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From Baxter, C.V., Kennedy, T.A., Miller, S.W., Muehlbauer, J.D., Smock, L.A., 2017.Macroinvertebrate Drift, Adult Insect Emergence and Oviposition. In: Hauer, F.R.,

Lamberti, G.A. (Eds.), Methods in Stream Ecology: Volume 1: Ecosystem Structure.Elsevier, Academic Press, pp. 435–456.

ISBN: 9780124165588Copyright © 2017 Elsevier Inc. All rights reserved.

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Chapter 21

Macroinvertebrate Drift, Adult InsectEmergence and Oviposition

Colden V. Baxter1, Theodore A. Kennedy2, Scott W. Miller3, Jeffrey D. Muehlbauer2 and Leonard A. Smock4

1Department of Biological Sciences, Idaho State University; 2U.S. Geological Survey, Southwest Biological Science Center, Grand Canyon

Monitoring and Research Center; 3National Aquatic Monitoring Center, Department of Watershed Sciences, Utah State University; 4Department of

Biology, Virginia Commonwealth University

21.1 INTRODUCTION

Aquatic invertebrates exhibit movements and behaviors that are ecologically important, not only because these processesare critical to the life cycles and population dynamics of individual species, but because they mediate key roles played bysuch invertebrates in the intricate web of life associated with streams, rivers, and their surrounding watersheds. Forinstance, although most stream invertebrates are benthic, they can also be found drifting with the current. This process ofdrift is essential to dispersal and colonization that help maintain populations of these animals (Brittain and Eikeland, 1988;Hart and Finelli, 1999; Downes and Lancaster, 2010). Similarly, of the many stream invertebrates that are insects, nearly alltransition to becoming winged, air-breathing adults via emergence and, after a period in this form, oviposit (i.e., lay theireggs) back in streams. These are key stages in their life cycle (Huryn and Wallace, 2000; Merritt et al., 2008), but alsoprocesses by which they become vulnerable to aquatic predators like fishes and are linked to terrestrial food webs, oftenserving as prey for animals like birds, bats, spiders and lizards (Baxter et al., 2005; Sabo and Hoekman, 2015). Therefore,stream ecologists study these processes of drift, emergence, and oviposition to address questions at scales from individualinvertebrates and populations of a species, to food webs and ecosystem processes in streams, rivers and their surroundinglandscapes. Increasingly, conditions necessary to support insects not just at the larval stage, but throughout their life cycles(e.g., egg, larvae, pupae, adult), are understood to be important to species’ management and recovery efforts (Peckarskyet al., 2000). These other life stages are traditionally understudied in stream ecology, but are nonetheless critical to thepersistence of aquatic invertebrate populations.

This chapter introduces key concepts and sampling methods concerning drift of benthic stream invertebrates, and theemergence, and oviposition behavior of lotic insects. In the previous edition of this text, Smock (2006) outlined methodsassociated with drift and emergence as a means of dispersal, and with an emphasis on techniques traditionally used in thecontext of invertebrate life history and population studies. Naturally, these methods, and the overarching topic of dispersal,continue to be of importance to the practicing stream ecologist. Here, however, we focus on these processes to includedescription of the growing array of methods for their quantification being applied in food web and ecosystem studies.Finally, we extend our treatment to encompass oviposition, a phenomenon closely linked to both emergence and drift, thedetails of which are receiving fresh attention as new studies (e.g., Lancaster et al., 2010a; Encalada and Peckarsky, 2012)have raised awareness of its importance to the ecology of stream insects and, with its potential consequence for thecommunities of organisms to whom they are linked (Kennedy et al., 2016).

21.1.1 Drift of Stream Invertebrates

Although most invertebrates that occur in streams and rivers are benthic, a net placed in the water column often will collectmany individuals. These organisms are drifting, an activity whereby they enter the water column and are transported

Methods in Stream Ecology. http://dx.doi.org/10.1016/B978-0-12-416558-8.00021-4Copyright © 2017 Elsevier Inc. All rights reserved.

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Methods in Stream Ecology, 3, 2017, 435e456


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downstream by the current. Entry into the water column by invertebrates can be broadly classified as accidental (passive) orbehavioral (active). Drift is one of the most important mechanisms for the dispersal to and colonization of downstreamhabitats by lotic invertebrates, and is also a stage at which invertebrates are particularly vulnerable as prey to fish. Becausedrift is such a fundamental process in streams and rivers, it is the most widely studied movement of benthic invertebrates(see reviews and syntheses by Waters, 1972; Müller, 1974; Brittain and Eikeland, 1988; Rader, 1997; Allan and Castillo,2007; Naman et al., 2016).

Early observations of diel periodicity in drift concentrations (# m�3; sometimes called drift density) motivated decadesof research by stream ecologists into drift dynamics. These studies demonstrated that the majority of species actively driftin maximum numbers sometime during the night. Changes in ambient light intensity, although not the ultimate cause foractive drift, serve as the trigger or proximate cause for this behavioral drift (Naman et al., 2016). Factors that can influenceactive drift include dispersal to more favorable habitats (Walton et al., 1977; Hershey et al., 1993; Koetsier et al., 1996;Siler et al., 2001), escape from predators and competitive interactions (Flecker, 1992; McIntosh et al., 2002), andmovements associated with key life-history events such as egg-hatching, pupation, and emergence (Otto, 1976; Kruegerand Cook, 1981; Ernst and Stewart, 1985).

Passive drift occurs when invertebrates are accidentally dislodged from the benthos. Changes in discharge are animportant variable that can influence the rate of passive drift (Gibbins et al., 2007a; Miller and Judson, 2014; Kennedyet al., 2014). Catastrophic drift represents an extreme case of passive drift and occurs when large numbers of invertebratesare physically removed from the channel bed and a high proportion of the available invertebrates become entrained in thedrift. If the conditions for initiating catastrophic drift persist for prolonged periods of time, benthic populations can bedepleted, reducing benthic invertebrate populations over long time-scales (i.e., months). The lowest threshold for cata-strophic drift occurs when shear stress is sufficient to entrain sand and “sand blast” invertebrates off the bed. At higherlevels of shear stress, large numbers of invertebrates may be physically removed from the bed, and at even higher shearstress the cobble substrates on which invertebrates live will be mobilized. Thus, catastrophic invertebrate drift can beinitiated at discharges that may or may not drive major changes in streambed or channel form (Gibbins, 2007a,b; Namanet al., 2016). Chemical pollution and abrupt changes in water quality have also been shown to precipitate catastrophic drift(Brittain and Eikeland, 1988; Wallace et al., 1989).

While invertebrates stand to benefit from actively drifting, this movement also poses significant costs, particularlyincreased risk of predation by drift-feeding fish. Minimizing predation risk by drift-feeding fish underlies the diel periodicity inactive drift exhibited by invertebrates (Flecker, 1992). For instance, the propensity of invertebrates to exhibit nocturnal driftincreases as the risk of predation by the visual-feeding fish increases, whereas drift is usually aperiodic in fishless streams(Flecker, 1992; Forrester, 1994; March et al., 1998). Because drift-feeding fish are visual feeders, their ability to detect andcapture prey increases with the size of invertebrates. Thus, predation risk also can affect the size distribution of drifting in-vertebrates, with larger individuals that are more vulnerable to predation being more prone to drift at night (Allan, 1978, 1984).

Invertebrate drift can constitute the primary prey source for a diverse guild of fishes (Grossman, 2014). Thus, in manycontexts, drift availability is a primary determinant of the capacity of streams and habitats to support fish populations(Rosenfeld et al., 2014). For instance, field studies have directly linked the magnitude of drift flux to fish abundance,growth, and survival (Fausch et al., 1991; Keeley, 2001; Rosenfeld et al., 2005; Naman et al., 2016). Linked drift-foragingbioenergetics models explicitly consider river hydraulics, invertebrate drift biomass, and the mechanics of drift-feeding todescribe foraging dynamics and how food availability translates into fish growth (Hughes and Dill, 1990; Hayes et al.,2000). These models are increasingly being used to estimate the capacity for streams to support drift-feeding fishes(Rosenfeld et al., 2014). These models can also be used to evaluate how fish growth rates will be influenced by alternativeprey and temperature scenarios that would be difficult to test in a field setting (Dodrill et al., 2016). Accurate estimates ofdrift biomass are a critical input parameter of these models.

Methods are provided in this chapter (Fig. 21.1) to: (1) quantify the rate of net clogging, which informs the design ofdrift studies; (2) quantify variation in drift among habitat types in wadeable streams; (3) quantify variation in drift betweenday and night; and (4) quantify the vertical distribution of drift in large rivers.

