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Insect making continuous sound in summers

Insect making continuous sound in summers


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In deciduous to tropical-deciduous forests and residential places with such vegetation I have found this insect to make continuous sounds all through the day (even in nights sometimes) during the summers. Average summer temperature > 30 C around noon and ~ 28-29 around nights with high humidity.

I want to know what is the reason they make this sound ?

Recently, I found several of them dead near light poles etc… , and some sit very close to windows of air-conditioned rooms. I think they want some cool environment.


These are Cicadas, of the superfamily Cicadoidea of the order Hemiptera.

These sounds are made by the males to attract mates. There are a lot of species, each making there own specific sound to attract the right mate. However, to the human ear, these sounds don't sound different. Cicadas are also able to produce other sounds, in distress or during courtship and mating, but the continuous sound during night and day is to attract mates.


These are definitely cicadas and mating is the primary reason for their long calls. As for the reason you've found many dead or dying cicadas, their lifecycle is primarily spent underground where they feed on tree roots. They will emerge and transform into their adult form which enables them to make their calls and mate, and many species die shortly after mating, having fulfilled their evolutionary function of reproduction. Some species spend up to 10+ years underground before emerging to mate for only a short week or so.


Top 6: Noisy Insects

Do you hear that? That music being broadcast outside your window of chirps, tweets and thumps is a full-on symphony of insects belting out their own unique tunes. Some are peaceful and serene others are loud and boisterous. No matter the melody, we’re counting down the loudest insects in this month’s Top 6: Noisy Insects.

6. Cicadas
The sound of the cicada is highly recognizable when summer rolls around. Sounding like a high-pitched rattle, this mating call sung by the males is the result of vibrating a part of their body called the tymbal. By doing this, cicadas are able to make loud noises, and groups of them can be heard from more than a mile away. Wow!

5. Flies
You know the sound. It’s common and all too annoying: you’re enjoying your favorite television show, and a fly buzzes past your head like a fighter pilot. The sound, like a miniature airplane propeller, is from the beating of their wings. These fast flappers can beat their wings about 200 times a second, allowing them to fly up to 4.5 miles per hour.

4. Crickets
Another common sound of summer is that of the cricket. They’re even named after the high-pitched chirping sounds males make to attract females. When crickets rub their front wings together, this long chirping sound is amplified by the wing’s surface. What’s even more interesting? Cricket chirps can be used to calculate temperature by the number of sounds they make in a fifteen-second period, making them the weather forecaster of the insect world.

3. Bees
Everyone knows bees buzz. You can hear when they’re zooming overhead looking for a flower to munch on, or when they congregate into large groups in hives. Just like flies, they rapidly beat their wings — creating vibrations in the wind. This is the buzz humans hear when they fly by. Some bees — namely bumblebees — can also vibrate their bodies. They do this when visiting flowers to shake pollen off. Pollination occurs when the pollen is disbursed by vibration at the next flower the bees visit.

2. Katydids
Not only are they masters of disguise they also know how to sing quite the tune. The song of the katydid is one that can be heard outside of any household. Almost like an increasing static resonance, katydids are able to produce this sound on a broadband level by rubbing their forewings together. This rubbing vibration has been compared to sounding like “katy did, katy didn’t,” hence their name.

1. Longhorned Beetles
Claiming the top spot of our noisy critter rundown is the longhorned beetle. With a hefty appetite for hardwood trees, this beetle’s noisy demeanor results from scraping the ridges on their head against their bodies. This produces a truly creepy, static-like squeaking sound that can only be described as rubbing two pieces of styrofoam together, which is like nails on a chalkboard for some people.


The Soothing Sound of Crickets

The sound crickets make is referred to as chirping, but they aren’t making the noise with their mouths. They’re also not making hose sounds with their back legs, as was once commonly thought. Instead, much like The Cricket In Times Square, they make noise by rubbing their wings together.

If you’ve ever run your fingernail along the tines of a comb and listened to the sound it makes, you have a bit of an idea of how a cricket makes its chirping sounds. Crickets have a thing on the tops of their wings called a scrapper that they use to rub along the bottom of the opposite wing. The bottom part of the wing is called a file.

