since the time of Darwin, and its evolutionary meaning is still being revealed [1–3]. Colour polymorphism, defined here as within-species phenotypic variation, occurs throughout the animal kingdom in several taxa of birds, fish, mammals, frogs, molluscs, spiders, several insect orders and also in plants [4–9]. The occurrence of colour polymorphisms in natural populations can result from biased mutation, pleiotropy and trade-offs, gene flow, spatially and/or temporally fluctuating selection and negative frequency-dependent selection that can counter loss of variation by genetic drift [10–13]. Furthermore, developmental plasticity and phenotypic flexibility, if they do not invoke significant cost, might allow the maintenance of polymorphisms. This can be particularly advantageous in unpredictably variable environments.

Insects offer a multitude of examples for the coexistence of two or more colour morphs in groups such as grasshoppers, mantoids, cicadids, damselflies, lepidopterans and beetles [13–15]. There is ample evidence for genetic and environmental effects, as well as genotype-by-environment interactions in colour determination [14–18]. Several species appear capable of modifying their colour in response to various environmental cues such as temperature, predation threats, behaviour stimulus (e.g., crab spiders which try to blend with their background to ambush prey, [6]), among others [19]. Within Orthoptera, colour polymorphism is present in dozens of species (reviewed in [15], see also [19, 20]). Two particularly eye-catching forms of colour polymorphism in orthopterans are a widespread green-brown polymorphism in grasshoppers and bush crickets and the famous phase polymorphism in locusts [14, 21]. Phase polymorphism is triggered by changes in population density which induces changes in colour (typically black patterns in gregarious versus pale green or brown colours in solitary phases) as part of more complex changes in morphology, physiology, behaviour and life history [22–26].

The green-brown polymorphism is far more widespread among orthopterans than phase polymorphism and does not correlate with obvious changes in morphology and/or behaviour. Many families and genera of orthopterans comprise species that display either green or brown morphs, while other species are polymorphic (e.g., in genera Decticus, Metrioptera, Oedaleus). In some species, one of the morphs is very rare (such as brown morphs in Decticus verrucivorus), while in others the ratios are far more even (as in Metrioptera roeselii) [14]. With respect to environmental effects, green morphs seem to develop primarily under high humidity, while brown morphs are favoured under dry environmental conditions [15, 27, 28]. Besides the two very striking forms of colour polymorphism mentioned before, there is a range of more fine-scaled within-species variation in colour pattern and colouration among orthopterans [29, 30]. Groundhoppers, for example, differ substantially in their colour patterns, which can be categorized into variable numbers of discrete morphs [31, 32]. In other species, differences in colour are more continuous such as with colouration of species in the genus Oedipoda. Such fine-scaled variation seems to be partly under genetic, partly under environmental control [15, 16, 33]. Many species also show occasional pinkish, purple, yellow or blue colour morphs [34], further illustrating the diversity of colour in orthopterans.

A very interesting phenomenon associated with colour polymorphism is homochromy, which describes matching of body colouration with variation in the background pattern of the local habitat [14, 15, 20, 35]. Such matching might arise for four different reasons: (i) local adaptation due to multi-generational history of selection on genetic polymorphisms, (ii) selective mortality within generations, (iii) individual-level choice of matching habitat patches [36], and (iv) developmental plasticity of body colouration to match local conditions [14]. Developmental switches are particularly intriguing, because they allow individual-level matching, which is likely advantageous if habitats are unpredictably variable across generations, but predictable from environmental cues over the lifetime of individuals. Developmental matching has been reported in orthopterans for species with fine-scale variation in colour pattern [37, 38], but also for species which present the green and brown colour polymorphism (Table 1 ).