21.1.2 Emergence of Adult Stream Insects

Sampling of stream insects has historically focused on their immature stages (e.g., see Chapter 20); however, nearly all ofthese insects metamorphose into adults that emerge, mate, and oviposit, so methods for quantifying emergence andoccurrence of adults in terrestrial habitats are important. Studies of insect emergence provide not only an understanding ofinsect life cycles, dispersal, colonization, and population ecology, but also food web and ecosystem processes that linkwater and land. Further, measures of emergence may reveal responses of insects to stressors (e.g., pollutants) or community

436 SECTION | C Community Interactions



interactions different from those detected through measures of benthic larvae alone (e.g., Walters et al., 2008; Schmidtet al., 2013; Wesner, 2016). In addition, regardless of the ecological questions under investigation, collection of adults canbe necessary to obtain species-level identifications, owing to the incomplete taxonomy of immature forms for many groupsof aquatic insects.

Emergence is a process that can be highly heterogeneous in time, which presents challenges to its measurement. Intemperate zones, emergence of individual taxa is typically seasonal and may be highly synchronous over a few days to afew months (Sweeney and Vannote, 1982), whereas it can be more continuous in tropical regions (Corbet, 1964) or mayexhibit variation with wetdry seasons (Suhaila et al., 2012). Emergence by the overall assemblage of insects in temperatestreams often peaks in early summer and declines sharply by late summer (Sweeney and Vannote, 1982; Malison andBaxter, 2010), but can also provide a low level of flux to riparian zones during autumn through early spring (Jackson and

(A) (B)

(C) (D)(D)

FIGURE 21.1 Approaches to sampling macroinvertebrate drift. (A) Traditional rectangular drift net equipped with recording flow meter. The secondflow meter outside the net is used to record ambient stream water velocity, needed for estimating filtration efficiency of drift nets (see Basic Method 1). (B)Large river invertebrate drift-sampling equipment. The whole apparatus is raised and lowered through the water column using a hand-power winch andchain attached to the end of the cable, and a lead weight attached to the bottom of the chain keeps the circular plankton net from moving downstream. Thissetup is from filtration-efficiency experiments, and the black cord running from the flow meters to the boat transmits velocity readings to on-boardcomputers for visualization; during routine drift sampling, stand-alone flow meters are used. (C) Preparing for drift sampling in the Colorado River,Grand Canyon, AZ, USA. The hand-powered winch (A-reel) is mounted to a chest-height platform, the winch cable is extended out beyond the bow of theboat by means of a crane and pulley and net is lowered to desired depth using the depth gauge on the left side of the winch. (D) Quantifying invertebratedrift on the Green River downstream of Flaming Gorge Dam, Utah, USA.

Macroinvertebrate Drift, Adult Insect Emergence and Oviposition Chapter | 21 437



Fisher, 1986; Nakano and Murakami, 2001), or may depart from these patterns altogether as has been observed in someMediterranean climate streams (Rundio and Lindley, 2012). Moreover, diel patterns in emergence and flight behavior ofadults occur (e.g., Jackson, 1988), which can influence effectiveness of the trapping methods. Large pulses of emergencecan be easily missed, so estimating total emergence flux, which is often needed for food web or ecosystem studies (e.g., asindex of secondary production; see Chapter 25), requires frequent and prolonged sampling (e.g., Jackson and Fisher, 1986;Nakano and Murakami, 2001; Malison and Baxter, 2010; Rundio and Lindley, 2012; Kennedy et al., 2016).

The flux of emergent insects also varies spatially, with attendant implications for sampling. Spatial patterns inemergence may be related to river and riparian habitat characteristics, as well as distribution of larval stages and behaviorduring metamorphosis. For instance, emergence flux may be greater from habitats that retain aquatic insects emergingthrough the water column (e.g., pools; Iwata, 2007), or provide insects predation refuge (e.g., floating algal mats; Poweret al., 2004), and could be amplified by habitat complexity such as sinuosity (Iwata et al., 2003), thermal heterogeneity(Uno, 2016) or increased surface area of the landewater interface (Sabo and Hagen, 2012). Emergence flux can even varyin relation to the availability and quality of oviposition substrates available during the egg-laying in the prior generation(Encalada and Peckarsky, 2012; Kennedy et al., 2016). Another source of spatial variation in emergence arises from thedifferent approaches that taxa use in their transition to land. For instance, most Diptera and Trichoptera actually emergethrough the water column, while taxa such as Plecoptera and some Ephemeroptera typically emerge from stream banks(Merritt et al., 2008). Thus, different methods of sampling may capture adults of different groups with varying efficiency(Malison et al., 2010).

The total flux of emergence from a given habitat or stream in turn influences the magnitude of flux into adjacentterrestrial habitats. Following emergence, the number of adult insects penetrating riparian zones typically declines withdistance from the stream edge (Jackson and Resh, 1989; Sanzone et al., 2003; Power et al., 2004; Briers et al., 2005). Arecent meta-analysis (Muehlbauer et al., 2014) showed this pattern followed a negative power function, though therelationship was mediated by stream size and 10% of flux was predicted to occur >500 m from the water’s edge, sug-gesting dispersal of some adults may extend the ecological “signature” of streams throughout watersheds, particularlywhen the complexity and extent of stream networks are considered (Sabo and Hagen, 2012).

Insect traits such as dispersal behavior or preferences for oviposition sites, along with riparian conditions, may alsoinfluence the distribution of postemergent adults, with additional implications for sampling methods. Many adult insectsundertake dispersal flights that can result in mating and/or laying of eggs far from the site of emergence, facilitatingcolonization of new habitats, genetically linking populations (e.g., Kovats et al., 1996; Turner and Williams, 2000;Winterbourn and Crowe, 2001; Macneale et al., 2005), and connecting food webs in mainstem rivers and tributaries (Unoand Power, 2015). Indeed, a “colonization cycle” has been hypothesized, the central component of which is that the flightof females prior to ovipositing should be primarily directed upstream, thereby compensating for the predominatelydownstream drift of the immature individuals living in the water (Müller, 1982). The results from field studies designed totest this colonization cycle, however, are equivocal (Hershey et al., 1993; Jones and Resh, 1988; Winterbourn and Crowe,2001; Macneale et al., 2004), with theoretical studies having resolved this “drift paradox” by demonstrating that randombenthic dispersal (i.e., upstream crawling), high benthic production, and density-dependent survival are sufficient tocompensate for downstream drift and allow for benthic population persistence in lotic environments (Anholt, 1995;Humphries and Ruxton, 2002). Dispersal distances may also be taxon-specific: Ephemeroptera tend to remain very nearstreams whereas Trichoptera and Chironomidae often travel farther inland, for example (Muehlbauer et al., 2014).Moreover, adult insect behavior and distribution may be mediated by riparian habitat characteristics (e.g., vegetation,microclimate; Collier and Smith, 2000; Briers et al., 2003; Petersen et al., 2004) and differences between the sexes(e.g., Petersen et al., 1999). Any of these factors may require consideration when choosing the type of trap or net, insampling design, and in interpretation of study findings.

Methods are provided in this chapter to introduce methodology for sampling emerging adults and estimating emergenceflux (Fig. 21.2) to (1) examine the differences in emergence that occur both spatially and temporally within a stream, (2)quantify dispersal distances of adult insects laterally from a stream, and (3) quantify emergence or postemergent adults inorder to compare different habitats in terms of insect prey availability to terrestrial insectivores.

21.1.3 Oviposition by Stream Insects

Much like emergence, oviposition by stream insects is understudied relative to larval life stages. This is likely due pre-dominately to practical considerations: insect oviposition often occurs only during specific, narrow windows of time withina day and within a year, and the time insects spend in the egg stage is commonly short compared to other life stages(Jackson and Sweeney, 1995). However, the egg-life stage is just as important to the survival of insects as any other stage,




and consideration of oviposition and egg development can reveal important life history bottlenecks that influence thepresence and persistence of insect groups within a habitat or ecosystem (Kennedy et al., 2016).

Insect oviposition is necessarily tied to the presence of emergent adults, which, as noted earlier, are often present onlyfor short periods of time (Sweeney and Vannote, 1982). Because the egg-laying stage occupies just a portion of this alreadylimited amount of time available to observe emergent adults, studying the oviposition can be challenging. For taxa such asmany Ephemeroptera and Chironomidae that can be particularly short-lived, mating swarms of adults can be a visual cuethat oviposition is about to commence (Sweeney and Vannote, 1982; Gray, 1993). Additionally, emergent adults tend to bemost active in crepuscular periods when the visual contrast between water and land surfaces is greatest (Bernáth et al.,2004). For a given taxon, the presence of eggs or spent adults in or around the water surface provides another line ofevidence that oviposition is ongoing or recently concluded, which allows inference about the timing, seasonality, andduration of egg-laying.

