Journal of Insect Physiology - COnnecting REpositories · Respiration patterns Ventilation movement...

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Respiration patterns of resting wasps (Vespula sp.) Helmut Käfer, Helmut Kovac , Anton Stabentheiner Institut für Zoologie, Karl-Franzens-Universität Graz, Universitätsplatz 2, A-8010 Graz, Austria article info Article history: Received 12 December 2012 Accepted 30 January 2013 Available online 9 February 2013 Keywords: Wasp Vespula Respiration patterns Ventilation movement Resting metabolism Temperature abstract We investigated the respiration patterns of wasps (Vespula sp.) in their viable temperature range (2.9– 42.4 °C) by measuring CO 2 production and locomotor and endothermic activity. Wasps showed cycles of an interburst–burst type at low ambient temperatures (T a <5 °C) or typical discontinuous gas exchange patterns with closed, flutter and open phases. At high T a of >31 °C, CO 2 emis- sion became cyclic. With rising T a they enhanced CO 2 -emission primarily by an exponential increase in respiration frequency, from 2.6 mHz at 4.7 °C to 74 mHz at 39.7 °C. In the same range of T a CO 2 release per cycle decreased from 38.9 to 26.4 llg 1 cycle 1 . A comparison of wasps with other insects showed that they are among the insects with a low respiratory frequency at a given resting metabolic rate (RMR), and a relatively flat increase of respiratory frequency with RMR. CO 2 emission was always accompanied by abdominal respiration movements in all open phases and in 71.4% of the flutter phases, often accompanied by body movements. Results suggest that resting wasps gain their highly efficient gas exchange to a considerable extent via the length and type of respiration movements. Ó 2013 Elsevier Ltd. 1. Introduction Insects may vary stupendously in their modes of gas exchange (Gibbs and Johnson, 2004), both among (Hadley, 1994; Lighton, 1996; Sláma, 1999; Terblanche et al., 2008c) and within species (Chown et al., 2002; Irlich et al., 2009; Kuusik et al., 2004; Marais and Chown, 2003), and even within the same individual (Chown, 2001; Kovac et al., 2007; Snelling et al., 2012). One particular res- piration pattern in both flying and flightless insects is well known as discontinuous gas exchange cycle (DGC, for reviews see Chown et al. 2006b; Lighton, 1996; Sláma, 1988). Many insects show this pattern when at rest, at least at the lower to medium temperatures of their thermal range. Typical DGCs consist of a closed or constric- tion phase with spiracles shut and little to no external gas ex- change (Bridges et al., 1980). O 2 inside the insect is metabolized, while CO 2 accumulates in the tracheae and in part is buffered in the hemolymph. This causes a drop in the total intratracheal pres- sure (Buck and Keister, 1955, 1958; Buck and Friedman, 1958; Hetz et al., 1994). In the following flutter phase single spiracles open and close rapidly. Gas exchange works here due to convection and dif- fusion. Small amounts of O 2 are inhaled to sustain a certain low le- vel of PO 2 for a minimum O 2 delivery to the insect’s metabolizing tissues (Hetz and Bradley, 2005; Lighton, 1996). The CO 2 level keeps rising in the hemolymph during the flutter phase, as only small amounts of CO 2 are exhaled (Buck, 1958). As accumulated CO 2 reaches a trigger threshold, a massive amount exits from the tracheal system to the environment in the open-spiracle phase (Lighton, 1996; Schneiderman and Williams, 1955). CO 2 is as- sumed to act directly at the spiracular muscles, with little central nervous control (Hoyle, 1961); however, Bustami and Hustert (2000), Bustami et al. (2002) and Woodman et al. (2008) found contrary evidence. Discontinuous gas exchange was hypothesized to be an adapta- tion aimed at minimizing water loss from the tracheae (hygric model, Chown, 2002, 2006a; Dingha et al., 2005; Duncan et al., 2002b; Hadley, 1994; Kivimägi et al., 2011; Williams and Bradley, 1998; Williams et al., 1998, 2010), though findings by Contreras and Bradley (2009), Gibbs and Johnson (2004) and Sláma et al. (2007) call into question the universal validity of this model. Other explanations suggest that it developed to allow sufficient gas ex- change in subterranean, CO 2 rich environments (chthonic model, Lighton and Berrigan, 1995). A combination of these two models is the hygric-chthonic hypothesis (Lighton, 1998). An alternative explanation suggests that it minimizes oxygen toxicity (Bradley, 2000; Hetz and Bradley, 2005). The variation of respiration pat- terns has been well investigated in different species (Basson and Terblanche, 2011; Chown et al., 2006a; Groenewald et al., 2012; Klok and Chown, 2005; Kovac et al., 2007; Nespolo et al., 2007; Terblanche et al., 2008a; Williams et al., 2010). Such an analysis is lacking in vespine wasps. This is especially interesting because Vespula sp. show an overall higher level and a steeper incline in resting metabolism with increasing ambient temperature (high Q 10 ) than many other insects (see Käfer et al., 2012). In this paper, therefore, we investigated the characteristics of the respiration 0022-1910 Ó 2013 Elsevier Ltd. http://dx.doi.org/10.1016/j.jinsphys.2013.01.012 Corresponding authors. Tel.: +43 316 380 5705; fax: +43 316 380 9875. E-mail addresses: [email protected] (H. Käfer), [email protected] (H. Kovac), [email protected] (A. Stabentheiner). Journal of Insect Physiology 59 (2013) 475–486 Contents lists available at SciVerse ScienceDirect Journal of Insect Physiology journal homepage: www.elsevier.com/locate/jinsphys Open access under CC BY-NC-ND license. Open access under CC BY-NC-ND license.