There are different types of crickets, and depending on the species, both males and females chirp. In some species, only males chirp, and some males are silent. Like fireflies have specific flash patterns to send messages, crickets have chirp patterns that are meant to attract mates that are far away, court those that are closer to them, and they even have a song of triumph once they score a mate.

What may be even more impressive than how crickets make their sounds is that you might be able to determine the temperature by the bug’s chirps without a thermometer. Science and math work together for this task, which requires you to count the chirps of a cricket over three fourteen-second intervals, then averaging the amount out. By adding that amount to forty, you get an estimate of the temperature in Fahrenheit—Scientific American has the detailed instructions as well as info on how this works.


Materials and methods

Data from the literature

The literature concerning insect gas exchange patterns was reviewed as far back as 1950, and all studies in the Anglophone literature reporting gas exchange patterns were included. Where authors provided figures of the gas exchange patterns of the species they studied, these were used for assessments of the type of gas exchange pattern (either DGC, cyclic or continuous). DGCs were identified on the basis of the presence of C and F periods(Lighton, 1996 Chown et al., in press) in the figures presented by the authors. For the other patterns our assessments were based on the protocol described in the next section. In those instances where figures were not available, the authors' view on the type of pattern was accepted as correct, although confidence in the pattern assessment was marked lower (confidence in the data was ranked either as high, medium or low, which reflects our access to original data, rather than the abilities of the original authors). These assessments were made independently by E.M. and C.J.K., and in instances of disagreement a consensus was reached following discussion or analysis. The data were then tabulated. In two instances original data from published (Shelton and Appel, 2000) and unpublished works (B. A. Klein, K. M. F. Larsen and A. G. Gibbs) were obtained to verify these assessments. Each species was also scored for whether it is winged or wingless, from a mesic or xeric habitat, or expected to have a subterranean lifestyle, based on comments provided by the authors in the original works, and/or information on the species or higher taxon available elsewhere in the literature.

Experimental investigations

The additional species collected for investigation were chosen based on Order-level deficiencies in the literature on gas exchange patterns. Adult individuals of 19 species representing the Archaeognatha (1 sp.), Zygentoma (3 spp.), Ephemeroptera (1 sp.), Odonata (2 spp.), Blattodea (1 sp.), Mantodea (1 sp.), Mantophasmatodea (1 sp.), Phasmatodea (1 sp.), Orthoptera (1 sp.),Dermaptera (1 sp.), Hemiptera (2 spp.), Neuroptera (1 sp.), Diptera (1 sp.),Trichoptera (1 sp.) and Lepidoptera (1 sp.) were collected from several localities in South Africa (Table 1) and returned to the laboratory within 1 week of collection. Most experiments started within 12 h of the arrival of the insects at the laboratory because little is known about how long they survive in captivity. Insects were held in an incubator at 22±1°C (12 h:12 h L:D photoperiod), with access to water but not to food (with the exception of the hemipterans, mantophasmatodeans, cockroaches and the stick insects, where food was provided, but where a period of starvation preceded respirometry), before their gas exchange patterns were examined. Assessments were made in dry air for technical reasons and because under these conditions discontinuous gas exchange would seem most likely as a means to conserve water(Duncan et al., 2002). Each individual was weighed using an analytical balance (0.1 mg resolution Mettler Toledo AX504, Columbus, OH, USA), and placed into a cuvette kept at 20±0.2°C, using either a water bath (Grant LTD20, Cambridge, UK) or a temperature-controlled cabinet (Labcon, Johannesburg, South Africa). This slightly lower temperature was selected because it improved quiescence and might have also induced discontinuous gas exchange. Previous work(Chown, 2001 Marais and Chown, 2003)indicated that gas exchange patterns, whilst repeatable, can be variable within individuals and species. In consequence, conditions favourable to the induction of discontinuous gas exchange were used, and particularly temperatures that are typically lower than mean summer microclimate temperatures in the region (which range from 24°C at sea level, to 22°C at the highest inland site of collection, with absolute maxima ranging from 50°C at the sea level site to 53°C at the high altitude site see also Botes et al., in press).

