Order Plecoptera

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933
Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00036-X
Copyright © 2015 Elsevier Inc. All rights reserved.
Order Plecoptera
R. Edward DeWalt
Prairie Research Institute, Illinois Natural History Survey, University of Illinois, Champaign, IL, USA
Boris C. Kondratieff
C. P. Gillette Museum of Arthropod Diversity, Department of Bioagricultural Sciences and Pest Management, Colorado State University, Fort Collins, 
CO, USA
John B. Sandberg
California Department of Fish and Wildlife, CSUC Research Foundation, California State University, Chico, CA, USA
Chapter 36
Chapter Outline
Introduction 933
Overview of the Insect Order Plecoptera 933
Phylogeny and Biogeography 933
General Biology 935
External Anatomy 935
Immature Stoneflies 935
Adult Stoneflies 936
Eggs 936
Life History 937
General Ecology and Behavior 939
Macro- and Micro-habitat Usage 939
Physiological Constraints on Distribution and Survival 939
Feeding Behavior 939
Parasites and Symbionts of Stoneflies 940
Conservation of Stoneflies 940
Behavior 940
Vibrational Communication (Drumming) 940
Diel Periodicity of Adults 942
Collecting and Rearing Stoneflies 942
Collecting Nymphs and Adults 942
Collecting Nymphs 942
Collecting of Adults 943
Preservation Techniques and Labeling 945
Preservation Techniques 945
Labeling Specimens 946
Rearing of Stoneflies 946
Acknowledgements 947
References 947
INTRODUCTION
Overview of the Insect Order Plecoptera
The order name Plecoptera comes from the Latin plecto, 
meaning “folded,” and the Greek pteron, meaning “wing,” 
and refers to the ability of adults to fold their wings. The order 
is an ancient one, with the fossil record extending into the 
Pennsylvanian, about 300 million years ago (Béthoux et al. 
2011). A combination of external characters identifies a stone-
fly from other insect orders: most adults with two pairs of 
folded wings, two multi-segmented cerci in nymphs and most 
adults (reduced in some families), a three-segmented tarsus, 
and two claws on each leg.
Stoneflies are hemimetabolous with egg, nymph, and 
adult life stages. Nymphs are almost exclusively aquatic and 
can be found in streams of all sizes, temperature regimes, 
and permanence. The greatest species richness is found in 
mountains of temperate latitudes, but considerable diversity 
can be found in warm-water streams. Some species live 
along the wave-swept shores of high altitude or latitude 
lakes. Species reside on every continent except Antarctica.
Phylogeny and Biogeography
The Plecoptera are a basal, aquatic order of the lower 
Neoptera occurring on all continents except for Antarctica 
(Fochetti and Tierno de Figueroa, 2008). The monophyly 
of the Plecoptera is supported by several apomorphic char-
acters such as gonads forming loops, two superimposed 
seminal vesicles, each forming a loop, the presence in the 
SECTION | VI Phylum Arthropoda934
larvae of strong oblique, intersegmental ventro-longitudinal 
muscles for laterally undulating swimming, and the general 
absence of ovipositors (with a few exceptions where they 
are secondarily derived) (Zwick, 2000). Terry and Whiting 
(2005), using a combined molecular and morphological 
approach, also presented evidence that the Plecoptera are 
monophyletic.
There are more than 3704 species recorded in Plecoptera 
Species File (PlecSF), including 225 fossil species (DeWalt 
et al., 2012a) (Table 36.1). Previously, the number of spe-
cies globally was underestimated by about 85% (Zwick, 
2000). The advent of PlecSF, populated by the substantial 
database of Dr. Zwick, has permitted a much more accurate 
picture of the richness of stoneflies worldwide.
Stoneflies have been placed into various hierarchical 
classifications proposed by several authors including Ricker 
(1950, 1952), Illies (1965, 1966), Zwick (1973, 1980), and 
Nelson (1984). Stewart and Stark (2002) provided a suc-
cinct summary of the higher classification of the Plecoptera 
emphasizing the Nelson (1984) and Zwick (2000) analyses. 
Zwick (2000) arranged the 16 recognized extant families 
into four groups under two suborders: the largely north-
ern hemisphere Arctoperlaria and the exclusively southern 
hemisphere Antarctoperlaria. Twelve of these families are 
arctoperlarian and four are antarctoperlarian (Table 36.1).
McCulloch (2010), using molecular sequence data, esti-
mated that the Arctoperlaria and Antarctoperlaria diverged 
during the Jurassic period, which is consistent with their 
vicariant formation driven by the breakup of Pangaea. 
McCulloch (2010) further concluded that a single Antarc-
toperlaria lineage apparently dispersed into the southern 
hemisphere during the Cretaceous period. The Antarc-
toperlaria, a relatively small group of approximately 343 
species (DeWalt et al., 2012a) distributed in Australia, 
TABLE 36.1 The Higher Classification of the Extant Plecoptera, Number of Valid, Extant Species, and World 
Distribution. Numbers of Species Modified from Plecoptera Species File (DeWalt et al., 2012a)
Hierarchy Taxon 1Valid Species Distribution
Suborder Arctoperlaria 3136 N & S hemisphere
“Group” Euholognatha 1521 N & S hemisphere
Family Capniidae 278 N Am, Europe, N Africa, Asia
Leuctridae 356 N Am, Europe, N Africa, Asia
Nemouridae 656 N Am, Europe, N Africa, Asia
Notonemouridae 118 Southern Africa, Australia, Madagascar, New Zealand, S Am [Argentina, 
Chile], Tasmania
Taeniopterygidae 105 N Am, Europe, N Africa, Asia
Scopuridae 8 Korean peninsula, Japan
“Group” Systellognatha 1615 N & S hemisphere
Pteronarcyidae 12 N Am, E Asia
Peltoperlidae 67 Asia, N Am
Styloperlidae 10 China, Taiwan
Perlodidae 309 N Am, Europe, N Africa, Asia
Perlidae 1024 N Am, Central Am, S Am, Europe, N Africa, Asia
Chloroperlidae 193 N Am, Europe, N Africa, Asia
Suborder Antarctoperlaria 343 S hemisphere
Family Austroperlidae 15 Australia, New Zealand, Tasmania, S Am (Argentina, Chile
Diamphipnoidae 6 S Am (Argentina, Chile)
Eustheniidae 22 Australia, New Zealand, Tasmania, S Am (Argentina, Chile)
Gripopterygidae 300 Australia, New Zealand, Tasmania, S Am (Chile to Columbia, Brazil)
Total extant 3479
1An additional 225 fossil species are known.
935Chapter | 36 Order Plecoptera
New Zealand, Tasmania, and South America, are united by 
the synapormorphies of sterna depressor of the fore tro-
chanter, absence of a tergal depressor of the trochanter, and 
presence of floriform chloride cells (Zwick, 1973, 1980, 
2000). Interestingly, as a group, adults apparently do not 
use vibrational communication which is common in species 
of the Arctoperlaria (Stewart, 2001). The Antarctoperlaria 
are divided into two superfamilies (Table 36.1): Eusthe-
nioidea (Eustheniidae and Diamphipnoidae) and Gripop-
terygoidea (Austroperlidae and Gripopterygidae) (Zwick, 
2000). The taxa of these two suborders have been treated 
by Illies (1963), McLellan (1990, 2006), Theischinger and 
Cardale (1987), Suter and Bishop (1990), McLellan and 
Zwick (2007), and Stark et al. (2009).