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Page 1: Journal of Insect Physiology - COnnecting REpositories · Respiration patterns Ventilation movement Resting metabolism Temperature abstract We investigated the respiration patterns

Journal of Insect Physiology 59 (2013) 475–486

Contents lists available at SciVerse ScienceDirect

Journal of Insect Physiology

journal homepage: www.elsevier .com/ locate/ j insphys

Respiration patterns of resting wasps (Vespula sp.)

Helmut Käfer, Helmut Kovac ⇑, Anton Stabentheiner ⇑Institut für Zoologie, Karl-Franzens-Universität Graz, Universitätsplatz 2, A-8010 Graz, Austria

a r t i c l e i n f o a b s t r a c t

Article history:Received 12 December 2012Accepted 30 January 2013Available online 9 February 2013

Keywords:WaspVespulaRespiration patternsVentilation movementResting metabolismTemperature

0022-1910 � 2013 Elsevier Ltd.http://dx.doi.org/10.1016/j.jinsphys.2013.01.012

⇑ Corresponding authors. Tel.: +43 316 380 5705; fE-mail addresses: [email protected] (H.

(H. Kovac), [email protected] (A. Stabe

Open access under CC

We investigated the respiration patterns of wasps (Vespula sp.) in their viable temperature range (2.9–42.4 �C) by measuring CO2 production and locomotor and endothermic activity.

Wasps showed cycles of an interburst–burst type at low ambient temperatures (Ta < 5 �C) or typicaldiscontinuous gas exchange patterns with closed, flutter and open phases. At high Ta of >31 �C, CO2 emis-sion became cyclic. With rising Ta they enhanced CO2-emission primarily by an exponential increase inrespiration frequency, from 2.6 mHz at 4.7 �C to 74 mHz at 39.7 �C. In the same range of Ta CO2 releaseper cycle decreased from 38.9 to 26.4 ll g�1 cycle�1. A comparison of wasps with other insects showedthat they are among the insects with a low respiratory frequency at a given resting metabolic rate(RMR), and a relatively flat increase of respiratory frequency with RMR.

CO2 emission was always accompanied by abdominal respiration movements in all open phases and in71.4% of the flutter phases, often accompanied by body movements. Results suggest that resting waspsgain their highly efficient gas exchange to a considerable extent via the length and type of respirationmovements.

� 2013 Elsevier Ltd. Open access under CC BY-NC-ND license.