Species examined for gas exchange patterns in this study

. Locality . Response time, lag time (s) .
Archaeognatha
Meinertellidae, sp. Helderberg Nature Reserve, Somerset West, South Africa (34°02.579′S,18°52.472′E) 6, 120
Zygentoma
Lepismatidae
Lepismatidae sp. 1 Sutherland, South Africa (32°34.105′S, 20°57.747′E) 6, 120
Lepismatidae sp. 2 Cederberg, South Africa (31°51.611′S, 18°55.122′E) 6, 120
Ctenolepisma longicaudata (Echerich, 1905) Stellenbosch, South Africa (33°55.923′S, 18°51.812′E) 6, 120
Ephemeroptera
Heptageniidae sp. Stellenbosch, South Africa (33°55.923′S, 18°51.812′E) 9, 210
Odonata
Coenagrionidae
Ischnura senegalensis(Rambur, 1842) Jonkershoek, Stellenbosch, South Africa (33°57.814′S,18°55.514′E) 7, 130
Libellulidae
Trithemis arteriosa(Burmeister, 1839) Jonkershoek, Stellenbosch, South Africa (33°57.814′S,18°55.514′E) 18, 270
Blattodea
Blaberidae, sp. Cederberg, South Africa (31°51.611′S, 18°55.122′E) 4, 90
Mantodea
Mantidae
Sphodromantis gastrica Stål Stellenbosch, South Africa (33°55.923′S, 18°51.812′E) 7, 130
Mantophasmatodea
Austrophasmatidae
Karoophasma biedouwensis (Klass et al., 2003) Cederberg, South Africa (32°05′S, 19°15′E) 9, 210
Phasmatodea
Phasmatidae
Extatosoma tiaratum(Macleay, 1826) Butterfly World, Klapmuts, South Africa, but originally from Australia 7, 130
Orthoptera
Pneumoridae
Bullacris intermedia(Péringuey, 1916) Zuurberg, South Africa (33°48′S, 25°14′E) 7, 130
Dermpatera
Labiduridae
Euborellia annulipes(Lucas, 1847) Stellenbosch, South Africa (33°55.923′S, 18°51.812′E) 6, 120
Hemiptera
Coreidae sp. Nigel, South Africa (26°25.422′S, 28°28.349′E) 6, 120
Lygaeidae sp. Somerset West, South Africa (34°03.806′S, 18°49.473′E) 6, 120
Neuroptera
Chrysopidae
Chrysoperla sp. Somerset West, South Africa (34°03.806′S, 18°49.473′E) 9, 210
Diptera
Glossinidae
Glossina morsitansWestwood FAO/IAEA, Vienna, Austria (Laboratory colony) 6, 120
Trichoptera
Leptoceridae
Leptocerina sp. Olifants River, Citrusdal (32°35′S, 18°40′E) 9, 210
Lepidoptera
Plutellidae
Plutella xylostella(Linnaeus, 1758) Somerset West, South Africa (34°03.806′S, 18°49.473′E) 9, 210
. Locality . Response time, lag time (s) .
Archaeognatha
Meinertellidae, sp. Helderberg Nature Reserve, Somerset West, South Africa (34°02.579′S,18°52.472′E) 6, 120
Zygentoma
Lepismatidae
Lepismatidae sp. 1 Sutherland, South Africa (32°34.105′S, 20°57.747′E) 6, 120
Lepismatidae sp. 2 Cederberg, South Africa (31°51.611′S, 18°55.122′E) 6, 120
Ctenolepisma longicaudata (Echerich, 1905) Stellenbosch, South Africa (33°55.923′S, 18°51.812′E) 6, 120
Ephemeroptera
Heptageniidae sp. Stellenbosch, South Africa (33°55.923′S, 18°51.812′E) 9, 210
Odonata
Coenagrionidae
Ischnura senegalensis(Rambur, 1842) Jonkershoek, Stellenbosch, South Africa (33°57.814′S,18°55.514′E) 7, 130
Libellulidae
Trithemis arteriosa(Burmeister, 1839) Jonkershoek, Stellenbosch, South Africa (33°57.814′S,18°55.514′E) 18, 270
Blattodea
Blaberidae, sp. Cederberg, South Africa (31°51.611′S, 18°55.122′E) 4, 90
Mantodea
Mantidae
Sphodromantis gastrica Stål Stellenbosch, South Africa (33°55.923′S, 18°51.812′E) 7, 130
Mantophasmatodea
Austrophasmatidae
Karoophasma biedouwensis (Klass et al., 2003) Cederberg, South Africa (32°05′S, 19°15′E) 9, 210
Phasmatodea
Phasmatidae
Extatosoma tiaratum(Macleay, 1826) Butterfly World, Klapmuts, South Africa, but originally from Australia 7, 130
Orthoptera
Pneumoridae
Bullacris intermedia(Péringuey, 1916) Zuurberg, South Africa (33°48′S, 25°14′E) 7, 130
Dermpatera
Labiduridae
Euborellia annulipes(Lucas, 1847) Stellenbosch, South Africa (33°55.923′S, 18°51.812′E) 6, 120
Hemiptera
Coreidae sp. Nigel, South Africa (26°25.422′S, 28°28.349′E) 6, 120
Lygaeidae sp. Somerset West, South Africa (34°03.806′S, 18°49.473′E) 6, 120
Neuroptera
Chrysopidae
Chrysoperla sp. Somerset West, South Africa (34°03.806′S, 18°49.473′E) 9, 210
Diptera
Glossinidae
Glossina morsitansWestwood FAO/IAEA, Vienna, Austria (Laboratory colony) 6, 120
Trichoptera
Leptoceridae
Leptocerina sp. Olifants River, Citrusdal (32°35′S, 18°40′E) 9, 210
Lepidoptera
Plutellidae
Plutella xylostella(Linnaeus, 1758) Somerset West, South Africa (34°03.806′S, 18°49.473′E) 9, 210