Studies on the effects of background colouration on the occurrence of colour morph switches in green-brown polymorphic (upper section) and other polymorphic (lower section) orthopterans

Proportion of Switches
Species Background Material matched non-matched matched non-matched Stages with Switches Reference
a) Green-Brown Polymorphism
Acrida turrita fresh/dry grass 62 145 4 % 86 % after ecdysis [48]
Acrida turrita painted sawdust 14 15 0 % 100 % after ecdysis [48]
Acrida turrita painted sawdust 32 0 % imago [48]
Oedaleus decorus NA 10 66 0 % 88 % after ecdysis [33]
Schistocerca americana coloured paper 74 0 % across stages [47]
Schistocerca gregaria coloured paper 58 312 0 % 63.5 % within stages [68]
Locusta migratoria coloured paper 58 237 69 % 24 % across stages [69]
Rhammatocerus habitat background 3000 > 90 % > imago [28]
Chorthippus biggutulus coloured paper 257 30 % across stages [70]
b) Other Polymorphisms
Oedipoda sp. earth, clay, coal, stone, chalk 12 92 0 % 80 % after ecdysis [37]
Tetrix subulata sand 312 16 % across stages [49]
Tetrix ceperoi sand 228 16 % across stages [49]

The proportion of switches was calculated for various studies based on multiple assays either on matched or non-matched background colour. Studies do not indicate precise time of colour morph switch occurrence, only final results of repeated colour assessments across nymphal stages are stated (in the case of Ergene all switches occur after an ecdysis event). Percentages reflect amount of indiv >

Orthopterans are preyed upon by a large diversity of species, including birds, lizards, amphibians, spiders and other insects and are frequently parasitized by parasitic flies and mites [24, 34]. Visually hunting predators might constitute a force that can favour homochromy and crypsis, since survival to the imago stage is critical to individual fitness. Predators might also impose frequency-dependent selection if they develop search images and preferentially prey upon the most common morphs [9]. However, there are other influences that might affect body colour and this may or may not be in conflict with crypsis. For example, body colour is likely to affect the absorption of radiant heat and therefore play an important role in thermoregulation [13, 39–41]. It has repeatedly been reported that orthopterans raised under cool conditions are darker than those raised under warm conditions [16, 19, 42–44].

The club-legged grasshopper Gomphocerus sibiricus is a highly sexually dimorphic alpine dwelling grasshopper that exhibits the green-brown polymorphism present in many other orthopterans. Green individuals are rarer than brown morphs in most populations. Despite substantial fluctuations in population density [24, 45], the species does not show any typical phase polymorphism [26]. It inhabits alpine pastures and grassland with a very heterogeneous composition of open terrain strewn with stones and mottled by various types of short grasses and herbaceous plants. Climate conditions in the mountains are very unpredictable and variable within and between years. The maintenance of the green-brown colour polymorphism could be aided by the heterogeneous habitat and/or temporal variability in climate conditions in the native habitat of G. sibiricus.

In the present study we aimed to test the effect of two known factors on developmental colour changes in G. sibiricus. First, we assessed the effect of background colour (green or brown) on colour morph development across almost the entire ontogeny. We were particularly interested in whether individuals are able to switch colour morphs to achieve homochromy as it has been described in other species (Table 1 ). We predicted that if individuals were able to switch colour morphs, then individuals whose colour morph mismatched the background colour would be capable of matching their background at an advanced developmental stage. Second, we assessed the effect of temperature by means of a radiant heat treatment on developmental darkness, while controlling for humidity, population density and food moisture content. Here we predicted that if individuals were capable of manipulate the degree of melanin in their cuticle, thermoregulation needs would promote a colouration darkening under conditions of low radiation. We followed individuals from the second nymphal stage through to the imaginal stage during two independent rounds of trials with two different radiant heat regimes. Individuals were exposed to experimental treatments from the second nymphal stage onwards. The long exposure to experimental conditions allowed us to evaluate if colour changes occur exclusively in connection with moults or if changes were possible even within nymphal stages.

A total of 78 indiv >2 = 5.05, df = 1, p = 0.025).

Colour morph switches

No colour morph switch was observed among the 34 individuals in cages with matched background colours (6 green individuals in green cages and 28 brown in brown cages in total for both rounds). Forty-four individuals (33 brown and 11 green) were exposed to unmatched backgrounds, but no colour morph switches were observed among these 44 individuals. Reasoning based on binomial sampling (see methods section) suggests that if G. sibiricus is capable of switch colour, the rate of colour morph switches is well below values reported in previous studies (c ≤ 0.07, Table 2 ). When we concentrate on the subset of the data where colour morph switches were most likely, given both non-matched background and temperature treatment (brown individuals on a green background under high radiant heat treatment), the probability of colour morph switch is still well below expectations (c ≤ 0.17, Table 2 ).