1. Introduction CO2 reaches a trigger threshold, a massive amount exits from the

Insects may vary stupendously in their modes of gas exchange(Gibbs and Johnson, 2004), both among (Hadley, 1994; Lighton,1996; Sláma, 1999; Terblanche et al., 2008c) and within species(Chown et al., 2002; Irlich et al., 2009; Kuusik et al., 2004; Maraisand Chown, 2003), and even within the same individual (Chown,2001; Kovac et al., 2007; Snelling et al., 2012). One particular res-piration pattern in both flying and flightless insects is well knownas discontinuous gas exchange cycle (DGC, for reviews see Chownet al. 2006b; Lighton, 1996; Sláma, 1988). Many insects show thispattern when at rest, at least at the lower to medium temperaturesof their thermal range. Typical DGCs consist of a closed or constric-tion phase with spiracles shut and little to no external gas ex-change (Bridges et al., 1980). O2 inside the insect is metabolized,while CO2 accumulates in the tracheae and in part is buffered inthe hemolymph. This causes a drop in the total intratracheal pres-sure (Buck and Keister, 1955, 1958; Buck and Friedman, 1958; Hetzet al., 1994). In the following flutter phase single spiracles open andclose rapidly. Gas exchange works here due to convection and dif-fusion. Small amounts of O2 are inhaled to sustain a certain low le-vel of PO2 for a minimum O2 delivery to the insect’s metabolizingtissues (Hetz and Bradley, 2005; Lighton, 1996). The CO2 levelkeeps rising in the hemolymph during the flutter phase, as onlysmall amounts of CO2 are exhaled (Buck, 1958). As accumulated

ax: +43 316 380 9875.Käfer), [email protected]).

BY-NC-ND license.

tracheal system to the environment in the open-spiracle phase(Lighton, 1996; Schneiderman and Williams, 1955). CO2 is as-sumed to act directly at the spiracular muscles, with little centralnervous control (Hoyle, 1961); however, Bustami and Hustert(2000), Bustami et al. (2002) and Woodman et al. (2008) foundcontrary evidence.

Discontinuous gas exchange was hypothesized to be an adapta-tion aimed at minimizing water loss from the tracheae (hygricmodel, Chown, 2002, 2006a; Dingha et al., 2005; Duncan et al.,2002b; Hadley, 1994; Kivimägi et al., 2011; Williams and Bradley,1998; Williams et al., 1998, 2010), though findings by Contrerasand Bradley (2009), Gibbs and Johnson (2004) and Sláma et al.(2007) call into question the universal validity of this model. Otherexplanations suggest that it developed to allow sufficient gas ex-change in subterranean, CO2 rich environments (chthonic model,Lighton and Berrigan, 1995). A combination of these two modelsis the hygric-chthonic hypothesis (Lighton, 1998). An alternativeexplanation suggests that it minimizes oxygen toxicity (Bradley,2000; Hetz and Bradley, 2005). The variation of respiration pat-terns has been well investigated in different species (Basson andTerblanche, 2011; Chown et al., 2006a; Groenewald et al., 2012;Klok and Chown, 2005; Kovac et al., 2007; Nespolo et al., 2007;Terblanche et al., 2008a; Williams et al., 2010). Such an analysisis lacking in vespine wasps. This is especially interesting becauseVespula sp. show an overall higher level and a steeper incline inresting metabolism with increasing ambient temperature (highQ10) than many other insects (see Käfer et al., 2012). In this paper,therefore, we investigated the characteristics of the respiration

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patterns of vespine wasps, Vespula sp., over their entire viable tem-perature range. We compare the specific features of their gas ex-change patterns with other flying and nonflying insects.

Respiration of adult insects is accomplished by a combination ofpassive diffusive gas exchange and active convective ventilation(Jõgar et al., 2011; Lighton, 1996; Terblanche et al., 2008b).Ventilatory movements are usually observed via automated opticalactivity detection. While this technique allows for an easy, semi-quantitative assessment of general activity (Lighton, 2008) it cannotgive information about the nature of an activity event. It cannot dis-tinguish between abdominal pumping and movement of other bodyparts. For a general perspective on the mechanisms of respiration,therefore, we investigated the connection between gas exchangeand respiratory movements in detail by infrared video observation.

2. Material and methods

2.1. Animals

A total of 37 yellow jacket foragers (24 Vespula vulgaris (Lin-naeus 1758) and 13 Vespula germanica (Fabricius 1793)) were bai-ted with sucrose solution at an artificial feeding place and caughtfor immediate analysis (29 individuals) or stored in cages over-night in a dark and cool area (8 wasps, 12–15 �C, sucrose solutionprovided) for use at low temperatures on the next day. As weneeded undisturbed, undamaged individuals for our experiments,species determination had to be accomplished after the experi-ments by assessment of head and thorax color markings, followingthe main characteristics in identification literature (temple, clyp-eus and pronotal markings; see Bellmann, 1995; Brohmer, 1977;Clapperton et al., 1989; Witt, 1998). As color markings are highlyvariable (Clapperton et al., 1989), this proved to be rather difficultin some cases. For example, we had 4 V. germanica individualswhich could be easily taken for V. vulgaris because of their thoracicpronotal markings.