Stream insects exhibit a range of oviposition behaviors, from generalized behaviors such as broad-cast spawning onthe water surface, to more specialized behaviors such as cementing eggs on emergent rocks or even females ovipositingfertilized eggs onto the back of the male with whom they recently copulated (Vieira et al., 2006). However, little isknown about oviposition for most stream insect species, with a recent synoptic database of traits for >14,000 NorthAmerican aquatic invertebrates only yielding a description of oviposition behaviors for 957 taxa (Vieira et al., 2006).Among mayflies, for instance, Encalada and Peckarsky (2007) identified five distinctive behaviors for 17 mayfly speciesin a single Rocky Mountain watershed, ranging from dropping eggs into the water from midair, to crawling down

(A) (B)

(D)(C)

FIGURE 21.2 Approaches to measuring emergence of adult stream insects, including a floating trap (A), a trap for capturing taxa that emerge alongstream banks (B), and methods for sampling postemergent insects in the terrestrial environment, including a sticky trap (C) and a light trap (D). Photocredits: (A & B) C.V. Baxter; (C) E. Kortenhoeven; (D) D. Herasimtschuk.




emergent rocks and depositing eggs under the water surface. At another extreme, at least one mayfly species has beenobserved to release parthenogenic eggs via bursting of the abdomen while still in the subimago life stage (Funk et al.,2008). In some cases, eggs are not even laid in water: many blackflies (Simuliidae), dragonflies (Odonata), and othertaxa lay eggs either on or within vegetation along the shoreline (Adler et al., 2004; Corbet, 1999), whereas the males ofmany giant water bug (Belostomatidae) species rear eggs on their backs (Smith, 1976). In general, however, results frompast studies indicate that egg masses are disproportionately found near the stream margins and are laid on emergentmineral or wood substrates in moderate water velocities (Peckarsky et al., 2000; Hoffmann and Resh, 2003; Lancasteret al., 2010a,b). Rock size and flow velocity were the only consistently important physical variables affecting thedistribution of egg masses for 17 taxa considered in one study, for instance (Reich and Downes, 2003), and populationdensities of both Baetis mayflies and Ecnomus caddisflies have been affected by experimental manipulations ofemergent rock and log substrates (Encalada and Peckarsky, 2012; Macqueen and Downes, 2015). Indeed, a synthesis ofoviposition behaviors for those species where behavior is known indicates that most cement eggs to underwater sub-strates (Statzner and Beche, 2010).

As with oviposition behaviors, the characteristics of insect eggs are generally not well-described for most lotic insectspecies (Vieira et al., 2006). Nonetheless, for those taxa that have been studied, there is impressive variability in eggcharacteristics (Fig. 21.3). Egg fecundity, for instance, can vary across orders of magnitude, from eggs distributed bybroadcast spawners as singlets, to egg masses with eggs in the thousands that may represent contributions from severalfemales (e.g., Hoffman and Resh, 2003). Similarly, some species diapause for months while in the egg stage, whereasothers hatch instantly upon being laid or are even ovoviviparous (e.g., some Baetidae and Capniidae, Lancaster andDownes, 2013). In general, egg masses tend to be on the scale of a few millimeters in size, and may be gelatinous andbulbous, or arranged in a flat mat or a stringy line (Fig. 21.3). This dimorphism may confer fitness benefits to speciesadapted for certain habitats and oviposition sites; the flat arrangement of Hydropsyche caddisfly eggs may allow them topersist in fast-water habitats, for example, whereas the thickly gelatinous egg masses laid by Ethochorema caddisflies mayrender them less vulnerable to predation (Bovill et al., 2015). The shape and other characteristics of these egg masses varywithin insect families or even genera (Lancaster and Downes, 2013); however, making species-specific egg mass iden-tification possible.

Methods are provided in this chapter to: (1) observe insect oviposition and search for insect eggs; (2) quantify eggdensities and oviposition-site selection; and (3) rear insect eggs and quantify hatch success.

21.2 GENERAL DESIGN

21.2.1 Site Selection

Naturally, the selection of sites for conducting studies of drift, emergence, or oviposition will depend upon the questionsunder investigation, and the choice of site can have important consequences for the types of methods that are mostappropriate or the adaptations to techniques that may be required. Below we provide some guidance along these lines, butthere is no single recipe for selecting the study sites. With respect to drift, for instance, investigations into its diel peri-odicity might focus on collecting replicate samples in habitat types where drift concentrations are high (e.g., downstream ofriffles), investigations into invertebrate population dynamics or habitat use by fishes might require drift measurements froma range of habitats within a stream reach, whereas investigations into the role of benthic density in controlling drift mightbe done across habitats and among streams. However, there are limits to the types of habitats where drift can be sampled.Specifically, drift nets become difficult to maintain or inefficient for measuring drift accurately under conditions of veryfast (>1e2 m/s) or very slow water velocity (<0.1 m/s). Nets can be positioned at any location within the channel, butplacement in midstream at the downstream end of a riffle usually is most productive in terms of the number of species andindividuals captured. Drift-sampling methods for wadeable streams are well established, and the vast majority of driftstudies have been conducted in these settings. Drift can also easily be sampled from a boat in large rivers by adaptingplankton sampling techniques originally developed by oceanographers (Kennedy et al., 2014). Drift sampling in largerivers is often easier and safer than traditional benthic invertebrate sampling, which are both important considerations inany study design. Further, because drift samples are spatially integrated, the variance in individual drift samples tends to bemuch lower than for benthic samples (Allan and Russek, 1985). Thus, regular measurement of invertebrate drift can be auseful and low-cost alternative or complement to traditional benthic sampling, such as in long-term invertebrate monitoringprograms whose purpose is to inform fish population trends.

The best tools to measure adult insect emergence and oviposition also depend upon study questions and context. Forinstance, the choice of emergence trap style, positioning (e.g., pools, riffles, bankside) within a stream reach (or reaches),




(A) (B)

(C) (D)

(E) (F)

FIGURE 21.3 Morphological diversity of aquatic macroinvertebrate egg masses. Included are photos of (A) Baetis spp., (B) Simulium vittatum,(C) Brachycentrus occidentalis, (D) Hydropsyche occidentalis, and (E & F) Chironomidae. Photos credit: M. Schroer.




and numbers of replicate samples may depend upon whether the study focus is description of insect life cycles anddispersal, or estimating these fluxes as part of food web investigations (see Davies, 1984; Malison et al., 2010). Trapsmay need to be adapted to sample particular habitats (organic debris jams, algal mats, floodplain wetlands; e.g., Whilesand Goldowitz, 2001; Power et al., 2004), or modified in the case of oviposition studies to capture insects as they returnto the water to lay eggs (Encalada and Peckarsky, 2007). Generally, as is the case for drift, far fewer studies have beenconducted on emergence or oviposition in larger rivers compared to small streams. As with any process quantified via anet or trap device that samples a limited habitat area, any increase in stream size without a change in the amount of areasampled is likely to result in a less representative quantification of the process. Thus, representative sampling ofemergence flux associated with large rivers may require larger traps, more traps, or the use of active techniques such aslight traps, which attract and integrate samples of adult insects over larger habitat areas (e.g., Kennedy et al., 2016).Additionally, trap designs that work in wadeable streams may become dangerous to deploy or are capsized in morepowerful currents of larger rivers. In some study contexts or where more spatially continuous sampling may be desired,remote sensing may prove useful, as it has for detecting large emergence events from lakes (e.g., via radar, Masteller andObert, 2000).

Finally, combining the study of drift, emergence and oviposition in the same stream or river ecosystem has thepotential to yield greater ecological understanding than may be obtained through study of any of these processesalone. Measures of any or all of these are, in our experience, likely to prove very complementary to more traditionalbenthic sampling (e.g., Malison et al., 2010; Kennedy et al., 2014, 2016). We highlight some of these possibilitiesbelow.

21.2.2 General ProceduresdDrift

Drift: Drift is easily sampled in most streams by using drift nets set in the water for short amounts of time (Fig. 21.1).Various factors must be considered when sampling drift, including the mesh size and length of the net, length of thesampling period, the number of nets needed for adequate replication, sampling location, and the manner of data analysisand presentation (Brittain and Eikeland, 1988).

The mesh size of the nets will depend on the objectives of the study, but the typical mesh size used is 200e300 mm.Nets with a larger mesh size will not retain small individuals that are abundant in drift such as Chironomidae, resulting ininaccurate conclusions regarding the species composition and magnitude of drift. Nets with a smaller mesh size can beused, for example with a 40e60-mm mesh to capture meiofauna in the samples, but these nets clog rapidly and can only bedeployed for short amounts of time during quantitative investigations of the drift.