Number of individuals of G. sibiricus on mismatched backgrounds, all of which did not switch colour morph during development

Round 1
Morph & Background Temperature
High Low Sum
Green on Brown n = 3 n = 0 n = 3
c ≤ 0.63 NA c ≤ 0.63
Brown on Green n = 9 n = 9 n = 18
c ≤ 0.28 c ≤ 0.28 c ≤ 0.15
Sum n = 12 n = 9 n = 21
c ≤ 0.22 c ≤ 0.28 c ≤ 0.13
Round 2
Morph & Background Temperature
High Low Sum
Green on Brown n = 4 n = 4 n = 8
c ≤ 0.53 c ≤ 0.53 c ≤ 0.31
Brown on Green n = 7 n = 8 n = 15
c ≤ 0.35 c ≤ 0.31 c ≤ 0.18
Sum n = 11 n = 12 n = 23
c ≤ 0.24 c ≤ 0.22 c ≤ 0.12
Round 1 + 2
Morph & Background Temperature
High Low Sum
Green on Brown n = 7 n = 4 n = 11
c ≤ 0.35 c ≤ 0.53 c ≤ 0.24
Brown on Green n = 16 n = 17 n = 33
c ≤ 0.17 c ≤ 0.16 c ≤ 0.09
Sum n = 23 n = 21 n = 44
c ≤ 0.012 c ≤ 0.13 c ≤ 0.07

Table depicts number of indiv >

Temperature-cued colouration darkening

We observed a significant change in colouration darkness in both rounds of trials. Individuals in the second and third nymphal stages (N2 and N3) of the round 1, and in the N2 and N4 stages of the round 2 experienced a change in darkness which depended on the direction of the temperature treatment. Individuals in the low radiant heat treatment became darker in colour, while those in the high radiant heat treatment became lighter in colour (Table 3 , Fig. 1 ). Individuals in the N3 stage in the R2 did not show a significant change in colour, yet the sign of the point estimate is the same as in stages N2 and N4. Individuals in the N4 stage of the R1 and under a high radiant heat treatment got significantly darker with age. Most of the low radiant heat treatment individuals from the R1 had perished, with the single remaining individual becoming lighter. Unlike the situation in nymphal stages, individuals in the imago stage undergo a darkening in colour in both treatments within the first week after final ecdysis (Fig. 1 ).

Results of the random-slope mixed effects model used to test for effects of radiant heat treatment and larval age on colouration darkness in both green and brown coloured morphs

Round 1

Changes in darkness across nymphal stages exposed to two different sets of radiant heat regimes (first round R1, and second round R2). In each set of panels, upper panels show low radiant heat treatments, bottom panels show high radiant heat treatments. Each line represents the trajectory of an individual (some individuals have only one observation per nymphal stage, and therefore appear only as a dot without line). Brown lines represent brown morphs, while green lines represent green morphs. Nymphal stages are measured in days since the start of each nymphal stage. Darkness has four levels, with 1 representing the lightest colour morph and 4 representing the darkest colour morph

The analysis of unambiguous cases of lightening or darkening using Fisher’s exact tests confirmed a significant colour change for the N2 and N4 stages in the R2 (N2 stage: n = 14, p = 0.023; N3 stage: n = 8, p = 0.14; N4 stage: n = 10, p = 0.015; IM stage: n = 12, p = 0.47, see also Fig. 1 ). Yet after exclusion of ambiguous cases, there was not significant effect in the R1 (N2 stage: n = 12, p = 0.091; N3 stage: n = 7, p = 0.28; N4 stage: n = 12, p = 0.33, see also Fig. 1 ).

The comparison of treatments per observation time (early or late observations within nymphal stages) also confirmed a significant effect of the treatment in the colour of nymphae (upper p values in Fig. 2 ). Individuals in the N2 stage in both rounds, and in the N3 stage of R1 show a significant difference in colouration darkness at the end of the nymphal stage. Individuals of the N4 stage differ at the start of the nymphal stage, but at the end colouration darkness had converged.