The Arctoperlaria stoneflies account for 3136 extant 
species (Table 36.1). Overall support for the suborder is 
weak, but the mate-finding behavior of drumming (Stewart, 
2001) strongly supports monophyly of this large group of 
taxa (Zwick, 2000). Zwick (2000) recognized the “groups” 
Euholognatha and Systellognatha within Arctoperlaria. 
Euholognathan families are united by three synapomor-
phies: the unpaired corpus allatum fused to the aorta, a soft 
egg chorion, and segmental nerves crossing under the lon-
gitudinal abdominal muscles (Zwick, 2000). This grouping 
includes the Scopuridae and the Nemouroidea (Capniidae, 
Leuctridae, Nemouridae, Notonemouridae, and Taeniop-
terygidae) (Table 36.1). The Notonemouridae are consid-
ered relics of the breakup of Gondwanaland (Zwick, 2000) 
and are the only Euholognathan family that occurs in the 
southern hemisphere, with more than 118 described species 
described from southern Africa, South America, Madagas-
car,New Zealand, and Australia. The Scopuridae are a relict 
family restricted to the Korean Peninsula and Japan (Jin and 
Bae, 2005). The remaining five families have widespread 
Holarctic and Oriental distributions.
The Systellognatha includes two recognized superfami-
lies, the Pteronarcyoidea (Pteronarcyidae, Styloperlidae, 
and Peltoperlidae) and Perloidea (Perlodidae, Chlorop-
erlidae, and Perlidae). This large group includes most of 
all extant stoneflies with approximately 1615 recognized 
species (DeWalt et al., 2012a). Zwick (2000) reviewed the 
apomorphies that unite the Systellognatha, including pro-
posing the female accessory glands on the receptacular duct 
as another apomorphic character. The monophyly of Ptero-
narcyoidea, including the three families noted previously, is 
supported by the paired but closely adjacent corpora allata 
and fused ends of the tegumental head nerves (Zwick, 2000). 
The well-known adults of the “giant stoneflies or salmon-
flies,” the Pteronarcyidae, are a small group of Nearctic and 
East Palearctic species characterized by ample wing vena-
tion and larvae with branched gills both on the thorax and 
first segments of the abdomen. Zwick (2000) suggested a 
new apomorphy of the Pteronarcyidae, the ability of the 
larvae to “curl the forebody under” when disturbed. The 
phylogenetic relationship of the Styloperlidae and Peltoper-
lidae was discussed by Uchida and Isobe (1989). Species of 
Styloperlidae are known from China and Taiwan, whereas 
the Peltoperlidae are widespread in North America and Asia 
(Stark, 1989; Stark and Sivec, 2007; DeWalt et al., 2012a).
The monophyly of the diverse Perloidea is supported 
by the occurrence of predaceous larvae, at least in latter 
instars, possessing slender mandibles lacking a mola and 
a distinctly toothed lacinia, and the greatly enlarged para-
glossae (Zwick, 2000). Relationships among the families 
of the Perloidea are not fully resolved (Zwick, 2000), with 
several hypotheses available (Zwick, 1973, 1980; Stewart 
and Stark, 2002). Stewart and Stark (2008) listed numer-
ous references for the identification of the Nearctic mem-
bers of this group: Stark et al. (2009) for South America; 
Zwick (2004) for the Palearctic; and Fochetti and Tierno de 
Figueroa (2008) for other parts of the world.
GENERAL BIOLOGY
External Anatomy
Immature Stoneflies
Immature stages of stoneflies may be referred to as nymphs, 
naiads, or larvae, but we will use “nymph” throughout the 
chapter. Most nymphs are drab in color, but some are highly 
patterned. General characteristics include a head with chew-
ing mouthparts, two stout antennae and cerci, three pairs 
of legs with two tarsal claws each, an abdomen, and thorax 
(Figure 36.1). The head is prognathus with two compound 
eyes and two or three ocelli. Mouthparts are of a chewing 
design and general food habits can be surmised from the 
shape of the mandibles and maxillae (Figure 36.2(a) and (b)).
The thorax is composed of pro-, meso-, and metatho-
racic segments (dorsally nota). The meso- and metanota 
support the developing wingpads beginning about halfway 
through development. Legs are composed of a round coxa, a 
stout femur, a more slender tibia, and three tarsal segments, 
the most terminal of which has two claws. Often the femur 
and tibia have long silky setae that act as swimming hairs.
The abdomen, which terminates in two long, stout cerci, 
is composed of 10 visible segments, the first often obscured 
dorsally by the thorax. The female of nymphs may often be 
determined by a gap in the posterior hair fringe of the eighth 
abdominal segment.
Gills are often present and may be found in combina-
tions on the head, thorax, or abdomen. Shepard and Stewart 
(1983) and Stewart and Stark (2002) provided a detailed dis-
cussion of the placement, type, and evolution of gills in Arc-
toperlaria stoneflies. Antarctoperlarian stoneflies also have 
gills, but these are mostly located on several abdominal seg-
ments as paired beaded structures, tufts of filamentous gills, 
clusters of filiform gills about the anus, or beaded structures 
arising from the paraproct region (Wichard et al., 2002).
SECTION | VI Phylum Arthropoda936
Adult Stoneflies
Most adult stoneflies have two pairs of pleated wings with 
simple venation (see Stewart and Stark, 2008). Hindwings 
are often larger than the forewings. Pleats allow stonefly 
adults to fold their wings, and this feature gives the order its 
name (Figure 36.3). The relative development of wings var-
ies from full to no wings. Polymorphism in wing develop-
ment occurs within and across species populations. No clear 
selective advantages are known for these conditions, but both 
environmental and genetic reasons for wing polymorphism 
have been postulated.
Males have complex internal or external primary and 
secondary sexual structures. The shape and armature of 
these structures are used to describe new species. For exam-
ple, capniid stoneflies have an external epiproct that is often 
separated into an upper and lower limb (Figure 36.4(a)). 
Conversely, perlids often have an internal aedeagus, with 
fields of spinulae and sclerotized plates, the organization 
of which is species specific (Figure 36.4(b)). Researchers 
often must extrude these to accomplish species-level deter-
minations. Males of the Arctoperlaria may have a ventral 
vesicle or hammer that is used in species-specific vibra-
tional communication (Figure 36.5(a) and (b)).
Female external genitalia are often more structurally 
conservative than in males. A subgenital plate often occupies 
all or part of the eighth abdominal sternum (Figure 36.6(a) 
and (b)), marking the opening of the gonopore and aiding 
in the formation of an egg mass. The gonopore receives 
sperm either from the epiproct, paraproct, or aedeagus and 
may be deposited internally or siphoned in from external 
placement.
Eggs
Although eggs are generally spherical, some with trian-
gular cross-sections are common among the perlodids 
(Kondratieff, 2004). Many euholognathan stoneflies have 
membranous eggs that are contained within a gel matrix 
(Figure 36.7(a)). Other stoneflies have sclerotized eggs with 
grooves and follicular cell impressions (Figure 36.7(b)). 
The ability to detect cryptic species is often possible using 
scanning electron microscopy to reveal egg chorionic sculp-
turing (Stark, 2004).