Comparisons of the species composition, drift concentrations, drift biomass (mg/m3), and mean size of drifting in-dividuals can be made between habitat units within a stream, across a range of discharges in a stream, across differentstreams, or across time periods (e.g., day vs. night, seasonally). Most investigations of invertebrate drift revolve aroundcomputing drift concentration (# m�3), which requires knowing the number of invertebrates captured by the nets per unitvolume of water filtered by the nets (Naman et al., 2016). Because drift concentrations are often small (i.e., <1 m�3), thisquantity is often expressed as numbers of invertebrates drifting per 100 m3 of water:

Drift Concentration ¼ ½ðNÞð100Þ�=½ðtÞðAÞðVÞ� (21.1)

where N represents number of invertebrates in a sample; t, time that the net was in the stream (s); A, the area of the netmouth that is submerged (m2); and V, mean water velocity at the net mouth (m/s).

As drift nets become clogged with debris, the velocity of water entering the net decreases. Many investigators attemptto account for this by measuring the water velocity in the net at the start and end of the deployment and then estimating theaverage water velocity over the duration of the deployment. However, decreases in velocity associated with clogging arenonlinear and typically follow a logistic pattern, where velocity is high and constant at the start of the deployment until netsbecome appreciably clogged and then velocity drops rapidly and approaches zero (Muehlbauer et al., 2016). Thus, spotmeasurements of velocity at the start and end of the deployment are insufficient, because they do not account for thenonlinear decreases in velocity. Although sophisticated procedures have been developed for estimating the volumesampled by clogged nets (e.g., Faulkner and Copp, 2001), we recommend instead that drift-sampling regimens be designedto avoid net clogging altogether because clogged nets and long deployment times introduce several other sources ofsampling error into drift-concentration estimates.

A major source of sampling error arises from clogged drift nets because they do not actually sample a representativeparcel of water (Smith et al., 1968; McQueen and Yan, 1993). A clogged drift net actually blocks the flow of the streamand forces water to flow around this barrier. However, suspended particles may not flow around the net to the same degree




as the water if their density is greater than water. That is, the inertia of suspended particles such as invertebrates may carrythem into a clogged net to a greater degree than the water that is flowing around the net (Sabol and Topping, 2013).Similarly, if the density of a suspended particle is less than water, these particles may avoid the net to a greater degree thanthe water. Back pressure in a clogged net can also cause particles that have been captured by the net to be flushed out overthe course of the deployment (Muehlbauer et al., 2016).

Filtration efficiency, or the ratio of velocity into the net relative to ambient stream water velocity, is a metric of netperformance (Tranter and Smith, 1968; McQueen and Yan, 1993). Ideally, filtration efficiency of drift nets should be w1,indicating that the sampler is isokinetic and flow conditions inside the net are representative of ambient conditions outsidethe net. At a minimum, filtration efficiency should not be allowed to drop below 0.85 during drift sampling (Smith et al.,1968; Muehlbauer et al., 2016). In our studies of invertebrate drift in rivers of the Western United States, we have foundthat 250-mm nets will only maintain high filtration efficiency for a maximum of 10 min (Kennedy et al., 2014). Thus, ourdrift-sampling collections are limited to 5 min, and by using a net with a large opening (i.e., 50 cm diameter) we are stillable to collect reasonably large samples (i.e., >30 m3). If short-duration drift samples do not adequately capture large taxaat a given site, we recommend that multiple short duration samples that maintain high efficiency be aggregated in the field,rather than extending the duration of individual drift samples beyond acceptable levels of filtration efficiency (i.e., <0.85).Below, we describe an exercise that demonstrates how filtration efficiency of nets declines over time that can be used toidentify maximum deployment times for drift studies (Basic Method 1).

Another source of sampling error that can be introduced when drift nets are deployed for a long period of time occursduring laboratory processing of samples. Specifically, long drift net collections will collect large numbers of invertebrates,which often necessitates subsampling or sample splitting procedures. Although numerous tools and statistical approachesexist for minimizing the error that is introduced with sample splitting in the laboratory (see Chapter 15), these approachesnecessarily introduce another source of sampling error that does not exist when an entire drift sample is processed forinvertebrates.

The length of the net is an important consideration in drift studies, because long nets have a greater surface area forfiltering water and will clog at a slower rate than a short net. The length of nets that are used for sampling drift in rivers isoften reported as the ratio of length:opening (e.g., a 5:1 net is five times as long as the net opening). Most nets are 3:1 butcustom lengths are also available. As the net length increases, however, deploying, retrieving, and washing nets downbecomes more difficult. In our experience a 5:1 net strikes a good balance and allows for reasonably long deployment timesunder most conditions while not being overly cumbersome to retrieve.

Accurate estimates of the volume of water sampled by nets are critical in quantitative studies of the drift. The bestapproach for estimating water volume sampled by a drift net involves mounting a small, inexpensive flow meter in themouth of each net (e.g., General Oceanics model 2030R; Fig. 21.1A,B). We also recommend that ambient velocity bemeasured during each drift net deployment so filtration efficiency of the net can be estimated on-site. This allows samplesto be discarded and new, shorter duration samples to be immediately collected if filtration efficiency is observed to havedropped below acceptable levels (i.e., <0.85). Alternatively, if nets are deployed for short amounts of time, and priorinvestigations have demonstrated deployment times are sufficiently short so as to avoid net clogging (Basic Method 1),then spot measurements of water velocity using a portable meter (e.g., Marsh-McBirney, Son-Tek) at the start and end ofdrift sampling will suffice. Identifying maximum deployment times for drift nets in a particular stream is best done usingportable flow meters that will continuously record and log water velocity readings, so filtration efficiency curves can bedeveloped.

Early studies of diel periodicity in invertebrate drift often involved continuous drift sampling over an entire 24-h cycle,and numerous drift studies continue to use this sampling design. Although this type of exhaustive sampling design isrequired for investigations into diel drift periodicity, there are many applications where this approach may not benecessary. For instance, drift-feeding fish are visual feeders, and their ability to capture prey declines at night. Thus,daytime drift may accurately represent prey availability to drift-feeding fish, and including night-time samples that haveelevated drift concentrations owing to behavioral drift may actually inflate estimates of prey availability (Allan and Russek,1985). Further, Allan and Russek (1985) demonstrated that in some settings a single night-time drift collection shortly afterdark was representative of total night-time drift. Moreover, even the most dedicated stream ecologist is prone to errorswhen sleep deprived, and in our experience this type of nonstop night work inevitably leads to poor quality samples (i.e.,filtration efficiency <0.85, errors on field notes, etc.) that have to be discarded. Thus, we recommend that practicing streamecologists first identify whether 24-h drift sampling is critical to their specific research question before embarking on acontinuous 24-h sampling design. In many cases, drift research questions can be addressed with short duration, high qualitysamples collected during daytime hours that are complemented by more limited and strategic collections of night-time driftsamples.




21.2.3 General ProceduresdEmergence and Postemergent Insects

Traps and nets: There are several techniques that may be used to collect and quantify adult insects as they emerge from astream, or insects that have already emerged and are occupying nearby riparian or terrestrial habitats. With respect to theformer, a number of different types of emergence traps have been developed; their designs, applicability for use underdifferent sampling conditions, and the factors affecting their performance are discussed in detail elsewhere (e.g., Davies,1984; Malison et al., 2010). A common type of emergence trap is pictured in Fig. 21.2 (see Malison et al., 2010 for detailsof trap construction). Measurement of emergence as a part of food web and ecosystem studies has placed new demands onthe techniques used. For example, trap designs have been adjusted (e.g., made more lightweight and inexpensive) tofacilitate studies in which large numbers of measurements must be made simultaneously. Traps typically consist of atriangle- or pyramid-shaped wooden or plastic frame, enclosing an area of 0.5e1.0 m2 and 0.5e1 m high. The sides arecovered with light-colored, fine mesh (typically 500-mm) netting. Many designs include a collection bottle with a funnel orcone-shaped entrance to prevent insects from returning to the net (mounted near the apex of the trap to facilitate removal ofcaptured adults), which can result in higher rates of capture (Cadmus et al., 2016). However, the presence of a collectionbottle can introduce bias because bottles can attract certain taxa more than others (Davies, 1984). Investigators may alsoavoid use of a bottle to keep the design of the trap simpler, to make traps easy to repair in the field, and to avoid the use ofchemicals (which may be undesirable in some settings; Malison et al., 2010). In such cases, a catch may be sewn to theinside of the net w1/3 the distance from top of the trap (see Fig. 21.2B) to capture insects that might otherwise reenter thewater, actively or passively. However, heavy rain and/or wind may compromise samples collected using such a trap. Itmust also be acknowledged that not using a bottle likely introduces its own biases as well (Cadmus et al., 2016), and thatmany potential forms of trap bias remain unevaluated.