The experiments took place overnight to ensure that the waspswere at rest for long enough periods (especially at high Ta). Theanimals would no longer have shown their natural resting behaviorand could have been physically damaged (especially at higher Tas)had we extended the experimental periods even further. Afterinsertion into the chamber, it took usually at least 90 min beforethe insects had calmed down enough in the measurement chamberto allow analysis of resting respiratory patterns. Individuals hadtime to accustom to a new experimental ambient temperature(Ta) in the respiratory measurement chamber for a minimum of15 min at the lowest temperatures (<10 �C). At medium to hightemperatures we waited at least 30 min before an evaluation wasstarted. Temperatures were set from 2.5 to 45 �C in steps of 2.5or 5 �C. After every change of Ta (ramp), however, it took time foran individual to stabilize in metabolism and behavior. So we hadto optimize the measurement regime in the course of the experi-ments through reduction of tested Tas per individual. The majorityof individuals (23 of 37) were tested at one Ta, six at two Tas, five atthree Tas, two at four Tas, and one individual at five Tas. Each Ta

lasted for 3.5 h minimum.As respiration data did not differ significantly between

V. vulgaris and V. germanica (P > 0.5, ANOVA; see Section 3.2 andTable S1; for metabolism data see (Käfer et al., 2012)), respirationdata were pooled and animals were referred to as Vespula sp. inthis paper (body mass = 0.1019 ± 0.0179 g, N = 37).

2.2. CO2 measurement

Carbon dioxide production of the yellow jackets was measuredin a flow through respirometry setup as described in Käfer et al.(2012), Kovac et al. (2007), Petz et al. (2004) and Stabentheiner

et al. (2012). The test chamber dimensions (volume = 18 ml) al-lowed unhindered movement of the wasps during the experiment.As the wasps stayed in the chamber over long time spans (>6 h,typically overnight) they were also provided with 1.5 M sucrosesolution ad libitum as a food source. Experimental temperaturewas set by an automatically controlled water bath (Julabo F33HT, Julabo Labortechnik GmbH, Seelbach, Germany; temperatureregulation to 0.1 �C). As the temperature inside the test chamberdeviated slightly from that of the water bath we measured the ac-tual experimental temperature (Ta) with a thermocouple inside thechamber, close (<10 mm) to the wasp.

Outside air was led through the reference channel of a differen-tial infrared gas analyser (Advance Optima URAS 14, ABB Analyti-cal, Frankfurt, Germany) sensitized to carbon dioxide, themeasurement chamber and subsequently through the measure-ment channel. Gas flow was set at 150 ml min�1 by a mass flowcontroller (Brooks 5850S; 0–1000 ml/min; Brooks Instrument, Hat-field, USA). This flow allowed for an accurate temporal resolutionas well as for a good CO2 signal in terms of signal to noise peak ra-tio (Gray and Bradley, 2006; Stabentheiner et al., 2012). Carbondioxide production of the tested wasps was recorded at intervalsof 1 s. The measurement gas (i.e. air) was dried via Peltier elementequipped cool traps prior to the reference and measurement chan-nel. Relative humidity in the test chamber was regulated by a set ofhumidifying bottles filled with distilled water, immersed in an-other Julabo water bath adjusted to the desired dew point temper-ature to keep the relative humidity in the measurement chamberat the desired level (50% at 45–15 �C, 60% at 12.5 �C, 70% at10 �C, 80% at 7.5 �C, 90% at 5 �C and 100% at 2.5 �C). Formulas fordew point calculation are given in Stabentheiner et al. (2012).

The empty test chamber was recorded for 5 min before andafter each experiment to determine any initial CO2 signal offsetfrom zero as well as a possible signal drift from the start to theend of the experiment. The long duration of each experiment re-quired regular (3 h intervals) automatic zero- and end point cali-bration of the URAS gas analyser, utilizing internal calibrationgas cuvettes containing a defined concentration of carbon dioxide.

The tube length between measurement chamber and measure-ment channel of the DIRGA resulted in a signal delay that was cor-rected for synchronization of the CO2 trace recordings withinfrared video sequences.

Data analysis and statistics were conducted using custom madepeak and valley finding formulas in Excel (Microsoft Corporation,Redmond, USA), OriginPro 8.5 (OriginLab Corporation, Northamp-ton, USA) and Stathgraphics Centurion XVI (StatPoint TechnologyInc., Warrenton, USA). The amount (ll min�1, ppm) of CO2 produc-tion refers to standard (STPS) conditions (0 �C, 101.32 kPa = 760 -Torr). All gas exchange referred to as respiration in the followingchapters is strictly speaking CO2 emission, as O2 uptake was notmeasured in this setup.