FIGURE 36.1 External morphology of stonefly nymph. (a) dorsal and (b) ventral. Figures modified from Stewart and Stark (2002) with permission from 
B. J. Armitage of the Caddis Press.
937Chapter | 36 Order Plecoptera
Life History
Stoneflies are hemimetabolous, exhibiting incomplete or 
gradual metamorphosis with egg, nymph, and adult life 
stages. The vast number of species reproduce in the spring 
and early summer; however, the so-called “winter stone-
flies” (capniids and taeniopterygids) emerge and oviposit 
in mid to late winter. Relatively few late summer and fall 
emerging species are found in the families of Leuctridae, 
Nemouridae, Perlidae, and Perlodidae.
Reproduction is semelparous in all stoneflies. Females 
live 1 to 2 weeks after transformation and produce one or 
more egg masses before dying. They drop egg masses from 
various heights to the stream surface or land atop the water, 
dislodging the egg masses. The eggs of euholognathans are 
clad in a gelatinous matrix and stick to substrates easily. 
Other stoneflies have filaments or a partial membranous cap 
that adheres to substrates, whereas the eggs of other spe-
cies are simply entrained in the sediments. Egg hatching 
may require less than a month, hatch many months later, or 
undergo cohort splitting in which eggs hatch at various time 
over several months, causing a large range in size of the 
nymphs in the stream and potentially leading to a stonefly 
egg seed bank (Zwick, 1996; Snellen and Stewart, 1979).
Most species are annual, or univoltine, being tied to the 
flush of resources that result from autumnal leaffall (Stewart 
and Stark, 2002). A smaller proportion of species require more 
(a)
(b)
FIGURE 36.2 Mandibles (left) and maxillae (right) of. (a) preda-
tory stonefly in family Perlidae; and (b) detritivore in the family 
Taeniopterygidae.
FIGURE 36.3 Fore (top) and hindwings (bottom) of Pteronarcys scotti 
Ricker.
(a)
(b)
FIGURE 36.4 Genitalia of stoneflies. (a) Male Allocapnia granu-
lata (Claassen) (Capniidae) epiproct over process on eighth tergum; (b) 
Aedeagus of Acroneuria internata (Walker).
SECTION | VI Phylum Arthropoda938
than 1 year to complete their growth. These semivoltine spe-
cies are often large bodied, live in spring-influenced habitats, 
or inhabit cold climates. Some nymphs require 3 years to com-
plete their growth. Only one species, the Palearctic Nemurella 
pictetii (Klapálek), has been demonstrated to have more than 
one generation per year (Lieske and Zwick, 2008). It also has 
been shown to be semivoltine at high elevation (Brittain, 1978).
Diapause is a common feature in stoneflies. Species 
that inhabit seasonally dry or warm-water streams often 
undergo diapause as eggs or early instar nymphs. Pre-
sumably, this reduces their risk to encountering drought, 
high water temperatures, and low oxygen concentrations 
as nymphs. Nymphs of winter adult-emerging species 
undergo diapause during the warmer months deep in the 
substrate (Coleman and Hynes, 1970). Egg diapause can 
last a year or more (Snellen and Stewart, 1979; Sandberg 
and Stewart, 2004). Both univoltine and semivoltine spe-
cies may diapause; in the latter, this may extend the life 
cycle into a fourth year for Pteronarcys californica New-
port (DeWalt and Stewart, 1995).
Up to 30 or more species may inhabit a pristine stream 
within mountainous areas of the northern hemisphere 
(DeWalt and Stewart, 1995). Each species has a unique life 
history, leading to unique phenologies of emergence, ovipo-
sition, and growth (Brittain, 1990). These differences lead 
to a succession of adults throughout the year, potentially 
limiting competition for space and other resources. Life his-
tory information is becoming increasing important as the 
use of taxon traits in water quality assessment and conser-
vation biology increases (Poff et al., 2006).
(a)
(b)
FIGURE 36.6 Female subgenital plates. (a) Leuctra sp.; (b) Agnetina 
flavescens (Walsh).
(a)
(b)
FIGURE 36.5 (a) Ventral abdominal terminalia of Amphinemura pal-
meni (Koponen); (b) Hammer (arrow) of Acroneuria lycorias (Newman).
939Chapter | 36 Order Plecoptera
GENERAL ECOLOGY AND BEHAVIOR
Macro- and Micro-habitat Usage
Stoneflies are distributed worldwide, with the exception of 
areas of permanent ice cover and large deserts (Fochetti and 
Tierno de Figueroa, 2008; DeWalt et al., 2012a). Nymphs 
have been collected from a wide range of sizes of fresh-
waters, from small seeps where there is little perceptible 
flow to the largest of rivers (Zwick, 1973). Adults generally 
remain close to the natal habitat.
Most stonefly species are associated with cool and 
cold waters and are the most diverse in ancient, temperate 
mountain areas the world over (Brittain, 1990). In North 
America, there has been significant radiation of species 
into warm-water streams. This radiation has occurred in 
many families, most notably in the Perlidae, where greater 
surface area of gills improves the ability to obtain oxy-
gen. A similar radiation has occurred in Central and South 
America (about 350 species known), where approxi-
mately 71% of the known species are perlids (DeWalt 
et al., 2012a). Species of many families have also radiated 
into intermittent streams, where diapause during summer 
months is a common life history strategy (Stewart and 
Stark, 2002).
A small number of species inhabit cool to cold oligo-
trophic lakes at higher latitudes or altitudes. These are usu-
ally a subset of the regional species pool that also inhabit 
nearby rivers, but find similar conditions in the wave-swept 
shores of lakes. One exception is Capnia lacustra Jewett, 
which lives its entire life within the confines of Lake Tahoe 
(Nevada and California, USA) (Jewett, 1965). Warming 
climates and increased pollution threaten the existence of 
stoneflies in large lakes throughout the world.
Most species occupy fast-flowing, riffle areas of streams 
where coarse mineral substrates dominate. Large per-
lids often occupy the undersides of boulders and cobbles, 
whereas detritivores inhabit leaf packs and woody debris 
piles. The nymphs of one Australian species, Riekoperla 
darlingtoni (Illies), can live in damp terrestrial habitats 
or in a burrow in the stream bank if the stream dries out 
(Australian Freshwater Invertebrates, 2012). Sand-bot-
tomed streams support stoneflies, but it is the wood in these 
streams where most of the production takes place (Benke 
et al., 1984). No stonefly species are specialized to burrow 
in sand, as do some mayflies and midges.
Physiological Constraints on Distribution 
and Survival
Stoneflies generally need highly oxygenated waters in 
which to live. Consequently, they are relatively intolerant 
of high stream temperatures (Quinn et al., 1994). Flowing 
waters ameliorate the effects of higher stream temperature, 
and time of year had a significant effect upon lethality of 
high temperature for 50% of a laboratory population of the 
perlid Paragnetina media (Walker) (Heiman and Knight, 
1972). This will be a fruitful area of research as the climate 
of the planet warms.
Stoneflies are absent from permanently ice-covered areas. 
Their ability to tolerate freezing is poorly understood (see 
Danks, 2007). Walters et al. (2009) found that nymphs of 
Nemoura arctica Esben-Petersen in subarctic Alaska, USA 
actually survived freezing to −15 °C for extended periods. 
The production of glycerol and ice-binding antifreeze proteins 
helped them resist the effects of freezing. This probably rep-
resents the extreme physiological response to freezing within 
Plecoptera. Bouchard et al. (2009) found freeze tolerance to 
−9 °C in winter-emerging adults of two Allocapnia species 
in Minnesota, USA. Darkly pigmented bodies and the use of 
thermal refuge are probably equally important as physiological 
responses in conferring freeze tolerance in winter stoneflies.