Some types of emergence traps are designed to be placed directly on and anchored into the streambed, thereby samplinginsects emerging from a known area (see Smock, 2006). In deeper water, or in studies whose aim may be to generateestimates of emergence flux integrating larger areas of habitat, it is common to employ floating traps (as in Fig. 21.2A).These float by virtue of foam or similar material attached to the frame, and can be secured by loosely connecting theupstream corners of the trap (e.g., using Zip ties) to pieces of rebar driven into the streambed to facilitate continuoussampling as water levels rise or fall. Choice of trap location can be important as well. As described above, habitat units(e.g., pools vs. riffles) may differ in rates of emergence, and composition of insects emerging from the water column islikely to differ from that emerging from the stream bank. Owing to the latter, investigators have adapted the floating trapdesign to capture bank-emerging taxa (Fig. 21.2B)dfor instance, by including overhanging netting on three sides that canbe buried into the bank substrate to prevent insects from escaping and potential predators from entering the trap (Paetzoldet al., 2006; Malison et al., 2010). In the case of large-bodied stoneflies that emerge from riverbanks, estimates ofemergence flux may even be obtained via visual counts of their exuviae (D. Walters, personal communication). Thedeployment duration for traps prior to sample collection will depend upon the questions but is typically not longer than4e5 days to avoid loss of insects via mortality. In addition to the use of a bottle, insects may be removed from a trap usingan aspirator or small vacuum (e.g., BioQuip Hand-Held Vac/Aspirator, BioQuip Products, Rancho Domingues, CA, USA).Below we describe an exercise (Basic Method 3) to quantify and compare emergence flux and composition from differenthabitats within a stream (i.e., pool, riffle, mid-channel, and bank).

Postemergent adult aquatic insects found in riparian or nearby terrestrial habitats are commonly sampled using stickytraps, light traps, funnel nets or a variety of other terrestrial arthropod sampling tools (pitfall, Malaise traps, beating nets,etc.). Sticky traps or light traps (Fig. 21.2C,D) seem best to determine the abundance and species composition of activeadults or the dispersal distances traveled laterally from the stream channel by individuals (Collier and Smith, 1995; Kovatset al., 1996), whereas funnel nets (Turner and Williams, 2000) and flat-design sticky traps (Bird and Hynes, 1981; Smithet al., 2014) may be used to quantify the direction of adult flights, such as up- and downstreams (but see Macneale et al.,2004). Sticky traps collect in a more passive fashion than do light traps, which in some cases may be advantageous. Theycan also be left unattended for several weeks, allowing time-integrated estimates of aerial insect abundance. Sticky trapsare often cylindrical (typically 0.1e0.2 m2), for instance constructed with sheets of acetate coated with sticky resin(e.g., Tanglefoot, The Tanglefoot Company, Grand Rapids, MI, USA) and suspended (from poles or sometimes vegeta-tion) 1e2 m above the ground. However, height above the ground may influence the composition of aerial insects captured(Jackson and Resh, 1989; Smith et al., 2016), and, indeed, there are likely important patterns in insect distribution withinthe “airscape,” which have yet to be investigated (Gurnell et al., 2016). Another sticky trap design, described by Smithet al. (2014), uses 150-mm Petri dishes coated with resin, suspended on poles. This trap, which may be used to quantifyflight direction, can be coated with resin prior to transport to the field with the lid on, which also facilitates efficientretrieval of the trap (cylindrical traps must be covered upon retrieval, often with a sheet of acetate or cellophane).




Disadvantages to using sticky traps include the negative effect of the resin on insect specimens (identification is typicallydone with the trap under magnification, rather than by removing insects from the trap), the potential for accumulation ofinsects to influence trap efficiency (Muirhead-Thompson, 2012), and the exposure of trapped insects to weather and evenpredators (e.g., birds, bears) that may cause specimen deterioration.

Light traps are an active-sampling tool used to collect and characterize postemergent adult insects; they range fromtowers with automated collection mechanisms to simple, battery-operated lamps set in close proximity to a killing agent(e.g., a bottle or tray of ethanol; Fig. 21.2D). Benefits to their use include rapid collection of a “snapshot” of insectoccurrence, often of a large number of individuals, good specimen preservation and the fact that, owing to insect attractionto the light, a single trap provides a sample integrated across a large area. On the other hand, the latter may be a disad-vantage if more absolute estimates of insect occurrence are desired. Light traps capture species that are nocturnally active,and differences in light attraction among taxa also introduce bias (Basset et al., 1997; Yi et al., 2012; Muirhead-Thompson,2012). Investigations of stream-riparian food webs linked by emergent aquatic insects frequently employ sticky traps orlight traps to estimate indices of emergence flux and/or availability of prey for terrestrial insectivores. However, the variousforms of bias described above require consideration in such studies, and more direct measurements of emergence may berequired if the aim is to generate absolute estimates of this flux from particular habitats. Below we describe an exercise(Basic Method 4) employing sticky traps to quantify dispersal distances of adult aquatic insects laterally from a stream. Wealso outline procedures (Advanced Method 3) by which measures of emergence or postemergent insects may be used tocompare the availability of emergent insects as potential prey for terrestrial insectivores among streams.

21.2.4 General ProceduresdOviposition by Stream Insects

Aquatic insect oviposition methods can be generally categorized in four groups: (1) methods for quantifying the presenceof egg-laying females; (2) methods for quantifying the contribution of egg-laying females to predator and detrital foodwebs; (3) observations of insect oviposition behaviors, and (4) methods for recording the distribution of eggs. Studiesfocused on the presence of food web contributions of gravid females generally utilize very similar procedures to theemergent adult and drift methods, often with modifications for capturing adults entering the water from the air (e.g., air-facing sticky traps, Encalada and Peckarsky, 2007). With some modification, sticky traps such as those described earlier inthis chapter (e.g., Smith et al., 2014), stream exclosures (e.g., Nakano et al., 1999), or drift nets deployed during specifictimes (e.g., Encalada and Peckarsky, 2007) could all conceivably be used toward these purposes (see Basic Methods 2e3).

Approaches for understanding the site selection process, behaviors, and related environmental cues for oviposition canrange from relatively simple field observations to rigorous protocols. A great deal can be learned about insect ovipositionby observing the behaviors of adult insects and thus such approaches are highly recommended, especially as a pilot formore involved field studies. Field observations do not require much sampling gear beyond a pair of binoculars and somesample vials to collect oviposited eggs in the event larvae or adults are needed for identification (see Basic Method 5).

Beyond field observations, the methods for studying oviposition behaviors or the distribution of eggs are similar toother procedures designed to quantify habitat site selection (Garshelis, 2000). Common approaches quantify both theavailable and utilized habitat to determine the environmental factors associated with either the presence or absence of aspecies. This often requires consideration of species-specific habitat needs in the context of the hierarchical arrangement ofstream and river systems: river segments, reaches, channel units, and microhabitats (see Chapter 2). Additionally, given thepatchy distribution of egg masses on substrates that are not present in equal proportions from segment-to-segment or reach-to-reach, structured, nonrandom sampling of commonly preferred microhabitats (e.g., emergent rocks or vegetation) maybe required for adequate egg detection (e.g., Encalada and Peckarsky, 2006). Alternatively, a two-stage sampling procedurethat characterizes both habitat availability and utilization can be employed in order to quantify egg mass densitiesthroughout a reach (Reich and Downes, 2003). Regardless, these methods are predicated on the ability to identify eggmasses as belonging to a given species. Because this information is not readily available for many aquatic insects, therearing of eggs to larvae or adults may be necessary to permit identification (see Basic Method 6).

21.3 SPECIFIC METHODS

21.3.1 Basic Method 1: Filtration Efficiency of Drift Nets

Objective: Determine maximum deployment times for drift nets in a stream.

1. Place a single drift net in a stream at a location with moderate- to fast-water velocity for that stream (Fig. 21.1A,D).Alternatively, if sufficient flow meters are available, drift nets can be placed in both slow and fast velocity habitats to




demonstrate how the rate of clogging varies with water velocity. Prior to actually deploying the drift net, dunk the net inthe stream repeatedly as air bubbles on a dry net can temporarily reduce filtration efficiency at the start of a deployment.Orient the net so that the mouth is perpendicular to the direction of flow and anchor it in place with rods driven into thesubstrate.