Localities are provided, and species names where these are available. However, the taxonomic impediment in South Africa means that the latter has not always been possible. Response and lag times refer to delay in first detection of CO2, and time to zero baseline, respectively, of each of the designs used to examine gas exchange patterns. With one exception these times are well within those calculated from designs typically described in the literature for gas exchange analyses in insects.

Air, scrubbed of CO2 (using soda lime) and water (using silica gel and then Drierite®, Xenia, OH, USA) was passed through the cuvette(see Table 1 for response times, regulated using a Sidetrak Mass Flow Controller, Monterey, USA) and into a calibrated infrared gas analyzer (Li-Cor Li7000 or Li-Cor Li6262Lincoln, NE, USA) to measure CO2 production. Flow rates and cuvette sizes varied according to the species and in a manner such that washout was unlikely to be significant (see Results, and Lighton, 1991b). A Sable Systems (Las Vegas, NV, USA) AD-1 activity detector was used to detect any movement of the individual in the cuvette during the experiment, and the output of the detector was fed into the auxiliary channel of the Li7000 or Li6262. The AD-1 registers activity as a value between –5 and +5 V,where little deviation from the mean indicates that the animal is inactive,and a large deviation indicates high levels of activity (for detail see www.sablesys.com/ad1.html). Each experimental assessment lasted for approximately 2 h, which is typically sufficient to detect variation in gas exchange traces(Chown, 2001) without dehydrating animals to such an extent that the gas exchange pattern might switch to continuous, owing to dehydration, as has been found in some species(Quinlan and Hadley, 1993 Chappell and Rogowitz, 2000). The data file generated by the Li7000 software was exported, viaMicrosoft Excel, to DATACAN V (Sable Systems,), whilst the data stream from the Li6262 was captured directly using Sable Systems hardware and software. DATACAN V was used for initial analysis of the respirometry data (corrected to standard temperature and pressure) for periods of inactivity only.