Feeding Behavior
Considerable literature exists for feeding of nymphs (see 
Stewart and Stark, 2002). All nymphs feed, whereas only 
a subset of adults does (Tierno de Figueroa and Sanchez-
Ortega, 1999). Adult feeding is largely limited to species 
whose nymphs eat detritus, but some predatory species also 
(a)
(b)
FIGURE 36.7 Eggs of stoneflies. (a) Gelatinous egg mass of Taeniopteryx 
burksi Ricker and Ross, From Frison (1935); (b) highly sculptured egg of 
Neoperla sabang Sivec and Stark. From Sivec and Stark, 2011.
SECTION | VI Phylum Arthropoda940
feed as adults, usually on pollen. Much of this research 
was painstakingly done by estimation of volume of various 
gut contents (Alan, 1982). However, stable isotope analy-
sis sometimes shows that feeding is often more complex 
than gut contents suggests (Miyasaka and Genkai-Kato, 
2009). The presence of specific enzyme activity may also 
be of great use in determining the types of food consumed 
throughout the life cycle (Tierno de Figueroa et al., 2011b). 
Ontogenic shifts in feeding have been documented, even 
within species that are thought of as highly predaceous 
(Miyasaka and Genkai-Kato, 2009).
Parasites and Symbionts of Stoneflies
Stoneflies are parasitized by a wide range of organisms 
including nematodes, larval mites, wasps, midges (Chiron-
omidae), fungi, and possibly even the bacterium Wolbachia. 
Giberson et al. (1996) found that the chironomid midge 
Nanocladius (Plecopterocoluthus) sp. parasitized nymphs of 
Pteronarcys biloba Newman in an eastern Canadian stream. 
Other midges may be only phoretic on nymphs of stoneflies. 
Nymphs are often parasitized by larval mites, the mites shift-
ing to the adult as thenymph molts. Females of Neoperla 
stewarti Stark & Baumann, an eastern North American perlid 
stonefly, have often been found with a hymenopteran parasit-
oid pupa in the spermathecum (DeWalt, unpublished data). 
Mermethidae nematodes are common endoparasites of fresh-
water arthropods. Gamboa et al. (2012) reported an aston-
ishingly high incidence of parasitism in two Anacroneuria 
species in Venezuelan rivers. The intracellular parasite Wol-
bachia (a bacterium), thought to infect a large proportion of 
insects, has yet to be documented in stoneflies. White and 
Lichtwardt (2004) reported that fungal symbionts in the order 
Harpellales commonly occur in the gut of stoneflies. Presum-
ably these fungi aid digestion of the decayed plant material 
that some stoneflies consume (Lichtwardt, 1986).
Conservation of Stoneflies
Ricciardi and Rasmussen (1999) recognized that aquatic 
fauna and habitat were being lost at a much greater rate 
than terrestrial ones. Stoneflies are among the most envi-
ronmentally sensitive of freshwater aquatic fauna (Master 
et al., 2000). Zwick (1992) early recognized their loss in 
European rivers, with the nearly complete disappearance of 
large river species and the reduction of refugia in the moun-
tains owing to acid precipitation. DeWalt et al. (2005) dem-
onstrated that >20% of the fauna of Illinois, USA had been 
lost in the twentieth century. It is feared that much of the 
developed world is similarly affected.
We cannot depend on the type of endangered species 
research conducted on vertebrates to help protect aquatic 
invertebrates (Strayer, 2006). However, we can use modern 
methods to capture information from museum specimens 
and new collections to help answer important conservation 
questions about stoneflies (DeWalt et al., 2012b). Species 
distribution modeling may be useful in predicting histori-
cal ranges using these data. Predicting individual species 
ranges, biodiversity hotspots, and reactions by species to 
climate change (Tierno de Figueroa et al., 2010) can all 
result from this approach (Cao et al., 2013). Species will 
continue to shrink their ranges and we must prepare to help 
them use corridors for migration or even assist them to 
move in the future.
Behavior
Vibrational Communication (Drumming)
Drumming is a type of adult intersexual vibrational commu-
nication practiced by arctoperlarian stoneflies (Rupprecht, 
1980; Stewart, 2001; Tierno de Figueroa et al., 2011a). These 
signals are usually produced by tapping or rubbing the pos-
terior-ventral abdominal segments upon suitable substrates. 
Drumming exchanges between sexes are thought to transmit 
information including the relative distance between individu-
als, relative location of the female, and the species identity. 
This behavior assists stonefly reproduction by bringing the 
sexes together within complex riparian habitats.
Drumming signals have been described for approxi-
mately 192 species worldwide (Table 36.2). Observations of 
TABLE 36.2 Summary of Drumming Signal Diversity 
in Stoneflies, Families, Number of Species Studied, 
Structures Used, Signaling Method. Citations for 
Primary Literature Too Numerous to List Here
Family
Number of 
Species
Male 
Structures
Signaling 
Method
Capniidae 12 None or 
vesicle
Percussion
Chloroperlidae 8 None or 
lobe
Percussion & 
tremulation
Leuctridae 7 Vesicle Percussion
Nemouridae 5 Vesicle Percussion
Peltoperlidae 15 Knob Percussion 
& rub
Perlidae 45 None or 
hammer
Percussion 
& rub
Perlodidae 83 None or 
lobe
Percussion 
& rub
Pteronarcyidae 9 None Percussion
Taeniopterygidae 8 None or 
vesicle
Percussion
941Chapter | 36 Order Plecoptera
the southern hemisphere stonefly suborder Antarctoperlaria 
in Australia, New Zealand, and South America have failed 
to detect drumming, which suggests that this suborder may 
not have evolved drumming. Antarctoperlarian stoneflies 
have undoubtedly developed other mate-finding behaviors 
(Stewart and Sandberg, 2006). Multiple drumming methods, 
specialized drumming structures, and the range of complex-
ity of signal types suggest that stonefly drumming behavior 
is one of the most diverse and complex forms of vibrational 
communication known in insects (Stewart, 2001).
Male stoneflies produce vibrational signals using three 
methods: percussion (tapping the distal postero-ventral abdo-
men onto the substrate), rubbing (the distal postero-ventral 
abdomen is scraped across the substrate), and tremulation (quick 
rocking movements of the entire body occur on the substrate). 
The percussive signal is most common and is accomplished 
by striking the unmodified abdomen or modified structures 
such as a vesicle (Figure 36.5(a)) or a knob or hammer (Figure 
36.5(b)) against a substrate. Rub calls are not percussive signals 
and are most prevalent in the Peltoperlidae and Perlidae, where 
the ninth sternum is modified as a textured ventral hammer or 
knob. The tremulation method has only been documented in 
the Chloroperlidae (Alexander and Stewart, 1997). So far as 
currently known, females only produce simple, ancestral, per-
cussive answer signals and rarely enter duets with males after 
they have been mated (Stewart, 2001).
Drumming duets are displayed in Figure 36.8(a) and (b). 
