Where does grasshopper come from rain

Where does grasshopper come from rain

Grasshopper Control

Grasshoppers are worse in hot dry years. When we have cool, wet springs, they can succumb to disease that keeps their numbers in check. Some years they don’t show up until the end of May and June. In 2006, they arrived in March, and even as late as August, young ones still appeared.

Control is difficult unless since it’s hard to hit them with a contact spray. The best thing to do, as soon as you see the nymphs, is to apply a bait. If you see leaves with tiny holes, or find more than 8 grasshoppers to a square yard, it’s time to treat.

Effective baits include Nolo Bait or Semaspore. Both contain a protozoa called Nosema locustae which is impregnated in bran flakes sweetened with sugar.

Apply by hand or with a rotary spreader, early in the morning, when grasshoppers are feeding. They don’t just drop dead – they’ll slow down and cling to plants. You won’t see any dead ones, since the healthy grasshoppers cannibalize the sick and weakened ones. Those grasshoppers will then ingest the protozoa; females will pass it along through their eggs. You’ll see a better reduction in population if you are diligent and enlist your neighbors in the project as well.

Be sure to use fresh bait that has kept chilled at the store, and bring it home in an ice chest. Grasshoppers will not be attracted to old bait. Look at the formulation date on the side of the container. If not chilled, it will last for 4 weeks. If refrigerated, a container will last 4 months.

You can also use a spray of Kaolin clay, available at craft stores that sell pottery supplies. Mix it up in a pump-up sprayer – 2 ½ cups of clay to a gallon water plus a teaspoon of soap as a surfactant. It’s important that you constantly agitate the sprayer while you work! Spray leaves lightly. It leaves a fine film of powder on the leaves, which gums up a grasshopper’s mouthparts, so it won’t eat there. The film also acts as sort of a shade cloth to protect leaves in summer’s heat! Obviously this won’t work if you use a sprinkler system, and must be reapplied after a rain.

Another prevention against grasshoppers is tightly staked row cover for plants that do not require pollination.


Where does grasshopper come from rain

Imperial College of Science and Technology, London

Imperial College of Science and Technology, London

Log in with Open Athens, Shibboleth, or your institutional credentials.

If you have previously obtained access with your personal account, Please log in.

If you previously purchased this article, Log in to Readcube.

You previously purchased this article on READCUBE_PURCHASE_DATE. Click on an option below to access.

View Enhanced PDF Access article on Wiley Online Library (HTML view) Download PDF for offline viewing

Logged in as READCUBE_USER. Log out of ReadCube.

1. The eggs of Acrididae are well adapted to survive adverse weather and mortalities in field populations from this cause are low in most years. The eggs of tropical species are generally only laid where conditions are suitable for growth, while an egg diapause is present in many temperate species, during which time eggs are resistant to extremes of both temperature and moisture. Only prolonged flooding or drought can cause high mortalities and then only at certain stages in the egg’s development. All eggs must absorb water to complete their development.

2. Nymphal and adult populations are far more dependent on the right weather conditions. Prolonged wet, cloudy weather greatly increases the mortality of both. Newly hatched nymphs are particularly susceptible. The way in which wet weather affects survival is little understood, but since the rate of feeding of Acrididae is so dependent on warm sunny conditions, it is possible that inclement weather may lead to starvation.

3. Fecundity is also greatly reduced by cool wet weather and the level indicated by laboratory studies is rarely reached in the field.

4. While both survival and fecundity are enhanced by hot, dry conditions, both are dependent on the presence of green food, which can be destroyed by excessive drought.

5. These opposing requirements of Acrididae make their numbers extremely sensitive to variations in weather and cause large fluctuations in their populations.

6. The eggs of most acridids are laid in bare ground, while vegetation is required by nymphs and adults for food and shelter. These requirements are again opposing. High‐density populations are often found in natural and man‐made ecotones, in which vegetation and bare ground occur in a mosaic pattern, thus providing both in their maximum availability. Ecotones are often unstable and the extent of the two vegetation components (bare ground and plant cover) is greatly influenced by weather. Variations in weather may then further affect acridid populations through effects on the carrying capacity of the habitat.

7. In locusts, gregarious behaviour and swarming are brought about by increases in population density, and variations in the relative extent of the components of the vegetation mosaic may cause crowding of one or other of the stages in the life cycle.

8. The influence of weather on acridid populations is so marked that correlations have been found for a large number of species between population size and particular weather conditions. The most important factor determining numbers varies between different species occupying different habitats and geographical regions.

9. Acridid populations are attacked by a large array of natural enemies–diseases, parasites and predators. Little quantitative work has been done on their effects, but biological considerations show that it is extremely unlikely that any are sufficiently density‐dependent to act in the way suggested by Nicholson (1933, 1954, 1958). It is likely that they do little more than damp the peaks in population fluctuations.

10. Many locusts and swarming grasshoppers emigrate from their habitats when their populations are large. This density‐dependent emigration possibly occurs in all Acrididae, though it is less well documented for non‐swarming species.