Traces of rates of CO2 production(CO2) were categorized as continuous, cyclic or discontinuous gas exchange by inspection. The DGCs were readily identified based on the presence of C-periods and F-periods. However,identification of gas exchange patterns in the absence of the C- and F-periods is less straightforward. Several statistical approaches were explored for distinguishing continuous from cycling patterns objectively. These included spectral analysis and the modification thereof that has been used to identify population cycles (Cohen et al.,1998). Unfortunately, these methods typically did not allow continuous and cyclic gas exchange to be distinguished, most notably because even continuous gas exchange has some periodicity. The variance approach adopted by Williams et al.(1997) is also unsuitable because it does not take temporal autocorrelation into account. In consequence, any comparison of variances between species would be confounded. Nonetheless, it is essential that some objective criterion has to be developed to allow traces to be classified or distinguished in a repeatable manner.


Cicadas vs katydids: Who’s singing this summer?

As a Southerner, I take my bugs seriously and I’m not so happy about the critter situation this summer.

Evenings on the porch have been sort of quiet, as cicadas seem to be sitting this one out while the katydids have taken over. It shouldn’t be surprising, given the unique life cycles of some North American cicadas. They can have 13- or 17-year lifespans, most of which is spent underground. When they emerge, it’s like a hallelujah chorus has taken over the trees, and the very air around you shakes. I’ve heard some so far this summer but not that all-encompassing thrum yet.

“Last year, you had an inordinate number of cicadas,” said Bennett Jordan, an entomologist and staff scientist at the National Pest Management Association. “The 17-years came out last year. They’ll come out again in 2030.”

If you miss them as much as I do this year, you may need to plan a road trip to Iowa or Illinois, where 17-year cicadas have emerged or Louisiana, Mississippi, Kentucky or Ohio, where the 13-year crowd is making an appearance, according to Gene Kritsky, editor-in-chief of American Entomologist magazine and author of “Periodical Cicadas: The Plague and the Puzzle.”

In addition to the 17- and 13-year bumper crops, we do get annual cicadas that emerge each year.

“Give it a little more time. We had a lot more rain this year than we’ve had in the past,” said Kritsky, who also serves as professor and chair at Mount St. Joseph University’s Department of Biology. “It’s been a little later of a spring, slightly cooler.”

While we wait on the cicadas, the katydids, which operate on one-year life cycles, are making plenty of racket. How to tell the difference? Easy.

Cicadas sound like a tiny tambourine rattling louder and faster until it’s just a wall of sound. Exoskeletal membranes on the insects’ abdomens make the noise.

Katydids, on the other hand, have a more halting, staccato sound. Imagine a bug imitating a goat. That’s what a katydid sounds like. They sing by rubbing their wings together, like crickets do.

In each case, it’s the fellas bringing the noise, for the same reason that males of other species can get loud and showy. They’re trying to attract the ladies. Like dudes on the make cranking the volume in their muscle-car speakers, cicadas will try to out-rattle each other to seem more attractive to the females. Occasionally they will make a more insistent sound, such as when they’re warning the others that a hungry bird is on the prowl, Jordan said.

Katydids use their song mainly to get the word out. Hey, girl. Hey.

Both feed chiefly on plants. Katydids chew and cicadas suction fluids from plants. Neither is harmful to people, and cicadas in particular can be helpful.

“Cicadas do an awful lot for our ecology,” Kritsky said. “When they emerge from the ground, those holes are natural aeration. When it rains, the holes allow water to get down to roots. When a female lays eggs in trees, that tends to weaken the branch. Leaves can wither and die. That turns out to be a natural pruning.”

A fruit or nut tree “pruned” by cicadas puts out more fruit and nuts the following year, he said.

Animals that eat cicadas, such as birds and raccoons, have better survival rates when large numbers of cicadas are present, and the little bugs can make for a tastier Thanksgiving feast. No, it’s not what you’re thinking.

“Turkeys weigh more in the fall on years when cicadas come out,” Kritsky said.

They’ll generally stick to the trees, but in case a cicada or katydid gets inside, say in the mouth of your cat, don’t freak out. You can sweep them out if you have a broom handy, or you can toss a dish towel over one, scoop it up gently, toss the whole kit and kaboodle out of the house and let it wriggle free to resume its singing. (Ask me how I know.)

Both Kritsky and Jordan agreed: There’s no reason to kill a katydid or a cicada. Let’s enjoy the former while we await the latter.