2. Use a recording flow meter (e.g., model 2030R, General Oceanics, Miami, Florida) affixed at the mouth of each netusing string, zip ties, or cable to record the velocity of water entering the net during the deployment at 1-min intervals(see Chapter 3). Position a second recording flow meter 0.5e1 m to the side of the net to continuously record ambientstream water velocity at 1-min intervals. Periodically, compare the ambient stream velocity with the velocity estimatesat the net to identify when the velocity into the net has slowed appreciably. Remove the net when the velocity into thenet is <20% of ambient.

3. Discard the contents of the net. Thoroughly clean the net and repeat steps one to two to obtain multiple estimates offiltration efficiency over time.

4. Make a plot showing the filtration efficiency (y-axis) over the duration of the deployment (x-axis), where filtration effi-ciency ¼ net velocity/ambient velocity. Identify how long it took for the filtration efficiency to drop below 0.85; thisrepresents the maximum amount of time that drift nets should deployed during future drift studies that have similar flowand suspended solid conditions. Use statistical software to fit linear, negative exponential, and logistic curves to thefiltration efficiency graph. Conduct analyses to identify which of these functional relationships best describes how wa-ter velocity changes over time and as the net becomes clogged.

21.3.2 Basic Method 2: Drift Concentrations Among Habitats

Objective: Quantify variation in drift concentration and biomass among habitats in a stream.

1. Place a single drift net in each of several different habitat units (e.g., riffle, glide, pool) in a stream reach (Fig. 21.1A).Orient the net so that the mouth is perpendicular to the direction of flow and anchor it in place with rods driven into thesubstrate. The net should be positioned at mid-depth in the water column or, if the stream is shallow, the bottoms of thenet should be 2e3 cm above the sediment to reduce the possibility of invertebrates crawling into the net.

2. Prior to actually deploying the drift net, dunk the net in the stream repeatedly as air bubbles on a dry net can temporarilyreduce filtration efficiency at the start of a deployment. After wetting down the net, attach the sampling bucket to theend of the net. As above, use the flow meter to record the velocity of water entering the net during the deployment.Record the starting value displayed on the flow meter before putting the net into the stream. At the same time the driftnet is put into the stream, place the second flow meter 0.5e1 m to the side of the net to record ambient stream watervelocity.

3. Record the width of the drift net and the height of water passing into the drift net. Measure height at three differentplaces if the drift net is not fully submerged.

4. Remove the drift nets from the stream after 15 min or when a filtration efficiency of 0.85 is estimated to occur (seeBasic Method 1), whichever is shorter. Simultaneously, remove the ambient flow meter. Record the ending value dis-played on both flow meters. Wash the contents of the net into a bucket partly filled with water. Use forceps to removeany invertebrates that remain clinging to the inside of the nets. Wash the contents of the bucket through a sieve with amesh size equal to or smaller than that of the net. Preserve the material from each net separately in bottles or sealablebags with 70% ethanol (final concentration). Label the samples with location, date of collection, habitat unit, timeperiod of sampling, and investigator.

5. Repeat steps one to four for a minimum of three consecutive samples in each habitat unit. Nets should be placed in thesame location during each sampling interval.

6. In the laboratory, separate all organisms from the debris in the samples. This is best accomplished using a stereomi-croscope at low power. Count the number of invertebrates in each sample.

7. To determine the species’ composition, identify and enumerate the taxa of invertebrates in the samples (see Chapters 20and 25). Conduct analyses to determine if there are differences in the species composition and relative abundance ofdrifting organisms among habitats.

8. Determine the size of each individual collected in the samples. This can be accomplished by various methods: (1) Mea-sure the length of all invertebrates in each sample using an ocular micrometer fitted in a stereo-microscope. Calculatethe mass of each individual using published regression equations relating the organism length to mass for a wide varietyof aquatic invertebrates (Benke et al., 1999). Note that this approach is required for drift-foraging bioenergetics modelsthat require as inputs the biomass of prey discretized into 1-mm size bins of length, because prey selection by fish is




related to the length of prey (Hayes et al., 2000; Dodrill and Yackulic, 2016); or (2) place all invertebrates collected in asample together in an aluminum weighing pan and dry them in a drying oven for 24 h at 60�C. Weigh the pooled in-vertebrates on an electronic balance.

9. Calculate the mean individual dry mass of the drifting organisms (¼pooled dry mass/number of individuals in the sam-ple) and the mean drift concentration of invertebrates in each sample. Conduct comparisons to determine whether thereare differences in drift concentration among habitat units.

21.3.3 Advanced Method 1: Quantifying Active Drift of Stream Invertebrates

Objective: Compare the drift concentration of invertebrates in a stream between day and night.

1. Place drift nets across the stream channel (Fig. 21.1D). Nets are placed in the stream with the net face perpendicular tothe direction of flow and anchored with rods driven into the substrate. Nets should be positioned at mid-depth in thewater column or, if the stream is shallow, the bottoms of the nets should be 2e3 cm above the sediment to reduce thepossibility of invertebrates crawling into the nets. Presuming channel width permits, use at least three nets positionedalong a cross-section to gather samples to encompass spatial variation in the drift estimate for a site.

2. Record the width of the drift net and the height of water passing into the drift net. Measure height at three differentplaces if the drift net is not fully submerged.

3. Remove the drift nets from the stream after 15 min or when a filtration efficiency of 0.85 is estimated to occur (seeBasic Method 1), whichever is shorter. Simultaneously, remove the ambient flow meter. Record the ending value dis-played on both flow meters. Wash the contents of the net into a bucket partly filled with water. Use forceps to removeany invertebrates that remain clinging to the inside of the nets. Wash the contents of the bucket through a sieve with amesh size equal to or smaller than that of the net. Preserve the material from each net separately in bottles or sealablebags with 70% ethanol (final concentration). Label the samples with location, date of collection, habitat unit, timeperiod of sampling, and investigator.

3. Repeat steps one to three for a minimum for three consecutive daytime samples and then repeat again after dark toobtain night-time samples. Nets should be placed in the same location during each sampling interval.

4. In the laboratory, separate all organisms from the debris in the samples. This is best accomplished using a stereomi-croscope at low power. Count the number of invertebrates in each sample.

5. Calculate the mean drift concentration of invertebrates in the stream during each time interval. Construct a curveshowing the change in drift concentration over time. Conduct analysis to evaluate any difference between day and nightdrift concentrations.

21.3.4 Advanced Method 2: Quantifying Drift in Unwadeable Rivers

Objective: Compare the vertical distribution of drift in deep rivers.

1. Prior to starting any boat-based drift-sampling program, familiarize yourself with safety procedures for working fromboats, read manuals describing the operation and maintenance of the equipment that will be used, and be cognizant ofthe hazards and risks that water quality sampling entails.

2. Collecting drift in large rivers from a boat requires different types of equipment than sampling drift in wadeablestreams. Plankton nets with a circular opening and bridle are used, because nets are raised and lowered through thewater column to collect drift samples rather than being staked into the benthos. Flow strength is high in large rivers,so large weights (e.g., 75-lb US Geological Survey Columbus-Type sounding weights; Rickly Hydrological) areneeded to keep nets from being pushed downstream. A hand powered winch (A-reel) that is mounted to a frame atchest height is used to raise and lower drift nets through the water column. A boom or crane that extends over thebow of the boat allows the winch cable to be raised and lowered smoothly over a pulley. Even with large weights, driftnets are often pushed downstream such that the winch cable may rub on the bow of the boat; a roller can be mounted tothe front of the boat to prevent wear and friction on the boat and cable. Connectors (Type B; Rickly Hydrological) areused to attach a 1-m length of chain to the end of the cable, and then the sounding weight is attached to the bottom ofthe chain (Fig. 21.1C). The circular drift net with bridle is then clipped halfway down the chain using a carabiner (seeFig. 21.1B). It is important to keep heavy-duty wire cutters nearby in case the drift net becomes caught at the bottomand the winch cable needs to be cut. Also consider the type of boat available (i.e., propeller vs. jet drive) when pro-curing nets, as long nets are easily destroyed in propellers.




3. Position the boat facing the upstream in moderate current and hold position by motoring or tying the boat off to a buoyor similar. Lift the sounding weight off the boat and slowly lower it without dropping it; the standard 0.25-cm diametercable that comes on A-reels is rated for static loads >500 lbs, but dropping a heavy weight, even from a short height,creates a dynamic load that can easily break the winch cable. Use the winch to lower the weight so that it is justtouching the water surface. Adjust the depth gage on the winch so it reads zero. Lower the weight to the river bottomto determine depth and record this value and then raise weight back to the surface. Record the starting value of the flowmeter that is positioned in the net mouth and clip the net into the drop chain. Affix the ambient flow meter to the sound-ing weight and record its starting value.