The red vertical lines represent the male call beats and the 
blue vertical lines are the female answer beats. Male calls 
last from 1 to 24 s; female answers are usually consider-
ably shorter. Interbeat intervals (indicated by i1–i4) are the 
rest intervals between beats and their duration is measured 
in milliseconds. The sequence of communication between 
the sexes follows three basic types: male-call (♂C); female-
answer (♀); and male-response (♂R). The third and less 
common male-response signal occurs only after the female 
answers the male call and is usually less complex than 
the initial male call or composed of beats with distinctly 
FIGURE 36.8 Stonefly drumming signals. (a) A simple duet, male call followed by female answer; (b) A three-way exchange, male call, female answer, 
and male response with different beat count and interval; and (c) A male diphasic call with three distinct interval patterns.
SECTION | VI Phylum Arthropoda942
different interbeat intervals. The female answer signal can 
follow the male call (sequenced) or begin before the male 
call has ended (interspersed or overlapped). The duet in 
Figure 36.8(a) represents a “two-way exchange,” signify-
ing a call and answer, whereas Figure 36.8(b) represents 
a “three-way exchange” whereby the male responds to 
the female answer. Recently, a four-way exchange was 
described and consisted of a call–answer–call–answer 
sequence with different beat intervals and no immediate ter-
minal response from the female (Sandberg, 2011a).
Male call complexity varies tremendously across spe-
cies. Monophasic signals have nearly even interbeat inter-
vals (Figure 36.8(a)) (Stewart and Sandberg, 2006). Varied 
beat-interval signals change interbeat intervals as the call 
progresses (Figure 36.8(b)) (Sandberg, 2011b). Diphasic 
calls are similar to varied beat-interval calls but contained 
three consecutive signal types composed of initial monopha-
sic beats, followed by a transition of decreasing varied beat-
interval beats, and ending with monophasic intervals (Figure 
36.8(c)). Table 36.2 provides a summary of known drum-
ming signal diversity for stonefly families relative to special-
ized abdominal structures and method of signal production.
Diel Periodicity of Adults
Adult behavior in the field has been little studied. This is 
undoubtedly because of the adult’s more mobile nature. 
DeWalt and Stewart (1995), working in the southern Rocky 
Mountains of the USA, found that many species emerged 
as adults just after dark. Males spent the next hours drum-
ming and seeking females to mate. Most species spent the 
remainder of the night under streamside cobbles, or more 
commonly within the leaf-choked bases of willows.Day-
light cued the ascendance of adults to the tops of the wil-
lows. Here they basked in the sun, males sought mates, 
females produced egg batches, and females flew to the 
stream for egg deposition. This is a pattern that can be 
counted on in open mountain streams. Elsewhere, we have 
observed adults transforming during the day. Poulton and 
Stewart (1988), working in the northern Rocky Mountains, 
USA, found that peak flight to the stream for male Calineu-
ria californica (Banks) was related to evapotranspiration 
potential, which led to their flying to the stream to drink. 
For females, the cue was light intensity. Males flew to pool 
sections in the mid-afternoon, whereas females flew to riffle 
sections during the last few hours before dark, presumably 
to drop egg masses. Much more work is necessary to eluci-
date what adults do once they leave the stream.
COLLECTING AND REARING STONEFLIES
Collecting Nymphs and Adults
Stonefly adults and nymphs may be collected from a 
wide variety of lotic, lentic, and terrestrial habitats. Many 
techniques are available for the collection and preserva-
tion of adults and nymphs. A thorough review of available 
equipment and techniques for collecting aquatic insects is 
provided in Merritt et al. (2008; and references therein). In 
addition, Voshell (2002) described the basic types of tech-
niques and equipment used to study aquatic invertebrates, 
including stoneflies. Steyskal et al. (1986) and Schauff 
(2005) included techniques applicable to collecting and 
preserving stoneflies, especially for permanent museum 
research or voucher collections.
The techniques employed will necessarily vary if the 
design involves qualitative, semiquantitative, or quantita-
tive sampling. For example, qualitative methods generally 
provide only information such as taxa present, whereas 
semiquantitative methods provide relative abundance and 
most quantitative methods allow for estimates of the num-
ber of individuals and biomass on a per-unit basis. Kerans 
et al. (1992) and Rosenberg et al. (2008) discussed the con-
siderations for use of any of these sampling protocols. The 
size of the mesh used in the catch net can impact sampling 
results (e.g., Mutch and Pritchard, 1982). Aquatic inverte-
brate sampling protocols are usually designed for wadeable 
streams (Cuffney et al., 1993). Most USA states have spe-
cific protocols for sampling streams (Barbour et al., 1999; 
Carter and Resh, 2001). Protocols have been developed for 
freshwater wetlands that may include stonefly taxa (Turner 
and Trexler, 1997; USEPA, 2002; DiFranco, 2006).
Major aquatic and terrestrial collecting techniques 
are summarized next. However, basic collecting supplies 
should include glass or plastic vials, a pair of forceps 
(featherweight or standard), a shallow white pan (enamel or 
plastic), and a preservative. Voshell (2002) recommended a 
pan size of 15.2 × 22.9 cm. Selection of substrates for sam-
pling will vary with the macro- and micro-habitats of inter-
est. Voshell (2002) and Merritt et al. (2008) reviewed the 
numerous types of habitats that include stoneflies.
Collecting Nymphs
Dip Net
The routine sampling device for collecting stonefly and 
many other aquatic insect immatures is the D-frame dip 
net. It is most often used qualitatively, but semiquantitative 
samples may be obtained if a consistent area is sampled. 
A dip net is the most versatile sampling device because it 
can be used in nearly all aquatic microhabitats. A 500- to 
1000-μm mesh bag is sufficient in most cases. Disturbing 
microhabitats (riffle substrates, leaf packs, logs, and root 
mats of undercut banks) upstream of the dip net will nearly 
always be productive (see Voshell’s [2002] description of 
the “stonefly stomp” technique). Nymphs may be picked 
alive from the sample debris by placing aliquots of the sam-
ple into a white tray with clear stream water or they may be 
removed from the preserved sample in the laboratory.
943Chapter | 36 Order Plecoptera
Life history studies often require much finer mesh nets so 
that they collect the smallest nymphs. In this way, a more accu-
rate growth histogram may be plotted (Mutch and Pritchard, 
1982). A two-stage ”life history” dip net may be designed 
by attaching a long, conical, fine mesh bag (about 300 μm 
mesh) to the downstream end of a dip net with a coarser 
(500- to 1000-μm) mesh inner bag (DeWalt and Stewart, 
1995). A plankton bucket may be attached to the fine mesh 
end to catch the finer sample fraction (smallest nymphs, eggs, 
silt, etc.). The two stages separate the coarse sample fraction 
from the fine, improving efficiency of sample processing.
Kick Screen
A kick screen or screen barrier net (a mesh sheet stretched 
between two poles) is often useful for deeper habitats or habi-
tats with larger rocky substrate or log jam piles. This strictly 
qualitative method will permit rapid assessment of the pres-
ence of large species. A commercially available kick screen 
(BioQuip™) consists of two 122 -cm poles with a 91 × 91-
cm mesh Lumite™ screen. Usually two persons are required 
to use a kick screen most effectively, especially with greater 
flows. Voshell (2002) described the operation of this device.
Hess, Surber, Drift Net, Peterson Grab, Ponar, 
Stovepipe Samplers
All of these devices are designed to sample a known area; 
hence, if carefully used they will produce quantitative sam-
ples. Unfortunately, they are not as versatile as dip nets and 
kick screens and may not be effectively used on very large 
substrates or in undercut banks. They work best where sub-
strate size is cobble size or smaller. A Hess sampler is a cylin-
drical device that is driven into the substrate. The upstream 
portion has coarse mesh that allows water to flow through to 
a downstream opening with a long, tapered fine mesh bag. 