Mean darkness per nymphal stage at the earliest and latest observation point, separated per radiant heat treatment for two different sets of radiant heat regimes. Upper panel shows the first round (R1), lower panel shows the second round (R2). White bars represent low radiant heat treatments, grey bars represent high radiant heat treatments. Numbers inside the bar show the number of individuals per observation point. P-values for two-sample t-tests comparing treatments within observation points are shown on top (labeled as “Early” and “Late”). P-values for paired t-tests comparing observation points within treatments are shown below (labeled as “High” and “Low”)

Finally the comparison between the earliest and latest observations per treatment within nymphal stages also found a significant effect of the treatment in the colouration darkness of nymphae (lower p values in Fig. 2 ). Individuals in the N2 stage of both rounds (statistically significant only in the low radiant heat treatment), N3 stage of R1 (statistically significant only in the high radiant heat treatment) and N4 stages of R2 change their colouration darkness within nymphal stages. The colour change in the IM stage is also significant and the change in both groups is towards a darker coloration.


We here show the effect of background colour and temperature on colour changes and darkening in G. sibiricus. There were no colour morph switches among 78 individuals tested, neither when exposed to matched nor to unmatched background. Colour morph switches have been widely reported in other orthopterans (Table 1 ), hence it is interesting that we cannot confirm this for our species. If this species is capable of switching colour at all, the probability of colour morph switch in individuals that mismatched their cage background was certainly very low for both rounds of trials (≤7 %). Additionally, we find that the amount of radiant heat affected colouration darkening within nymphal stages, with darkening at low amounts of radiant heat and lightening at high amounts of radiant heat. Imagoes tended to darken within the first week after final ecdysis independent of radiant heat treatment, suggesting that imagoes may face different life-history trade-offs than nymphae.

Green-brown switches

Colour morph switches have been previously reported, with several observations of the phenomenon in a diverse array of species (Table 1 ). Several of the species for which colour morph switches have been reported are members of the Acrididae (the family that also includes Gomphocerus), but none of them is a member of the subfamily Gomphocerinae [46]. It is possible that the capability for colour morph switches has been lost somewhere in the branch of Gomphocerinae, but this remains speculative in the absence of information about other species. The lack of a mechanism for switching colour during nymphal stages in G. sibiricus may also be due to the really fine-grained structure of the habitat inhabited by the species. It would be very costly for an individual to move at all within the matrix of colours of the habitat if this would require an active colour switching to match their background, even if this could be done in a relatively short time window.

Background colour was visually perceivable to the developing individuals and based on previous studies, we assume visual perception to be the main input that triggers colour change to match the background (see [14] and references therein). However, it might be argued that our coloured paper was not of the right kind for triggering colour changes. Previous studies have used a large diversity of materials for the background manipulation, such as stones, sawdust, sand, coal, clay, paper and paint (Table 1 ). These different materials have typically elicited colour morph switches, which suggests that the effect does not depend on the exact kind of materials used as background. We consider it unlikely (albeit possible) that our paper type was so substantially different from previously used materials that it would not be suitable for triggering colour switches.

In previous experiments, individuals have been tested for different periods of time in order to assess colour morph switches, usually starting the experiments at early nymphal stages [27, 32, 47–49]. Colour morph switches have typically been reported to occur across nymphal stages, often quantified a few days after ecdysis, though detailed information on the exact timing of switches is usually lacking. The only exception to colour morph changes occurring within nymphal stages are changes to black colouration, which have been reported to occur during the imaginal stage [14]. We started with our experimental treatment very early in the life of the grasshoppers (also in comparison with previous studies), giving scope for switches within and/or between developmental stages. Therefore it is rather unlikely that the duration of the treatment prevented colour switching.

Switches might also have been expected due to the radiant heat treatment, since brown individuals tend to be darker on average than green individuals and we would expect them to be better able to heat up if radiant heat is limiting. High temperature, as well as high humidity, high food moisture content and low individual density, are known to drive green body colouration in grasshoppers [14, 15]. We expect that in our experiment high radiant heat conditions would have served to cue individuals of a green habitat. High alpine habitats are typically characterized by high humidity regimes due to high condensation of air humidity at night, which is available as dew drops early in the mornings, and also due to high precipitation regimes [50]. Under these conditions, high temperature and high humidity will promote vegetation growth and produce greener habitats than conditions of low temperature, where vegetation would have weaker growth. Such conditions could have promoted colour morph switches from brown to green, possibly as a means for habitat matching. In contrast, low radiant heat conditions would have served to cue individuals of a browner habitat with less flourishing vegetation. These conditions, in the case of green individuals, could have promoted colour morph switches from green to brown, either for habitat matching and/or to improve thermoregulatory capacity.