2.3. Behaviour and activity observation

To evaluate the wasps’ behavior and to determine the periodswhen the tested individuals were at rest we applied state of theart infrared thermography techniques that particularly enabledus to distinguish between rest and activity without disturbingthe wasps in their natural behavior (Käfer et al., 2012; Kovacet al., 2007; Stabentheiner et al., 2012).

The top of the measurement chamber was transparent to infra-red (IR) radiation (covered with plastic film permeable in the rangeof 3–13 lm). It enabled us to record both the wasps’ body surfacetemperature and activity with an infrared thermography camera(ThermaCam SC2000 NTS, FLIR Systems Inc., Wilsonville, USA; fordetails see Kovac et al., 2007; Schmaranzer and Stabentheiner,1988; Stabentheiner and Schmaranzer, 1987; Stabentheiner et al.,

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H. Käfer et al. / Journal of Insect Physiology 59 (2013) 475–486 477

2012). Not only visual clues (e.g. body movements), but also thethermal state of the individual (ectothermic or endothermic) couldbe evaluated. This thermal state was determined by the differencein thoracic and abdominal surface temperature (Tth � Tab). An indi-vidual was assessed as resting when it was ectothermic (Tth � Tab)and showed no or only scarce body movements for a minimumtimespan of 10 min (see classification according to Crailsheimet al., 1999; Stabentheiner and Crailsheim, 1999; Stabentheineret al., 2003); single flips of legs or antennae were allowed (compareKaiser, 1988). At higher Ta (>27.6 �C) the duration was reduced to5 min if no 10 min sections were available. In the course of evalu-ation we had to redefine ‘‘rest’’ in such a way that individuals notmoving for a longer period of time were allowed to show weakendothermy (Tth � Tab < 2 �C, usually <1 �C) over a few periods inthe experiment (see Käfer et al., 2012; Kovac et al., 2007).

IR sequences were recorded on hard disk at 3, 5 or 10 Hz. Anal-ysis of the yellow jackets body surface temperatures was con-ducted with AGEMA Research software (FLIR Systems Inc.,Wilsonville, USA) controlled by a proprietary Excel (Microsoft Cor-poration, Redmond, USA) VBA macro.

2.4. Respiration frequency and abdominal ventilation movements

A respiration cycle was determined from one minimum in CO2

emission just before the open phase to the next one. For discontin-uous gas exchange cycles (DGCs) this included a closed and a flut-ter phase. In cyclic respiration at higher temperatures the samescheme applied. From minimum emission to minimum emission,every CO2 peak was assumed to be a respiration cycle. Abdominalventilation movement (pumping, etc.) was assessed from IR videosequences recorded at a frequency of 10 Hz. A minimum of 10 res-piration cycles were assessed in the evaluation of respirationmovements, resulting in time spans of 13 min at the highest Ta

(36.3 �C) and 287 min at the lowest Ta (5.9 �C) tested. The abdomenhad to be well distinguishable from the background over this per-iod of time.

Respiratory ventilation within one cycle consisted of one or sev-eral successions of single abdominal pumping movements. Thesesuccessions were counted as single ventilatory events. The dura-tions of these ventilatory events were determined, and related tothe whole cycle as well as the cycle phases (open, closed or flutter).

3. Results

As we tested two species of vespine wasps, Vespula vulgaris andV. germanica, we had to analyze our data regarding the possibilityof inter-species differences in respiration parameters. ANOVA re-vealed no influence of the tested wasp species on respiration cycleduration (P = 0.5449, F-quotient = 0.39, DF = 1) and CO2 release percycle (P = 0.9239, F-quotient = 0.01, DF = 1; see Supplementarymaterial, Table S1; data for the two species in Table S2). Therefore,species was not considered for the further analysis.