“Cicadas are creatures of wonder, not of concern. Growing up I really liked the call of the cicada,” said Jordan, a Wisconsin native who usually heard them toward the end of summer. “It fit in so well with what was happening in the weather. It signals the end of summer. It was like music playing summer to a close.”


RISK-SPREADING AND BET-HEDGING IN INSECT POPULATION BIOLOGY

AbstractIn evolutionary ecology, risk-spreading (i.e. bet-hedging) is the idea that unpredictably variable environments favor genotypes with lower variance in fitness at the cost of lower arithmetic mean fitness. Variance in fitness can be reduced by physiology or behavior that spreads risk of encountering an unfavorable environment over time or space. Such risk-spreading can be achieved by a single phenotype that avoids risks (conservative risk-spreading) or by phenotypic variation expressed by a single genotype (diversified risk-spreading). Across these categories, three types of risk-spreading can be usefully distinguished: temporal, metapopulation, and within-generation. Theory suggests that temporal and metapopulation risk-spreading may work under a broad range of population sizes, but within-generation risk-spreading appears to work only when populations are small. Although genetic polymorphisms have sometimes been treated as risk-spreading, the underlying mechanisms are different, and they often require different conditions for their evolution and thus are better treated separately. I review the types of evidence that could be used to test for risk-spreading and discuss evidence for risk-spreading in facultative diapause, migration polyphenism, spatial distribution of oviposition, egg size, and other miscellaneous traits. Although risk-spreading theory is voluminous and well developed in some ways, rarely has it been used to generate detailed, testable hypotheses about the evolution of risk-spreading. Furthermore, although there is evidence for risk-spreading, particularly in facultative diapause, I have been unable to find any definitive tests with unequivocal results showing that risk-spreading has been a major factor in the evolution of insect behaviors or life histories. To advance our understanding of risk-spreading in the wild, we need (a) explicit empirical models that predict levels of diversifying risk-spreading for several insect populations in several environments that vary in uncertainty, and (b) tests of these models using measurements of phenotypes and their fitnesses over several generations in each environment.


A guide to identifying nocturnal nature sounds at the cottage

Going to the cottage is a great way to escape the noises of civilization. No blaring fire trucks, no car honks, no rowdy teenagers hanging out past curfew. But being less on-grid doesn’t mean you fully escape from all noises, especially the nocturnal ones produced by nature. Spend one night at a cottage and you might hear a variety of sounds from mammals, birds, insects, and reptiles. Here’s a primer on how to identify some of the coos, hoots, and caws when you’re at the cottage.

The wildlife call of a bullfrog is far from the high-pitched “ribbit” that we were taught in elementary school. Their sound is marked by a low, repetitious drone, which is made by males to attract females during breeding season. They also make the noise as a defence mechanism.

Evenings are when you’ll most often hear crickets chirping. This two-tone attraction call is made when male crickets rub their wings together. It can be heard two or three times per second when it’s 25 degrees Celsius and above. The rate slows down when the temperature drops.

Burrowing Owl

Like most owls, most of their calls can be heard at night. These owls like to reside in holes dug up by other animals such as prairie dogs or skunks. They make two kinds of calls. Listen for a soft, two-note hoo hoooo—the last note trails a bit longer—which is known as their typical hooting call. The alarm call is used when they’re in defence mode and sounds more aggressive and high-pitched. It’s often compared to the chatter sounds of a rattlesnake.

Whippoorwill

These birds might be hard to spot but their sounds are easily identifiable by their three-syllable, sing-songy whistle. Accents are on the first and last syllable, with a rise in the last. Males often repeat it throughout the night, sometimes for hours, on warm summer nights.

The coyote howl can be a frightening sound for some cottagers, but these shy animals mostly avoid confrontation with humans. They’re considered one of the most vocal wild mammals in North America, so familiarize yourself with some of their common, attention-grabbing noises: a lone, chattering howl is used to contact other coyotes, a group yip howl is used to respond, and dog-like barking is used to warn their pups to retreat to safety.