Green individuals on a brown background and under the low radiant heat treatment thus constitute the subgroup for which colour morphs switches from green to brown appear particularly advantageous. It is possible that the combination of relative high humidity, high food moisture content and individual housing (i.e., low population density) counteracted the effect of the background, hampering the occurrence of the colour morph switch [14, 15]. This is different for brown individuals on a green background and under the high radiant heat treatment, since for those individuals the combination of low density, high humidity, background mismatching and no need for improved heat absorption are all expected to favour colour morph switches towards green. Yet none of the 16 individuals under this suit of conditions switched colour.

Our limited sample size does not allow us to exclude the possibility that G. sibiricus is capable of colour morph switches under some conditions. Still it strongly points against frequent, general developmental switches in response to background colour and temperature. We had expected that if colour morph switches occurred in G. sibiricus, they would occur at the nymphal stages, given that matching the habitat background is expected to improve survival in natural conditions. It is possible that nymphs of G. sibiricus achieve homochromy even in the absence of developmental switches by actively seeking out matching (micro)habitats [36]. Habitats of G. sibiricus are spatially highly heterogeneous and this might distinguish them from many of the other species that show developmental colour morph switches. Microhabitat variability might favour behavioural over developmental homochromy, while more global (temporal) variability in less structured habitats might favour developmental switches.

Colouration darkening

While the temperature treatment in our experiment did not elicit colour morph switches, it elicited a more subtle response in colouration, causing darkening under low radiant and lightening under high radiant conditions. Radiant heat can limit behaviour of individuals by hampering thermoregulation and this in turn can constrain activity levels, growth and development, and ultimately fitness [51–53]. An increase in the amount of melanin under the cuticle surface would improve thermoregulatory capability due to a difference in heat absorbance between black and brown or green colours, and this would help counteract the effect of a short window of radiant heat exposure. Hence the darkening that we found in the low heat treatment is in line with what would be expected for improved thermoregulation [39–41].

The lightening in colouration in response to the high radiant heat treatment can result from a trade-off between melanin as a colour pigment and other functions of melanin. Melanin plays a role in several functions in insects, such as immune defence, integumental colouration, wound healing and cuticle sclerotisation, among others (reviewed in [54]). It has also been documented that melanin production in insects can be costly, mostly because of the many possible functions of melanin, but also because of dietary limitations of melanin precursors or lack of enzymes necessary to process precursors [51, 55–57]. Therefore lightening of colouration can be seen as an option to avoid investing melanin in body colouration when it is not necessary for absorbing more radiant heat. This reasoning might give an adaptive explanation for the lightening in our high radiant heat treatments.

A change in darkness within nymphal stages implies a mechanism which allows individuals to adjust the amount of visible melanin in their epidermis during the relatively short time spanned between moults. Such a mechanism would include cells at the epidermis capable of spreading pigment granules under the cuticle surface, but also capable of withdrawing the pigment granules under proper stimulation [19, 58]. Relatively little is known about the physiology of colour changes in grasshoppers, but different physiological and morphological mechanisms have been described in other arthropods.

A very intuitive mechanism which could explain the changes in darkness under different radiant heat regimes is pigment dispersal and concentration within chromatophores [19, 59]. This type of cell is known to be present in several taxa, such as fish, reptiles, amphibians, crustaceans and bacteria [60]. The shape of chromatophores is typically highly branched, allowing for pigment to disperse to the branches or contract to the centre to achieve colour change [19, 59, 61]. Another physiological mechanism involved in colour change is granule migration. Here granules of pigment are transported along microtubules which are perpendicular to the cuticular surface, and which branch distally, allowing the pigments to spread and therefore causing colour change. A striking example of this plastic mechanism is observed in the temperature-controlled daily changes in the colour of the chameleon grasshopper Kosciuscola tristis. In this case granules of pigment migrate from the epidermis when the grasshopper is exposed to temperatures above 25 °C, giving males a bright turquoise colouration [19]. A similar mechanism is used in the stick insect Carausius morosus [62]. The colour change that we found in G. sibiricus is much slower, but since granule migration might simultaneously explain changes in darkness in both directions, it might contribute to our observations.