3.1. Respiration (CO2 release) pattern

Over the entire temperature range, spiracle control was func-tioning well for Vespula sp. At the lowest experimental tempera-tures (Ta = 2.9 �C) yellow jackets showed discontinuous gasexchange resembling an ‘‘interburst–burst’’ pattern similarly tothat described by Marais and Chown (2003) for Perisphaeria sp.cockroaches. Interburst phases with a minimum of 0.6 and a max-imum of 81.73 min duration (mean: 11.86 ± 12.05 min) were fol-lowed by 0.42–14.57 min long burst phases (mean:6.19 ± 4.91 min) consisting of 1–5 initial higher peaks and severalsubsequent lower ones (see Fig. 1A). A flutter phase could not be

observed at this Ta. Sporadic single CO2 spikes with similar peakheight and duration as the initial peaks of the burst phases werecounted as separate open phases. They caused the rather high SDin duration of closed as well as open phases (Fig. 3).

With increasing Ta DGC appeared in a more common fashionwith a closed phase followed by a distinct flutter phase and themain peak or open phase (Fig. 2A). The open phase oscillations ofthe CO2 signal merged (but remained detectable), and flutter be-came visible (Fig. 1B and C). At temperatures of 15–25 �C Vespulasp. showed typical DGC patterns (Hetz and Bradley, 2005; Lighton,1996) with closed, flutter and open phase (Fig. 2B). Exceptionalbody movements, e.g. when the wasp lost and regained hold witha leg or flipped the wings (Fig. 1B–D, arrows) were clearly distin-guishable from the ‘‘normal’’ respiration pattern. At Ta = 26.2 �Cthe CO2 level inside the measurement chamber did not alwaysreach zero between two respiration cycles. However, CO2 emissionbefore the open phase resembled a flutter pattern consisting ofmerging single peaks. In certain individuals, residues of this partic-ular pattern could be observed in some cycles as slight increases inthe CO2 signal prior to the main respiratory peak even at Ta = 31.4and 36.4 �C (Fig. 2B, D; see large triangles in Fig. 3). At Ta > 31.4 �Call individuals showed cyclic respiration (Fig. 2C). At the highestexperimental temperatures (Ta = 39.7 and 42.4 �C), resting periodswere scarce in yellow jackets. CO2 emission was always cyclic,sometimes on the verge of continuous respiration (Fig. 2D).

Fig. 3 shows the duration of cycles, and of open, closed and flut-ter phases (where present) as a function of experimental ambienttemperature. The course of all components of DGC follows expo-nential curves. With rising ambient temperature the open phasedecreased slower in duration than the flutter and the closed phasesat low to medium Ta. Closed phases were only detectable up to Ta -6 26.3 �C. Fig. 4 shows the duration of the respiration cycles andcycle phases in dependence on resting metabolic rate (RMR). How-ever, the courses of data points indicate a higher order of depen-dence than a simple exponential decrease. Good linear regressionin a double logarithmic graph (inset) strengthens this finding.

3.2. Respiration frequency and CO2 release per cycle

With rising Ta the cycle frequency (f) increased (Fig. 1, Fig. 2)following an exponential curve (Fig. 5). Data fitted best with anexponential function of the type f = y0 + A1Ta/t1, withy0 = 0.12716, A1 = 2.18932, t1 = 11.2997 (R2 = 0.51337, P < 0.0001,N = 37). Respiration cycle frequency was 2.55 ± 3.58 mHz at4.7 �C, 9.33 ± 13.2 mHz at 9.8 �C, 13.0 ± 24.66 mHz at 19.8 �C,39.92 ± 25.35 mHz at 31.1 �C and 73.97 ± 28.85 mHz at 39.7 �C.Data at 42.4 �C were not included in the fitting curve because sin-gle CO2 ‘‘peaks’’ merged to ‘‘plateaus’’. Comparison of variances ofcycle frequency at the same Ta revealed significant differences be-tween individuals (P < 0.05, N = 2–10, ANOVA). Over the entiretemperature range these tests indicated significant differences in69.5% of comparisons.

An ANOVA with the means per animal and Ta (of both species)indicated a slight negative temperature dependence of CO2 releaseper cycle (P < 0.05; R2 = 0.06685, N = 62, F = 5.36977, DF = 60). Thecorrelation was more pronounced in an analysis with all cycles ofall animals, which includes the intra-individual variation (Fig. 6).CO2 release per cycle as estimated from the regression line chan-ged from 39.51 ll g�1 cycle�1 at 2.9 �C to 25.4 ll g�1 cycle�1 at42.4 �C,

Single individuals compared at the same temperature showedsignificant differences in the variances of mean CO2 emission percycle and animal (P < 0.05, N = 2–8, ANOVA; see large circles inFig. 6). Over the entire temperature range these within-Ta compar-isons showed inter-individual differences in 56.8% of cases. Thisimplies that the other 43.2% of cases indicated no difference.