11. Locusts change both biologically and morphologically when crowded. These changes appear to be an adaptation to life away from their permanent habitat. Basically they reflect a change in the individua’s metabolism to favour greater mobility. These ‘phase’ changes also affect fecundity, and crowding results in a marked reduction in the number of eggs laid.

12. Population size may be controlled by emigration and phase changes. When numbers become high, emigration of part of the population brings them down again. In locusts, phase changes could theoretically reverse the trend at both the peaks and troughs in population size.

13. The dynamics of acridid populations are discussed in the light of the main theories in population control, and it is concluded that Milne’s theory (1957a) is closest to the known facts.


Getting those grasshoppers

Most areas won’t have grasshopper problems in 2014. But there are some places where farmers should hone their >

For the most part, Saskatchewan farmers are unlikely to be fighting waves of grasshoppers this year, according to Saskatchewan Agriculture’s 2014 hopper forecast.

But Saskatchewan’s 2014 forecast does show one bull’s eye of severe risk.

“And in the middle of that bull’s eye is Meadow Lake,” Peter Walsh told farmers at Cavalier Agrow’s farm forum in North Battleford this April. Walsh teaches courses in insect, weed and disease management, along with crop agronomy, at Lakeland College in Vermilion.

A count of at least 12 adult hoppers per square metre in the fall adds up to a severe risk forecast the following year.

Meadow Lake farmers aren’t the only Prairie producers likely to suffer hopper plagues this year. Alberta Agriculture and Rural Development’s forecast shows very severe risk (more than 24 hoppers found per square metre) near Grande Prairie, along with areas in the northeast and west-central regions.

Alberta also has severe risk areas elsewhere in the Peace, in the northeast, between Calgary and Lethbridge, and in the east-central part of the province.

Most surveyed areas in Manitoba last year rated very low risk. But hoppers numbered 18 per square metre at a site near Crystal City and 22 per square metre near Wawanesa.

Farmers in zones not rated as high risk aren’t necessarily going to get off scot-free this year, though.

Forecasts partly depend on weather and natural enemies, Dr. John Gavloski, entomologist with Manitoba Agriculture, Food and Rural Development, said in an interview. And local populations can vary.

“We try to do our best to make sure that the counts are representative, but that’s not always the case,” said Gavloski.

Identifying pests

Not all grasshoppers are crop killers and it’s well worth knowing which are friend and which are foe.

Manitobans walking the ditches might see bright green insects resembling grasshoppers. They are katydids, Gavloski said. “They’ll never move in and damage the crop.”

Both Gavloski and Walsh cited the Russian Thistle Grasshopper as a species that only eats weeds.

But farmers don’t have to be able to name the specific specie to know if a hopper will gorge itself on grain. Any grasshopper that makes a clacking noise while flying, has colourful wings (yellow or rose-coloured), or is flying in April or May is not a pest, Walsh told farmers.

“All of our pest species over-winter as eggs. That’s why in June they’re just hatching out,” said Gavloski. Pest species usually aren’t adults until July.

But some non-pest species over-winter, said Gavloski. “So they’re nearly mature early in the season.”

Females that are common pests will lay eggs in the fall “until the frost shuts her down,” said Walsh.

Nymphs emerge and begin feeding immediately, Walsh added. “The grasshopper does not have a resting stage. There is no stage in the hopper like a cocoon.”

Scouting and thresholds

Alberta Agriculture and Rural Development’s website recommends checking field edges, fence lines and ditches for grasshoppers. Egg beds generally line field edges so instars will be found there first.

To count grasshoppers, Manitoba Agriculture’s site suggests starting in one field corner, walking diagonally past the centre, and then walking straight out to one side of the field. While walking, note how many nymphs jump from a square foot area.

The ministry suggests taking at least 20 of these counts per survey. Dividing the total by two will give an approximate number of hoppers per square metre.

The economic threshold for grasshoppers is eight to 12 hoppers per square metre, according to Manitoba Ag’s site. Gavloski said this threshold is nominal, meaning there isn’t quantitative data correlating insect damage to yield loss.

“It’s basically a best guess of what people think is likely economical,” he said.

Farmers with lower-value crops should err on the high end of the economic threshold, Gavloski said. But for high-value crops, action is needed when grasshoppers breach the lower end of the spectrum, he added.

And crops such as lentils and flax likely require action before the grasshoppers hit the low end of the threshold. In fact, lentils at the flowering and podding stage have a recommended threshold of two hoppers per square metre.

Soybeans and canola aren’t preferred snacks for most hoppers, but there are species that will eat them once their preferred food sources dry down, Gavloski said.

“Cereal crops are favoured by some of our pest species, so they’re certainly more vulnerable,” said Gavloski.

Control timing

Gavloski suggested controlling hoppers at roughly the third or fourth instar stage, before they start damaging crops.