The unpredictable climate in the habitat of G. sibiricus, characterized by long spans of time with few favourable climatic conditions, might explain the occurrence of a darkening and lightening mechanism. In this environment, given the duration of each nymphal stage (about one week, albeit likely to be substantially longer in the field) nymphae may need to adjust colouration even within nymphal stages to be able to cope with climatic variability. In species dwelling in habitats where radiant heat is a limited factor, energy balance and thus activity levels during early developmental stages could be hampered by limited sun exposure conditions. Being able to adjust colouration darkness could greatly improve the use of resources by an individual, in this case melanin which is a multipurpose and apparently costly pigment [63].


Colour polymorphism is a striking feature of several species, which involves variation in the types of morphs and different environmental triggers and genetic determinants which combine to help define the colour morph of an individual [14, 38]. Here we report the absence of colour morph switches in the alpine grasshopper G. sibiricus. The lack of switches could be driven by the heterogeneity of the habitat of G. sibiricus. Additionally we found that nymphae of G. sibiricus are capable of modifying the amount of colouration darkness in response to radiant heat. This capability could help nymphae adjust colouration within nymphal stages according to the unstable climatic conditions characteristic of their habitat, and in turn improve allocation of pigment resources to other physiological processes. The complex net of interactions between fine-scaled variation in polymorphisms, unpredictable and highly variable environments, and melanin pathways which may be costly and also shared by different developmental processes are not only a very promising avenue of study in behavioural ecology, but they will also change the way in which we assess colour polymorphism in natural populations.

Last-instar individuals of G. sibiricus were collected in the field (near Sierre, Valais, Switzerland) in July 2013 and brought to the laboratory where they moulted into imagoes. We housed imagoes in separate cages (dimensions 22 × 16 × 16 cm 3 ) containing one male and one female and provided a cup of sand-vermiculite mixture as substrate for egg laying. Eggs were collected once per week, kept for approximately 6 weeks at room temperature and subsequently stored at 4 °C for diapause. After seven months in the refrigerator, eggs were taken out of their diapause and kept at room temperature until hatching. In total, 116 hatchlings hatched from 37 egg pods. Full-siblings from the same egg pod were housed together in the same cage (white cages lined with black mesh, dimensions as above) throughout the first nymphal stage, but were separated after the first nymphal stage (i.e., after about one week) and thereafter housed individually. We refer to the different nymphal stages as N2 (second nymphal stage), N3, N4 and IM (imago stage).

Experimental setup

The experiments were conducted under artificial full-spectral light conditions (Biolux L 58 W/965, OSRAM, Munich, Germany) with a 14:10 H light dark cycle (7:00 till 21:00). Average humidity was kept at 70 % by humidifiers. In order to maintain a high moisture content in the food, we provided fresh grass in small plastic vials (height x diameter: 5.8 cm × 2.1 cm) filled with water, and replaced the grass every other day in order to avoid withered or yellowing grass. Experiments were conducted in two rounds (we refer to the two rounds as R1 and R2) that differed in details of the radiant heat treatment, because of high nymphal mortality under one of the conditions in R1. Experimental setups consisted of blocks of four cages (11 and 10 replicate blocks in the R1 and R2, respectively) flanked on both sides by isolating dividers (22 cm × 55 cm × 1 cm) (Fig. 3 ). The dividers were installed to isolate alternating blocks exposed to different experimental conditions. Individuals were transferred to experimental setups on the day of moulting into the N2 and assignment to treatments was done at random. All individuals remained under the same experimental conditions until one week after moulting into IM (unless they died before completing the experimental period). Only three individuals reached the last nymphal stage under low radiant heat conditions in R1, two of which died shortly after, and therefore the experiment was stopped at this point due to lack of one treatment group. We did not score the sex of individuals during early nymphal stages because this is easily visible during the N4 stage. In the case of the R1, individuals died rapidly while the experiment was running, and sex scoring was not assessed in time. For this reason data on colour morph frequencies for both sexes is not presented in the results.