4. Slowly raise and lower the drift net using the A-reel for the duration of the 5e10 min sample. Never lower the drift netto closer than 0.5 m from the bottom to prevent benthic material from being dislodged and captured by the net, andnever allow the net to break the surface of the water. Each round trip vertical sample represents a “transit”; strivefor a minimum of two transits for each drift sample (Topping et al., 2011). Maintain a constant rate of movementboth up and down the water column so that each transit samples all depths equally. After 5e10 min, pull the netout of the water and record the ending time of the deployment and the values displayed on both the net and ambientflow meter. Estimate the filtration efficiency of the net by comparing flow meter readings to ensure it was above 0.85.Discard samples and shorten deployment times if filtration efficiency is less than 0.85. Use an on-board washdownsprayer to hose the sides of the net off and into the collecting bucket. Alternatively, dunk the net in the river severaltimes to flush the contents of the net into the collecting bucket.

5. Follow procedures in the Basic Methods for preserving and laboratory processing of samples.6. Repeat steps one to four and collect multiple depth integrated samples and also samples where the net is parked near

bottom, at mid-depth, and near-surface. The ordering of different types of samples should be randomized over thecourse of the collection (i.e., do not collect all depth integrated first, then all near-bottom next).

7 Calculate the mean drift concentration and biomass of invertebrates for each type of sample (i.e., depth integrated anddifferent fixed depths).

21.3.5 Basic Method 3: Quantifying Emergence of Adult Stream Insects

Objective: Measure and compare the numbers, fluxes, and composition of adult insects that emerge from different habitatswithin a stream. Optionally, determine if emergence differs between day and night.

1. Construct (e.g., as per description above and, for instance, Malison et al., 2010; Cadmus et al., 2016) and deployfloating emergence traps (Fig. 21.2A) over the primary habitats (e.g., pools, riffles, glides) in a stream. For instance,some investigators (e.g., Iwata, 2007) have reported greater emergence from pools compared to riffles; is this whatyou observe? In a paired fashion, deploy modified traps encompassing the stream bank adjacent to each of those withina habitat unit. It is common to deploy six to eight traps per habitat unit (three to four floating, three to four bankside)within a stream reach. However, determining the number of traps to deploy per-habitat type should depend upon theanticipated variability of emergence and magnitude of difference, and a more sophisticated way of addressing the ques-tion of sample size is to conduct a power analysis (e.g., see Aho, 2014). This requires use of an existing emergence dataset, which ideally is generated via pilot measurements, but often draws upon published data from a site that may havesimilar characteristics to those being studied.

2. After 24e48 h, remove all insects from the trap using an aspirator (or from the collection bottle), and preserve them in70% ethanol. Label the sample with location, trap number, date, sampling time period, and investigator. As describedabove, emergence can be highly heterogeneous in time, so many time periods may need to be sampled if a comparisonis to be representative of the habitats or the temporal scope of inference is intended to be seasonal or annual.

3. Optional: Divide sampling into day and night periods, recording the number of hours the traps were in place duringboth periods.

4. In the laboratory, identify (using keys such as those found in Merritt et al., 2008) and count the number of insectscollected in each trap. In addition, measure the lengths of individuals.

5. Calculate the flux, or emergence flux, or production [g dry mass (DM) m�2 d�1] using direct weighing or publishedlengthmass regressions (e.g., Rogers et al., 1977; Sample et al., 1993; Stagliano et al., 1998; Sabo et al., 2002). If regres-sions do not exist for a particular taxon, then use one from a related taxon that has a similar morphology or weigh directly.

6. Express the results as the mean number of taxa, individuals and/or total dry biomass emerging per square meter perhour from each habitat or during the day and night. Conduct analyses describing differences in the number of taxa,individuals or total biomass emerging from the different habitats or during the day and night.




21.3.6 Basic Method 4: Investigating Lateral Dispersion of Emergent Stream Insects

Objective: Quantify the extent to which adult aquatic insects disperse laterally from a stream. Optionally, determinedirectionality of adult insect flight.

1. Construct (e.g., as per description above and, for instance, Smith et al., 2014) sticky traps (Fig. 21.2C) and deploy themat sampling locations at a minimum of five locations, such as 0, 25, 50, 100, and 200 m on a transect away from thestream. Other spacing regimes may be used depending on the size of the drainage basin and its geography.

2. Set up sticky traps at each sampling point along the transect, noting the time of their deployment.3. Optional: Evaluate directionality of flight by aerial insects. Use a flat-trap design and deploy traps at each point along

the transect to capture insects flying in each cardinal direction. For instance, if using the Smith et al. (2014) trap design,position four traps on each pole, mounted at right angles.

4. Allow sticky traps to collect for at least 24 h (ideally a few days, as temporal heterogeneity of emergence can be high,and comparisons may be more representative if integrated over more time). Collect traps (noting time) and label themas above.

5. In the laboratory, identify, count, and measure the number of insects of aquatic taxa collected on each trap. Dependingupon the size of the trap, this may require use of a stereomicroscope mounted on a movable arm, or, in cases wheredetailed taxonomic resolution is not required, a magnifying lens mounted on an arm (such as are sometimes used forreading) may also be used.

6. Calculate the emergence biomass intercepted by each trap [g dry mass (DM) m�2 d�1] using published lengthmassregressions, as described in Basic Method 3.

7. Express the results as the number of taxa, individuals, or total emergence flux captured at each trap. If traps were oper-ated for different time periods, express the results per a standard time period. Develop a graph showing the change innumber of taxa, individuals, and/or flux with distance from the stream and/or compare directionality of insect dispersalat each location.

21.3.7 Advanced Method 3: Investigating Availability of Emergent Insects as PotentialPrey for Terrestrial Insectivores

Objective: Compare the availability of emergent adult insects as potential prey for terrestrial insectivores among stream-riparian ecosystems.

1. Deploy emergence traps (as per Basic Method 3) in reaches of the same or different streams across which you intend tomake comparison. If the goal is to represent what may be available to terrestrial insectivores, then it is important thattraps sample emergence from the suite of habitats that contribute the most to this availability (a pilot comparison usingBasic Method 3 may be conducted to determine this). Construct a map that will allow you to estimate the area of eachhabitat type in each reach (see Chapter 3) stratify the habitats within the site you wish to represent, and install at leastsix to eight traps per habitat unit throughout each reach (e.g., three to four floating, three to four bank side; or conductpower analysis to determine trap number using data from Basic Method 3).

2. Optional: Use sticky traps to make comparison among reaches of an index of postemergent insect availability as preyfor terrestrial insectivores. This may be advantageous if it is not possible to visit sites on the short time interval requiredfor emergence traps, or if study questions pertain to distribution of potential insect prey extending beyond habitatsimmediately adjacent to the stream. Install at least four transects of sticky traps (as per Basic Method 4), alternatingthe side of the stream from which each extends. Alternatively, in settings where traps cannot be left unattended forlonger periods and where a rapid “snapshot” is required, light traps may be used in a similar fashion (but see caveatsregarding biases associated with both trap types above).

3. Collect insects from emergence traps for at least three sample periods, with each deployment encompassing at least24 h, but ideally 2e3 days. If the temporal scope of inference is intended to be seasonal or annual (as would typicallybe necessary to gauge prey availability and possible responses by insectivore populations), more sampling periods willlikely be needed.

4. Follow procedures in the Basic Methods for preserving and laboratory processing of samples, and estimate the totalbiomass flux for each sample, as well as for different taxa [g dry mass (DM) m�2 d�1].

5. Calculate the average emergence flux from different habitats within each reach, the average for each reach-sampleperiod combination, and the average for each reach across all sampling periods. To obtain the most representativeestimates of these, calculate them as averages weighted by the occurrence of each habitat type sampled within each




reach. In addition, riparian insectivores may respond to the total flux of emerging prey that crosses the stream-riparianboundary (i.e., the total food available), rather than the mean flux per unit area across the stream surface (Gratton andVander Zanden, 2009). Therefore, estimate total emergence flux at the reach scale (Benjamin et al., 2013) as averageflux associated with each habitat type times the surface area of each habitat occurring within the reach (as derived fromthe map for each reach).