Tribe Cicadini

Brown Bunyip Tamasa tristigma, body length 20mm Brown Bunyip is one of the most commonly seen cicada in Brisbane. It is light brown in colour with black pattern on thorax. Across the abdomen there are the black, brown and light brown narrow bands. Wings are clear, males have three dark spots on tips of each forewings. They were found trees in our backyard in Eight mile Plains in Brisbane during mid summer. They are small and usually rest on the tree trunks about two meters from ground. Because they often sit in the shadow, together with their camouflaged colour, they are not easily seen. Their song is a long continuous low pitch zeep which may continue for minutes. For more information and pictures please click on here. Reference: 1. Insects of Australia, CSIRO, Division of Entomology, Melbourne University Press, 2nd Edition 1991, pp 464. 2. The cicadas of central eastern Australia - L. W. Popple, Zoology and Entomology, the University of Queensland, Australia, 2006.

NCPR provides this essential service.

&ldquoAnd the way they do that is the males have a hollow abdomen. That's basically a big resonant, buzzing drum, and they make this big loud noise. So we all hear that and so to the females.&rdquo

A large, stationary insect, making lots of noise represents a juicy target for predators such as birds or foxes.

Stager says thousands of cicadas emerging all at once is actually an evolved survival mechanism called predator saturation.

&ldquoIf they come out all at once in gigantic numbers, it pretty much guarantees that a lot of them will make it and reproduce,&rdquo he explained.

The good news for North Country residents is that Brood X is not expected to be seen in New York State.

Cornell University biologist Cole Gilbert said in an email that the closest known cicada groups are Brood VII in the Finger Lakes and Brood II in the Hudson Valley.


Yessotoxins produced by phytoplankton caused summer mass shellfish mortality events in Washington

Dying clams on Hood Canal, Rocky Bay, 2019. Credit: King et al, Harmful Algae, 2021

Back in the summers of 2018 and 2019, the shellfish industry in Washington state was rocked by mass mortalities of its crops.

"It was oysters, clams, cockles—all bivalve species in some bays were impacted," said Teri King, aquaculture and marine water quality specialist at Washington Sea Grant based at the University of Washington. "They were dying, and nobody knew why."

Now, King and partners from NOAA National Centers for Coastal Ocean Science, NOAA Northwest Fisheries Science Center, Northwest Indian College and AquaTechnics Inc. think that they have finally sleuthed out the culprit: high concentrations of yessotoxinss, which are produced by blooms of certain phytoplankton. The researchers' findings were published last month in the open-access journal Harmful Algae.

Because yessotoxins are not a threat to human health, their presence in Washington has not been closely monitored. The researchers dug through data that had been collected by the NOAA Northwest Fisheries Science Center and NOAA National Centers for Coastal Ocean Science for different purposes, coupled it with current observations from the SoundToxins phytoplankton monitoring program, and discovered that these algae species, Protoceratium reticulatum and Akashiwo sanguinea, are correlated with shellfish mortality events stretching as far back as the 1930s.

The algae species Protoceratium reticulatum, seen under a microscope. Credit: Teri King/Washington Sea Grant

In 2018 and 2019, with SoundToxins partners' eyes on the water, and reports of dying shellfish from the Washington Department of Fish & Wildlife and the shellfish industry, the research team was able to collect shellfish and water samples for analysis. This set the table to help answer the mystery of what was causing summer mortality in Washington state shellfish.

These findings have significant implications for shellfish growers in the region.

"We are working towards being able to help growers count the cells of yessotoxin-producing organisms in the water and correlate it to an action level," King explained. "SoundToxins has been conducting similar work for the Washington Department of Health for three 'human health' marine biotoxins since 2006. Adding the 'shellfish killing' plankton species to the real-time mapping capability of the SoundToxins partnership would allow for shellfish producers and natural resource managers to make informed decisions, such as harvesting their product early or otherwise strategizing to save as much crop as possible."

Clams on Rocky Bay watershed, Case Inlet, July 2019. Credit: Teri King/Washington Sea Grant

King said this research is also a demonstration of the value of partnerships between shellfish producers, plankton monitors, Native tribes, agencies and researchers.

"We were a team of oceanographers, biologists and chemists working together to answer these questions," King said. "People are able to think differently when you have different people at the table."

Sometimes, it's even the key to solving the longstanding mysteries that have been taking place right in your backyard.