This bag often has a plankton bucket to improve handling 
of specimens and debris. A Surber sampler (Surber, 1937) is 
composed of an open metal frame that demarks the area being 
sampled. To this frame is connected a conical net of ≤ 500-
μm mesh, often with a plankton bucket attached. Stonefly 
nymphs are known to drift (Benke et al., 1991) in rivers and 
streams and have been collected effectively in a drift net. This 
cylindrical net may be suspended in the river current to assess 
the phenology and rate of drift. If the current speed is known, 
a catch per unit volume may be assessed. Petersen, Ponar, 
and Ekman grabs are devices with heavy jaws used to sample 
fine substrates, usually in slowly flowing or still water. The 
latter may be used in conjunction with a pole release, and 
hence may be used in waters up to 1.5 m deep. The other 
grabs may be used in deeper waters. A core sampler is a tube 
designed to be pushed into fine sediments, most often in <1 m 
water. Peterson, Ponar, Ekman, and core samplers cannot be 
effectively used in swift currents over hard sediments, which 
limits their effectiveness for collecting stoneflies.
Collecting of Adults
As Resh and Unzicker (1975) and Lenat and Resh (2001) 
indicated, species-level identifications are crucial for 
higher-resolution analyses. For most stonefly species, for-
mal descriptions are based on male specimens. This often 
leaves females and especially nymphs undescribed (Stew-
art and Stark, 2002). In North America, Australia, and 
Europe, approximately 40% of the nymphs and a larger 
percentage of the females are known. These percentages 
drop substantially in other parts of the world. Moderate 
effort expended collecting adult stoneflies in addition to 
sampling for nymphs will improve a researcher’s chances 
of species-level identification and allow for association of 
environmental variables with species, not just a family- or 
genus-level identification.
Hand Picking
Adults are often collected from riparian vegetation and from 
the surface of emergent cobbles and boulders. They may 
also be obtained by turning loosely embedded cobbles and 
boulders alongthe shoreline. Winter-emerging stoneflies 
(Frison, 1929; Ross and Ricker, 1971)—including Capni-
idae, Taeniopterygidae, Nemouridae, and Leuctridae—may 
be collected from similar habitats, but they may also be 
found crossing snow or ice or retrieved from leaf packs just 
above the water line. Adults of stoneflies often crawl to the 
highest locations near streams. Look for them atop bridge 
abutments, the tops of signs, and on the windows of build-
ings near nymphal habitats.
Sweep and Aerial Nets
Sweep nets are used for sweeping insects out of riparian 
vegetation. We suggest you use a net with a 38-cm net ring, 
91-cm handle, and white muslin bag. Remove captured 
adults with forceps or an aspirator. Adults of the Nemouri-
dae, Leuctridae, Chloroperlidae, Perlodidae, and Peltoperli-
dae are often captured in large numbers by sweeping through 
streamside vegetation. Aerial nets are fitted with a long, fine 
mesh net on a longer handle than is found on a robust sweep 
net. Most stoneflies are generally erratic, clumsy fliers and 
spend little time in the air. However, adults will fly to the 
stream to drink or lay eggs (Poulton and Stewart, 1988). 
In addition, we have seen many adults near dusk fly from 
dark wooded areas into well-lighted forest openings. At this 
time, the more fragile, but rapidly swept aerial net may be 
used to good advantage.
Beating Sheet or Beating Net
The beating sheet or beating net is one of the most effec-
tive methods for collecting adult Plecoptera from riparian 
vegetation year round. It is often the only efficient means 
of sampling within dense, streamside vegetation. Using a 
beating sheet, adults of Capniidae, Nemouridae, Leuctridae, 
SECTION | VI Phylum Arthropoda944
Taeniopterygidae, Chloroperlidae, Peltoperlidae, and Per-
lodidae are often abundantly collected, especially in early 
morning hours during cooler temperatures. Beating sheets are 
especially useful for collecting winter-emerging stoneflies 
from riparian zones. The best method is to strike branches 
and associated vegetation with the handle of an aerial net or 
a stick using one hand while holding the sheet at one corner 
with the other (Figure 36.9). Specimens that drop onto the 
sheet are then gathered with forceps or an aspirator. Beating 
sheets are great for separating specimens from wet or dry leaf 
material and wood debris, especially winter-emerging species 
that avoid freezing by inhabiting organic debris just above 
the waterline. Place this material on the sheet and bounce the 
stoneflies free of the organic matter, tossing the excess debris 
off the sheet as you progress through it.
Light Traps
Use of ultraviolet light traps is a standard technique for gen-
eral insect surveys (Schauff, 2005) and is often the most 
efficient type of collecting for adults in the family Perli-
dae. Standard bucket traps may be used, but most stonefly 
researchers use one or two portable 15-W black light tubes 
either set atop a white bed sheet spread on the ground or on 
a sheet hung vertically between two trees. Power supplies 
can be 120 V AC or 12 V DC current. Set the ultraviolet 
wands at a 45° angle to the sheet to improve reflectance. 
Traps should be set up at least 30 min before dark and run 
for up to 2 h afterward. Few stoneflies come to light after 
this time. Lights may be unattended and set up along sev-
eral streams each evening to increase coverage of an area. 
Unattended lights may be turned on remotely using pho-
toelectric switches or mechanical timers. White trays with 
2 cm ethanol (concentration to purpose) should be left to 
capture stoneflies as they erratically fly about the light. It is 
important to attend some lights to collect adults of Perlidae 
and Perlodidae alive for later extrusion of the aedeagus, an 
internal reproductive organ of some stoneflies. The pattern 
of sclerites, spinules, and fine hair patches on the aedeagus 
is often diagnostic for a species (Figure 36.4(b)). In addi-
tion, live specimens may be taken back to the laboratory for 
behavioral observation, mating, and egg batch collection.
Malaise Traps
Malaise traps are useful for capturing many types of fly-
ing insects and have been employed in ecological studies 
of stoneflies (e.g., DeWalt and Stewart, 1995; Griffith et al., 
1998) and stonefly surveys (e.g., Parker et al., 2007). Vari-
ous commercial designs are available or can be constructed 
(e.g., Townes, 1962). In protected situations, these may be 
left for extended periods of time and repeatedly emptied 
of specimens to monitor the phenology of emergence. In 
long-term operations of Malaise traps, the choice of killing 
agent or preservative becomes important. A trap head with 
70–80% EtOH may be left for several days between visits.
Emergence Traps
Emergence traps sit atop water bodies and trap insects as 
they transform to the adult stage. There are two major advan-
tages to their use in ecological, biodiversity, and emergence 
phenology studies: The origin of the adults is indisputable, 
and if emptied daily, the number of days since emergence 
is known. Several designs for emergence traps are avail-
able (Merritt et al., 2008) and have been used extensively in 
sampling Plecoptera for ecological studies (e.g., Harper and 
Pilon, 1970; Masteller, 1983; Kerst and Anderson, 2006).
Miscellaneous Collecting Techniques: Pit Traps, Pan 
Traps, Sticky Traps, and Canopy Fogging
Pit traps (for the general methodology see Spence and 
Niemela, 1994) or pan traps (see Roulston et al., 2007) 
have been used to collect stonefly adults as they move from 
stream or lake edges into surrounding riparian zones after 
emergence or returning to the water to lay eggs. Secretive 
adults, especially Perlodidae, can be collected using these 
techniques. Generally a propylene glycol and 95% ethanol 
mixture is used as preservative in pit traps. These may set 
for a day to a week and then be emptied of specimens and 
replenished with preservative.