“Adult grasshoppers are much harder to control than juveniles,” said Gavloski. “And the juveniles are often concentrated along field edges and borders.”

Walsh also recommended farmers delay spraying until nymphs hit the third instar. At that stage they’ll have small wing pads.

“What you’re waiting for is the complete hatch. If you jump the gun the first time you see those little nymphs out there, you may have to spray a second time,” said Walsh.

But farmers will need to weigh this advice against how much damage is already being done to the crop and the crop type, said Walsh. Grasses, such as wheat, will outgrow some of the early damage, Walsh added.

“And if I’m still waiting for the rest of the hatch, I probably don’t have peas or canola into bud or any kind of stage like that,” Walsh said.

It ultimately comes down to individual farmers’ risk tolerance, Walsh said. “Where’s your line? Everybody’s got their line. And just some cautions that if you can hold off a bit, you may only have to spray once.”

Common wisdom says that a wet spring is hard on newly-hatched hoppers, but spring rains don’t always drown grasshoppers.

Instars “breathe through their abdomens. And they can’t lift their abdomens out of the muck and the mud and the water,” Walsh explained.

But wet weather doesn’t affect the eggs, so whether or not spring rains kill instars comes down to timing.

“Heavy rains in June could potentially kill lots of grasshoppers. Heavy rains in April will do next to nothing,” said Gavloski.

And a cold spring will delay the hatch, and so if the wet weather passes, the eggs will still be viable, said Walsh.


Cold Blooded – But Not Cold Hearted

Insects are ectothermic and therefore their body temperature and activity is heavily influenced by their environment. So generally, the colder it is, the slower insects move and the warmer it is, the faster and more active insects are. However, if your life is completely dictated by the environment it makes it hard to exist for very long, especially if you’re a little insect. You don’t want to just be frozen in place every time an unexpected chill blusters by.

So insects have some tricks up their sleeves to help them deal with the inclement weather.

If bees get too cold, they’ll huddle together and shiver – just like us – to keep themselves warm. Queen bumblebees disengage their wings and use the flight muscles to generate heat to help them forage for food in the early, chilly spring.

And some arthropods simply have antifreeze proteins to help them when it gets extremely cold – like this snowflea that cheerily hops along snow banks in sub zero temperatures.

But if it’s raining, it’s probably not below freezing. So what do insects do in the rain?
That varies considerably based on the insect and their behavior.

Staying In

Many insects can sense atmospheric pressure differences. Honey Bees for instance, just stay home if they sense a storm coming. Other bees, like Mason Bees may stay out and forage in light rain but will take shelter when it starts raining too heavily or the wind gets too intense.

Since the water can weigh them down, it’s harder for insects to fly when it’s cold and the rain can damage their wings, many insects just seek shelter. After a particular rainy afternoon in Ecuador, I looked under a small leafy plant and found several butterflies, hunkered down, just waiting for the storm to pass.

Some resting butterflies I found after a big storm in the Ecuadorian rain forest.
PC: Nancy Miorelli

Also, after a particularly wet morning, I found some Red Flat Bark Beetles (Cucujus clavipes) hiding away under some bark, clearly disgruntled that I disturbed their slumber.

Out Past Curfew

Raindrop size in relation to a mosquito.
PC: Dickerson et al. 2012

Small insects that thrive in warm and humid areas fly in the rain anyway. Mosquitoes, in particular, were studied to determine how they can resist the rain. For the most part, they’re just small and many raindrops don’t hit them. If they are hit by the raindrops, the mosquitoes just kind of become assimilated into it and fall with the raindrop. They then escape the falling raindrop with the help of their water resistant hairs. Every mosquito in this experiment managed to escape the water droplets.While we don’t really have the answers, it’s assumed that other small, flying, and agile insects do the same.

Being able to stay active when other insects are not reduces competition. There are lots of other insects that would like to blood feed on humans and other animals, so being able to stay out in conditions when others can’t, allows you to feast. You snooze you loose … literally.

Just Waiting for the Party

Some insects specifically wait for the rain to complete their reproductive cycles. The winged reproductives of a leafcutter ant in Texas fli]y just after rain on moonless nights in the early spring.

Rain beetles are another group that specifically wait for the rain. They live underground and during the rain, the females will come to the surface of their burrows and release pheremones. The males, also aware of the rain, escape their soil forts and fly to the find the females in the very early morning. By following the pheromone trail, the males find a female’s burrow and mate with her, where she then lays eggs, and they all live underground again until the next rainy winter.

Depends on the insect. Some wait for the storm to pass, some don’t go out at all, some just dodge the raindrops or escape them, and others specifically wait for the rain to get their funk on. Also, as a short aside, there are lots of insects just live in water, in which case, the rain doesn’t really affect them.

These Shield Bug nymphs don’t wanna go out there with the weather like that!
PC: Peter Nijenhuis (CC by NC ND 2.0)


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.


No Comments

Leave a Reply

Your e-mail will not be published. All fields are required.