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A B

C D

Fig. 1. Discontinuous gas exchange (DGC) of resting wasps at ambient temperatures (Ta) <20 �C. CO2 release changes characteristically in pattern as well as in frequency withrising Ta (A–D). Arrows mark CO2 release exceptional in peak height (B) or pattern (C and D), caused by bodily activity (e.g. in (B) the insect lost and regained grip on the filmcovering the experimental chamber for two times). Dotted lines indicate mean CO2 emission over timespan. Insets show details in the CO2 registration.

478 H. Käfer et al. / Journal of Insect Physiology 59 (2013) 475–486

However, measurements where data of only one individual couldbe evaluated indicate also considerable intra-individual variance(Fig. 6, Ta = 22.5 and 42.4 �C). In direct comparison, wasps differedfrom honeybees significantly in slope and intercept (P < 0.0001 inboth cases, ANOVA; see Fig. 6).

Cycle frequency (f) increased linearly with the mass specificRMR (Fig. 7, f (mHz) = �2.54647 + 0.65394 � RMR CO2 (ll g�1

min�1), R2 = 0.976, P < 0.0001, N = 37, means per animal). A com-parison with other (flying and non-flying) insect species revealedyellow jackets to be among the insects with the lowest respiratoryfrequency at a given RMR though Vespula has a rather high mass-specific RMR (Käfer et al., 2012). This comparison also showed thatthis relation differs largely between different insect species (Fig. 7).However, in spite of the high variation in RMR levels as well as inslopes of the single species data, a tendency is obvious in insects toincrease respiration frequency with an increase in emission of CO2.

3.3. Respiration movements

CO2 emission of wasps at rest was accompanied by convectiveabdominal respiration movements (pumping, etc.) in all observedcases (100%) where CO2 emission took place, during discontinu-ous as well as during cyclic respiration. Respiratory ventilationconsisted of a succession of single abdominal pumping move-ments (see Supplementary material, IR video S3). Such a succes-sion was counted as one single ventilatory event. However,typical abdominal ventilation movements were often accompa-nied by leg or antenna movement, flipping of the wings (see Sup-plementary material, IR video S4) as well as sideward jerking ofthe abdomen, leading to spasm-like twisting of the whole waspbody (24.2% over the tested temperature range; for details see Ta-ble 2, Fig. 8). Additional body movements, therefore, contributedto a considerable amount to respiration movements. During a

DGC, some kind of respiration movement could be observed inall open phases and also in 71.4% of the flutter phases (66.7% ifthe distinct increase in the CO2 signal before an open phase at Ta -P 26.3 �C was not counted as a flutter phase). Ventilation move-ments during flutter were in the majority of cases single or fewabdominal movements with small amplitude often accompaniedor masked by body movement. They differed visibly from thewasps’ pumping in open phases. Fig. 8A shows the percentagedistribution of abdominal respiration movements (resp), abdomi-nal respiration movements accompanied by leg and antennamovements (resp&mov), and body movements possibly maskingrespiration movements (mov) in closed, flutter and open phases.All types of movement occurred in all phases of respiration,though at some Tas some types were missing. Abdominal respira-tion movements (pumping) were in all tested individuals accom-panied by other body movements in at least one phase of arespiration cycle. Whole-body movements possibly masking theabdominal ventilation movements (mov; see Table 2 and Supple-mentary material, IR video S5) were rather rare. They occurred in9.7% of the cycles (over the tested temperature range), in closedas well as in flutter and open phases. Fig. 8B shows the relativeamount of ventilation movements (resp, resp&mov, mov) in theclosed, flutter and open phases of respiration cycles. In the openphase of the gas exchange cycle clearly definable respirationmovements (resp and resp&mov) were observed at all Tas. Inthe flutter and closed phases, however, this did not always occur.Including those body movements that might have maskedabdominal pumping (mov) did not change this resultconsiderably.

Abdominal movements did also occur in closed phases (see alsoGroenewald et al., 2012; Hetz et al., 1994; Jõgar et al., 2011). Themovements resembled abdominal respiration movements asobserved in flutter phases (without additional leg or body

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Fig. 3. Duration of cycles, flutter, open, and closed phases (where existent; meanvalues with SD). Open phase values are 100% in every cycle (i.e. no cycle withoutCO2 emission). Flutter data points at Ta > 30 �C (large triangles) represent respira-tion patterns resembling flutter phases without preceding closed phases (seeSection 4).