Pan traps are usually made of yellow (highly attractive 
to insects) plastic dishes with soapy water to help capture 
the specimens that alight in them. These should be set 
up near streams and examined daily. Specimens are then 
placed in ethanol for preservation. Sticky traps have also 
been used to collect adult stoneflies (e.g., Collier and Smith, 
1995). Generally, a flat surface is coated with Tanglefoot™ 
and placed on stakes or hung from trees or other substrates 
near the stream. Insects trapped in the Tanglefoot™ may 
be removed from the trap with ethyl acetate or methyl 
FIGURE 36.9 Plecopterologists often use a beating sheet to capture adult 
stoneflies. Here, Dr. Richard W. Baumann checks his sheet for stoneflies.
945Chapter | 36 Order Plecoptera
chloroform. Specimens then should be washed in Cello-
solve™ before storage in ethanol.
Sometimes stonefly adults in the tropics occupy the high 
canopy of trees near streams and are much more abundant 
than one might suspect using traditional sampling devices. 
Shepard and Baumann (2011) used canopy fogging with 
an insecticide to collect stoneflies in the forests of south-
ern Chile, unexpectedly yielding large numbers of Grypop-
terygidae and Notonemouridae adults from the canopy. A 
standard agricultural fogger is used with diesel fuel mixed 
with a pyrethroid insecticide. Trees are fogged for a specific 
time. Individual specimens fall onto sheets placed around 
the base of the trees, and are then preserved.
Preservation Techniques and Labeling
Preservation Techniques
Preservatives such as ethanol will protect a specimen from 
bacterial or fungal degradation while at the same time pro-
tecting color patterns and external morphology. Fixatives 
such as formalin have these same properties but are better 
at protecting the integrity of soft tissue because the fixa-
tive forms cross-bridges in protein and DNA molecules. It is 
important to understand this difference because the choice 
of a preservative or a fixative depends on the objectiveof 
the study being conducted.
Under most circumstances, ethanol is the preservative 
of choice for stoneflies. In concentrations ranging from 
70% to 80%, it preserves color well and produces supple 
specimens. In addition, it is pleasant to use and stores 
safely, and its disposal is much less problematic than for 
other preservatives/fixatives. A few specimens may be live 
sorted into 70–80% ethanol and yield well-preserved color 
patterns without appreciable shrinkage of the body. These 
specimens will be suitable for a wide range of objectives 
including identification and enumeration, life history mea-
surements, and taxonomic description, as teaching collec-
tions or for long-term museum storage. However, when 
preserving large numbers of individuals or bulk samples in 
the field, so much water is associated with the samples that 
70–80% ethanol would become diluted to levels that would 
not preserve specimen integrity. In these cases, it is wise to 
preserve with 95% or 100% ethanol applied directly to the 
sample. If bulk samples are large (>1 L volume), a second 
rinse with 95% ethanol may be required within a few hours 
of the initial preservation. Because ethanol takes time to dif-
fuse into animal tissue, it is best to store samples under cool 
or cold conditions while in the field and under refrigeration 
in the laboratory. Sorting of these samples and long-term 
storage may then take place in 70–80% ethanol. If traveling 
abroad, ethanol may be difficult to obtain. Try a pharmacy. 
Even here, you may find the ethanol to be denatured with 
additives to make it unpalatable. Denatured ethanol is fine 
for general collecting, but the additives may render it unfit 
for preserving samples for RNA/DNA extraction. Some-
times the only source of ethanol is drinking spirits. If this 
is your last resort, obtain the highest possible proof (per-
cent concentration is half of proof listed on the container). 
If you are left without ethanol, adults and nymphs may be 
pinned and dried until they can be relaxed and examined in 
the laboratory.
Isopropanol is a poor preservative because it does not 
diffuse into animal tissue quickly, often leaving specimens 
transparent, flaccid, and fragmented. Its major advantage is 
its low cost. An inexpensive alternative to ethanol is 10% 
buffered formalin (10% formaldehyde to 90% water, with 
stock being 37–40% formaldehyde). This will actually pro-
duce a concentration of 3–4%. Often, formalin is acidic 
and may need to be buffered with a thin layer of sodium 
bicarbonate or household borax on the bottom of the con-
tainer. In addition, Kahle’s fluid is a good general fixative; 
it is composed of 11% formalin (the 3–4% concentration 
described previously) plus 28% ethanol (the 95% concen-
tration) plus 2% glacial acetic acid plus 59% water. The 
water may be added in the field, thereby reducing the vol-
ume of fixative taken into the field (Edmunds et al., 1976). 
Fixatives that contain formaldehyde are far less pleasant to 
use and require more careful storage, ventilation, and dis-
posal of waste. Note: It should not be disposed of down the 
drain! These fixatives are not suitable for permanent stor-
age; specimens may be rinsed in water for several hours and 
then transferred to 70–80% ethanol. A useful resource for 
preservation/fixation of aquatic invertebrates is contained in 
Pennak (1989).
Storage containers are another important consideration. 
The right container will store bulk samples for decades or 
individual specimens for a century or more. For collection 
and storage in the field, Nalgene™ plastic containers come 
in a wide variety of sizes and are hard to beat. Whirlpacs™ 
may be used for temporary storage but must be stored in 
larger, sealed plastic bags or jugs for to prevent leakage and 
evaporation. In the laboratory, avoid using vials and jars 
with flat paper or plastic liners in lids because these allow 
rapid evaporation of the preservative. Natural history muse-
ums often use glass, patent-lip vials of 3 or 4 dram size. 
These vials are sealed with rubber or neoprene stoppers that 
swell past a narrowed shoulder of the vial, creating a tight 
seal. A thin wire or forceps allows release of pressure as the 
stopper is pushed into the vial. Avoid natural cork stoppers 
because they allow rapid evaporation of the preservative. 
Screw-cap vials with cone-shaped plastic inserts are good 
alternatives to patent-lip vials; their only detraction is that 
the openings for smaller vials are not wide enough to pass 
standard forceps to the bottom. Glass shell vials with plastic 
snap caps may also be used effectively, but with repeated use 
the plastic threads of the snap caps are damaged and evapo-
ration of preservative may occur. Clear plastic cryovials of 
SECTION | VI Phylum Arthropoda946
2 and 8 mL volume with lids that have neoprene O-rings are 
now becoming widely used (one supplier is Sarstedt). They 
have several advantages over glass vials: reductions in cost, 
weight, space, and ethanol use, and shatter resistance, and 
they may be stored in freezers to −80 °C without becoming 
fragile or losing their seal.
If DNA is to be extracted from the specimens, it is 
imperative that consultation with a molecular biologist takes 
place so that collection and storage protocols meet the study 
objections. Generally speaking, collection and storage in 
95–100% ethanol are sufficient for Sanger type sequencing 
of mitochondrial (Sweeney et al., 2011) and nuclear DNA. 
Cold storage of specimens is preferred (−20 °C or colder).
Labeling Specimens
Regardless of the method used for collecting, proper label-
ing of all samples should be routine. The minimum label 
data needed to identify a field sample includes at least a field 
notebook number (e.g., initials of collector–year–consecu-
tive number) or sample code, the water body sampled, and 
a date. This combination of information provides a check 
against any one part of the label being recorded incorrectly. 