6. Compare estimates of mean and total emergence flux among reaches. Also, assess whether reaches differed in thetiming of emergence among the sample periods. Finally, compare the composition of emergence flux among reaches.Different terrestrial insectivores may exhibit preference for or avoidance of different adult aquatic insects as prey(Marczak et al., 2007) and may respond to heterogeneity in prey availability at different scales (Power and Rainey,2000). For instance, most web-building spiders are unlikely to capture large-bodied taxa such as dragonflies or largestoneflies and may make more use of aerial, smaller taxa such as midges or mayflies (e.g., Kato et al., 2003), whereasground-dwelling predators (e.g., many spiders, beetles, reptiles and amphibians) may be responsive to availability ofprey that spend more time crawling than flying (e.g., Paetzold et al., 2006). Similarly, predators that capture insects viaaerial, hawking behavior (like many birds and bats) are likely to be more responsive to flying insects, whereas thosethat mainly glean from vegetation or other surfaces may exhibit other preferences (e.g., Hagen and Sabo, 2011). Withthese possibilities in mind, estimate emergence flux that may represent availability of prey for different types of pred-ators and compare among reaches.

21.3.8 Basic Method 5: Observing Oviposition by Stream Insects

Objective: Record the timing and behavior of insect egg-laying.

1. Consult fly-fishing “hatch” charts, species descriptions in taxonomic journals, or other local or species-specific sourcesto anticipate likely emergence times. In the absence of such information, dusk in late spring or early summer is a peakegg-laying time for many insects.

2. Find a reach of stream that allows observation of multiple habitats such as riffles, rocky shores, littoral vegetation, andbackwaters.

3. Watch for insects flying around the stream. Record any behaviors of these insects, such as mating swarms, mate pairsflying together, or females dipping their abdomens in the water, along with the timing of these activities. Payspecial attention to any adult insects that appear to be flying toward the water or crawling on rocks or other emergentsubstrate.

4. After adults leave a habitat (e.g., a boulder), check the spot for any eggs. Special attention should be paid to large emer-gent substrates, which may need to be partially removed from the water to locate eggs. If possible, collect any eggsalong with their corresponding adults, as this makes identification much simpler (compare with Basic Method 6). Stan-dard insect nets or sticky traps (see Basic Method 4) may be useful for this purpose.

5. If any gravid females are captured, pay special attention to any eggs that may be laid in the net or on the sticky trapduring capture to assist in the identification of unknown egg masses.

6. Record the habitats used or avoided by adults, both for mating and for egg-laying. If possible, also record the charac-teristics, locations, and densities of the egg masses.

21.3.9 Basic Method 6: Rearing Stream Insect Eggs

Objective: Hatch insect eggs to larvae for the purposes of identifying egg masses to species, or establishing an experimentallaboratory stock.

1. Search for eggs in stream habitats. If specific taxa are of interest, descriptions in taxonomic journals may include pic-tures of the species’ egg masses for reference. For many species, check specifically for eggs in stream margin habitats,and on the sides or undersides of rocks, wood, or other large or emergent substrates.

2. Scrape eggs as delicately as possible from substrates, using a scalpel or similar tool. Place eggs in vials containing riverwater for transport to lab. If you seek to minimize egg-hatching outside of the lab, maintain water temperatures at orbelow that of the studied system.

3. Egg-rearing in the lab is relatively easy for taxa with short gestation times, such as many Diptera, Ephemeroptera, andTrichoptera. In these cases, simply replace the water in the vial every 2e3 days to minimize fungal or bacterial growth.For all lab-rearing scenarios, the water should be river water (which, in some cases, may need to be filtered) or dech-lorinated water, kept at or below room temperature.




4. For taxa with longer gestation times or sensitivities to high temperatures (e.g., Pteronarcyidae), care needs to be takento regulate temperature (w20�C), ensure sufficient oxygenation, and mimic natural diurnal light patterns. In suchinstances, eggs can be reared in containers with an air stone in a climate controlled room.

5. Count the eggs and record physical characteristics of the eggs as they mature, and check on the vials at least daily tolook for hatched larvae. Counting the eggs can be made easier by first photographing the eggs under a dissectingmicroscope and performing the counts on the image.

6. Once hatched, more care is required to rear individuals to late instars or adults for identification. Rearing containerssuch as small plastic food storage containers can be fitted with an air stone and situated in a climate controlledroom to rear larvae. Water should still be changed regularly, and basic habitat for larvae such as small gravel or sticksshould be provided. Small pipettes are very helpful for manipulating larvae.

7. Depending on the feeding ecology of the aquatic taxa, biofilm covered stream rocks, leaves conditioned in stream waterfor approximately one week, small midges, or store-bought fish food need to be supplied regularly. In many instances,mortality rates are minimized by providing more than one food supply.

8. Record proportional hatch success, the growth of instars, the feeding regimen, and identify larvae once they reach suf-ficient size.

21.3.10 Advanced Method 4: Characterizing Habitat Availability and Site Selection forOviposition

Objective: Quantify differences in oviposition site selection as a function of environmental factors such as channel unittypes, location within channel units, and substrate types.

1. Identify and delineate the stream segments to be sampled using maps, imagery, or field-based methods presented inChapter 2.

2. Randomly choose reaches for sampling within each segment. The number of replicate reaches will depend on themagnitude of differences to be detected among segments and the variance within each reach.

3. Within each reach, two types of points will be sampled: Systematic points to quantify both habitat availability andoviposition-site selection and stratified points for oviposition-site selection for underrepresented habitats.

4. Distribute five transects, oriented perpendicular to the flow, equally throughout the reach. Transects will span the extentof wetted channel or from the wetted edge to the channel thalweg.

5. Sample 10 points along each transect to characterize the channel unit type; water depth and velocity; substrate size, typeand emergent area; distance from bankfull; other desired physical characteristics of the stream or river for a total of 50systematics points per reach. Equally space the points across the entire channel, or out to the extent to which wading issafe.

6. Supplement the 50 systematic points per reach with up to 15 additional random points per strata (i.e., stratified points),where strata can include microhabitats such as submerged mineral substrate, emergent mineral substrate, submergedwood, emergent wood, or emergent vegetation.

7. At each of the systematic and stratified sample points, examine a single substrate particle for egg masses, enumeratingthe number of egg masses by egg type.

8. Measure and record values for the physical variables described in step five at each of the systematic and stratified sam-ple points.

9. Conduct analyses to investigate differences in egg density (response variable) as a function of reach type, channel unittypes, substrate types, egg location within a channel unit, and velocity.

21.4 QUESTIONS

1. What aspects of drift nets influence the time at which clogging occurs? What aspects of the deployment site influencethe time to net clogging?

2. What factors might be responsible for observed differences in drift among habitat types? Which habitats afford thehighest prey delivery rates for drift-feeding fishes?

3. What factors might be responsible for observed differences in the vertical distribution of drift in a large river?What type of sampling approach (i.e., vertically integrated or fixed depth) is most appropriate for estimating total driftloads for a reach (# s�1) versus estimating the amount of drift biomass that is available to drift-feeding fish in a deepriver?




4. What differences might occur in drift concentrations and the species drifting as stream size (order) increases? Wouldyou expect seasonal differences in drift concentrations? Why?

5. What factors might be responsible for differences in the numbers or fluxes of emerging insects among different hab-itats? What factors might cause day-to-day, day versus night variation, or differences in emergence timing amongdifferent streams?

6. If composition of emergence differed among habitats, stream reaches, or samples collected using different traps, whymight this be?

7. Benthic sampling devices (e.g., Chapter 15) and emergence traps often provide very different estimates of the speciescomposition of stream invertebrate communities. What are some of the reasons for these differences?

8. Do the numbers of adult taxa collected laterally from the channel (via sticky traps) show an exponential decrease withdistance from the channel? Why or why not?

9. What ecological advantages might the specific timing of oviposition confer on population success? Consider this bothin terms of time-of-day and any synchronous timing of egg-laying within a population.

10. Where are insect eggs being laid? How might egg morphology vary depending on habitat conditions?

21.5 MATERIALS AND SUPPLIES

Field MaterialsRectangular drift nets and metal holding rods (wadeable streams)Circular drift (plankton) nets, sounding weights, and winch (deep rivers)Emergence traps (floating, and bank-side)Sticky traps (including covers or cellophane for retrieval)Current velocity meterMeter sticks or tapesBuckets (5 gallon)Sieves (series of mesh sizes)ForcepsAspiratorEthanol (70%)Bottles or vials or sealable bagsLabeling paper and pencilsHip boots or wadersJeweler’s loupe or magnifying glassBinocularsScalpel

Laboratory MaterialsStereomicroscopesOcular micrometer (optional)Digital camera mounted on microscope (optional)Drying oven (optional)Electronic balance (optional)Rose BengalAluminum weighing pansAcrylate adhesiveSoft forcepsClear, multiwell plates or dishes with lidsEthanolDechlorinated waterVialsSubsampling deviceAir stone




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