Experimental setup. Two blocks are shown, one exposed to the high radiant heat (orange dot) and one to the low radiant heat (yellow dot) treatment. Blocks are separated by dividers (Di). Each cage has its floor and two of its sides lined in green or brown paper, and each block has two cages lined with each colour. Time of exposure to radiant heat varied between rounds (see details in Temperature treatment section). The doors (do) on the sides allow easy access to cages

Background colour treatment

Temperature treatment

The second component of our experimental setup consisted of a temperature treatment and was applied to entire blocks. Grasshoppers regulate their body temperature behaviourally, elevating it substantially above ambient temperature by sun-bathing [24]. We aimed to simulate high (sunny) and low (overcast) radiant heat conditions, both of which occur in the natural habitats of G. sibiricus. Therefore, we placed 150 W infra-red heat bulbs (IOT/90, Elstein, Northeim, Germany) at the vortex of each block. In the R1, blocks were exposed to either 5 h (high treatment) or 1 h (low treatment) of radiant heat, but due to high mortality, this was increased to 4 + 6 h (high treatment) or 2 h (low treatment) per day in the R2 (Fig. 3 ). The hours of exposure in the R1 for the high treatment were from 9:00 until 12:00, and again from 13:00 until 15:00, while the exposure time for the low treatment was from 12:00 until 13:00. The hours of exposure in the R2 for the high treatment were from 8:00 until 12:00, and again from 14:00 until 20:00, while the exposure time for the low treatment was from 12:00 until 14:00. The temperature at exposed locations was around 44 °C when heat lamps were active, in contrast to the 25 °C when heat lamps were switched off. In the high radiant heat treatment, individuals were able to adjust the amount of radiant heat that they received by positing themselves at the mesh ceiling or by moving away from it or also by hiding behind the beams of the cage.

Scoring of body colour

Colour morphs are > 90 % of black. Our analysis is based on a total of 433 scorings from the R1 and 306 scorings from the R2.

Statistical analysis

Since our sample size is necessarily finite, it is at least possible that despite an absence in our sample, colour morph switches occur with a low probability. We therefore estimated the lowest probability of colour morph switches that would still be consistent with our data. We used the following approach to assess the highest colour change probability that is consistent with our data. We define c as the probability of colour change in unmatched backgrounds, (1-c) as the probability of no change and k as the number of individuals exposed to unmatched backgrounds. Based on binomial sampling, we expect to observe no colour morph switches among k individuals with a probability of (1-c) k . We defined a probability threshold of α = 0.05 and searched for the value of c at which the expectation of no-colour morph switches is equal to α. This gives the highest possible switching probability that is still considered reasonably consistent with our data.

To test for the effect of the temperature treatment in the body colour we used a random-slope mixed effects model. We chose this model because the possible response to the treatment could vary physiologically from individual to individual, making it necessary to account for this variability in our analysis [64]. The p-values for the fixed effects in the mixed models were approximated by a t distribution with degrees of freedom equal to the number of individuals minus the number of fixed effect parameters estimated in the model. This approach is conservative in that it considers only individuals as independent datapoints. However, such random-slope models depend on assumptions about the linearity of the response-covariance relationship and about the distributions of random-deviations from the population mean and population slope. Therefore, we applied additional, simpler tests for the same question in order to verify the robustness of the results (see below for details).

In addition to the analysis of the data using random slope models, we cons >

Availability of supporting data

The data sets supporting the results of this article are available in the Dryad repository: doi:10.5061/dryad.j95t2 [67].


Thanks to Amy Backhouse for the valuable feedback and additional help provided. Thanks to Bahar Patlar and Petra Dieker for valuable comments on the manuscript. Thanks to Axel Hochkirch for the data provided for the polymorphism table. Thanks to Veronica Dahm for her valuable comments and support. The project was funded by an Emmy Noether grant from the German Research Foundation (DFG; SCHI 1188/1-1).


Round 2
slope SE t p slope SE t p
N2 (n = 42) N2 (n = 35)
Intercept 1.79 0.25 7.16
N1 First nymphal stage
N2 Second nymphal stage
N3 Third nymphal stage
N4 Fourth nymphal stage
IM Imaginal stage
R1 First round
R2 Second round

Competing interests

The authors declare that they have no competing interests.

Authors’ contributions

JPV and HS perceived and designed the experiment. JPV performed the experiments and drafted the manuscript. JPV and HS analysed the data and the manuscript. All authors read and approved the final manuscript.


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