The field notebook number serves as the key to a hardcopy 
notebook entry or electronic record that includes the fol-
lowing: country, state, county, distance (km), and direction 
(N, NE, E, SE, etc.) from a small nearby town or other rec-
ognized landmark, road crossing (if applicable), geographic 
coordinates (the coordinate system choice is the reader’s, 
but latitude/longitude in decimal degrees to four or five 
significant figures is commonly used), date of collection 
(consistency in format is important, recording months as 
Roman numerals and a four-digit year will allow for unam-
biguous dates), collector(s), sample protocol(s), and reach-
level habitat and water physical-chemical measures. Other 
data may also be present. Use carbon-based inks or pen-
cil on white, high–cotton rag content paper for temporary 
labels and place it inside the container. A secondary label 
may be affixed externally to aid in the sorting of samples. 
Of course, the reference standard is to provide a temporary 
label containing the entire suite of locality information as 
described previously, but under certain conditions we err 
on the side of collecting several more samples in a day in 
contrast to fully labeling each individual sample.
Once sorting of specimens into individual vials has 
been accomplished, high-quality interior labels are pro-
vided. These include the locality and the determination or 
identification labels. Although it is easy to recognize the 
value of the locality label, the determination label is often 
overlooked or done in shorthand. Key features of the deter-
mination label include the binomial (may be left at family 
or genus level (e.g., Taeniopteryx sp.)), the author of the 
species (“Ricker and Ross”), the stages and numbers of 
individuals, the determiner, and the determination year. The 
author is provided just in case there is a homonym (different 
taxa given the same name) involved, in which case the 
author helps to distinguish which taxon is being presented.The determiner name helps reveal the trustworthiness of the 
identification. Not all taxonomists are created equal, and the 
opinions of some are trusted more than others. The year of 
determination helps a reader assess what concept of the spe-
cies (current or obsolete) was used when the specimen was 
identified.
Labels may be handwritten if only a few specimens are 
involved. However, the number of labels needed for a single 
site/date event might be large, and a word processor, spread-
sheet, or database is more efficient at producing duplicate, 
high-quality, appropriately sized (depends on the container) 
labels. This not only relieves the researcher of the tedium of 
hand printing labels, but reduces errors in interpretation by 
others viewing them in the future. Most moderately priced 
laser printers can produce long-term labels if used in con-
junction with high–cotton rag content papers.
Rearing of Stoneflies
Rearing immatures to the adult stage is often the only means 
of determining what species a nymph represents. Because 
of this, stonefly researchers rear many nymphs to adulthood 
to associate life stages and to obtain species-level identifica-
tions. Shepardson (1968) and Merritt et al. (2008) described 
many of the rearing methods available. We discuss some 
methods that work for us. Nymphs are carefully collected 
for rearing from their natal habitat. It is wise to segregate 
detritivores (slowly moving, drab forms) from predators 
(rapidly moving, often highly patterned). Styrofoam cups 
with lids work well as temporary rearing chambers. On 
the lid, identify each cup with stream name, date, and field 
notebook number. Many cups may be transported in an ice 
chest by neatly stacking them on cardboard spacers. Gen-
eral rules of thumb are to keep the water depth in the cup 
shallow (2–4 cm) to improve diffusion of oxygen and not 
to place more than 15–20 detritivores or five predators in a 
cup. Nymphs do not tolerate rapid increases in temperature, 
but they can withstand substantial cooling so long as it is 
gradual. Ice in plastic bags may be placed next to the cups, 
enough to lower the temperature in the cooler and cups to 
5–10 °C. Nymphs will settle quickly onto small leaf scraps, 
wood, or gravel provided for them.
Detritivores are easiest to rear, mainly because they 
have a high surface area to body size ratio, are often found 
in decaying leaves and wood where oxygen is already 
depressed, and can be fed conditioned leaves and wood 
from their natal habitat. These stoneflies often may be 
reared without current, as long as the water temperature is 
cold. In fact, an ice chest and cups are all that are needed in 
many cases for rearing detritivores. Collect the adults and 
exuviae from these cups daily and preserve them in separate 
vials, recording the field notebook number, stream name, 
and date of emergence for each vial.
947Chapter | 36 Order Plecoptera
Predators are more difficult to rear. They are often larger 
than detritivores, are active, live in the most highly oxygen-
ated habitats, and require live food. Here, water flow and 
low temperatures are extremely important for successful 
rearing. The simplest rearing chamber is the stream itself. 
Creation of rearing bags from window screening works well 
(Frison, 1935). Make a pillow of the screening, leaving one 
end open. Fill the bottom with gravel, leaves, and sticks. 
This substrate will hide the nymphs and provide access to 
prey items. Fold over the top and clip it tightly, but leave the 
top 4–6 cm above the water level so that nymphs will have 
a place to crawl out of the water, and adults a place to stay 
dry. These bags should be checked daily, removing adults 
and exuviae. For summer-emerging predators, jars with 
stream water and an aerator will often suffice for rearing. 
These can be reared at air-conditioned room temperature, 
but be sure to provide some substrates above the water sur-
face for transformation of the nymph to the adult.
A commercial rearing chamber such as the Living 
Stream™, produced by Frigid Units, will provide both water 
current and temperature control. Stonefly researchers often 
float Styrofoam sheets atop the water and sink cups through it 
with screened windows for current flow. Cup lids or clear Petri 
dishes may be used to cover the cups. Reared male and female 
individuals can be transferred to containers with damp paper 
towels and allowed to mate in a cool area (18–21 °C). Often 
after 1–3 days, females may produce egg masses for study. 
Rearing nymphs individually will allow for absolute associa-
tion of the nymph characters represented by the shed exuvium 
with the adult and provide virgin females for drumming study.
ACKNOWLEDGEMENTS
The authors thank the efforts of Dr José Manuel Tierno de Figueroa 
of the Universidad de Granada, Spain for providing information on 
European species whose drumming has been recorded. We also thank 
Marilyn Beckman of the Illinois Natural History Survey for conduct-
ing queries on Plecoptera Species File that helped to differentiate the 
number of valid extant species from the fossil species in the database.
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	36 - Order Plecoptera
	Introduction
	Overview of the Insect Order Plecoptera
	Phylogeny and Biogeography
	General Biology
	External Anatomy
	Immature Stoneflies
	Adult Stoneflies
	Eggs
	Life History
	General Ecology and Behavior
	Macro- and Micro-habitat Usage
	Physiological Constraints on Distribution and Survival
	Feeding Behavior
	Parasites and Symbionts of Stoneflies
	Conservation of Stoneflies
	Behavior
	Vibrational Communication (Drumming)
	Diel Periodicity of Adults
	Collecting and Rearing Stoneflies
	Collecting Nymphs and Adults
	Collecting Nymphs
	Dip Net
	Kick Screen
	Hess, Surber, Drift Net, Peterson Grab, Ponar, Stovepipe Samplers
	Collecting of Adults
	Hand Picking
	Sweep and Aerial Nets
	Beating Sheet or Beating Net
	Light Traps
	Malaise Traps
	Emergence Traps
	Miscellaneous Collecting Techniques: Pit Traps, Pan Traps, Sticky Traps, and Canopy Fogging
	Preservation Techniques and Labeling
	Preservation Techniques
	Labeling Specimens
	Rearing of Stoneflies
	Acknowledgements
	References