A B

C D

Fig. 2. Representative CO2 release patterns of resting wasps at ambient temperatures (Ta) >20 �C. (A) Typical DGC pattern with closed (C), flutter (F) and open (O) phases. (B)DGC on the verge of cyclic respiration. No closed phases (i.e. CO2 trace reaches zero), and flutter phases merge with open phases. (C) and (D) Cyclic respiration. Dotted linesindicate mean CO2 emission over timespan.

Fig. 4. Duration of cycles and flutter, open, and closed phases (where existent;mean values with SD) as a function of resting metabolic rate (RMR). Large trianglesrepresent the same respiration patterns as described in Fig. 3. Inset shows data withlogarithmic scaling on both axes. Regression lines follow log10 RMR VCO2 (ll g�1 -min�1) = a + b � log10 Duration (s). Cycle: a = 3.6982, b = �1.221, R2 = 0.93908; Openphase: a = 2.68333, b = �0.73177, R2 = 0.95925; Closed phase: a = 3.19209,b = �1.1129, R2 = 0.64334; Flutter phase: a = 3.47489, b = �1.09373, R2 = 0.87907.

H. Käfer et al. / Journal of Insect Physiology 59 (2013) 475–486 479

movement), but were not accompanied by CO2 emission. Withincreasing Ta the total duration of abdominal ventilation move-

ments decreased exponentially (Fig. 9), which coincided with theincrease in cycle frequency reported in Section 3.2 (Fig. 5).

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Fig. 5. CO2 release cycle frequency (fcycle) of resting wasps in dependence onambient temperature (Ta). Squares indicate mean values with SD (vertical bars),numbers indicate evaluated cycles/individuals. Data fit best with the indicatedexponential function f = y0 + A1Ta/t1, with y0 = 0.12716, A1 = 2.18932, t1 = 11.2997;R2 = 0.51337, P < 0.0001, N = 37. Data at 42.4 �C were not included into the fittingcurve because single CO2 peaks merged. Solid line shows honeybee data from(Kovac et al., 2007).

480 H. Käfer et al. / Journal of Insect Physiology 59 (2013) 475–486

CO2 emission per cycle correlated positively with the durationof abdominal ventilation movements if calculated throughout allexperiments (Fig. 10, F = 0.6211, P < 0.0001, N = 9). However, linearregression in 5 of 9 wasp individuals showed insignificant results,probably due to low variation of duration (compare inset inFig. 10). Slopes of the individual wasps’ regression lines(F = 0.07872, P = 0.78715, N = 9) as well as y-intercepts(F = 0.35149, P = 0.10295, N = 9) did not change significantly withTa.

Fig. 6. CO2 release per cycle in resting wasps (small circles, overlapping values shifted hrelease values of single individuals. At Ta = 42.4 �C, cycles were identifiable only in one ouWhiskers indicate 1.5 interquartile range (IQR, def. Tukey). Numbers indicate the cyc� 0.35678 � Ta(�C), R2 = 0.02998, P < 0.0001, n = 5336 cycles, 34 animals. Solid line shohoneybees significantly in intercept (F-quotient = 4731.92, P < 0.0001, ANOVA) as well as(Kovac et al., 2007)).

4. Discussion

4.1. Respiration patterns

At rest, many insect species show a particular respiration pat-tern of discontinuous gas exchange cycles (DGC; for review seeChown et al., 2006a; Lighton, 1996; Sláma, 1988). The illustrationof respiration patterns depends on flow rate, measurement cham-ber size (i.e. volume) and metabolic rate of the animal (Gray andBradley, 2006; Lighton, 2008; Terblanche and Chown, 2010). Alarge measurement chamber dilutes the animal’s CO2 trace, leadingto a smoothed away signal at the CO2 detector. Last but not least,the metabolic turnover of the tested animal is a crucial parameter(Gray and Bradley, 2003; Moerbitz and Hetz, 2010). In restingyellow jackets the CO2 emission varied in a wide range, from5.6 ll g�1 min�1 at 7.7 �C to 101.3 ll g�1 min�1 at 40 �C (Käferet al., 2012). With a measurement chamber size of 18 ml –as smallas possible, but without impairing the animal’s natural movement– and a flow rate set to 150 ml min�1 the respiration patterns ofVespula sp. could be displayed throughout their entire viable tem-perature range.

Typical DGCs consist of a closed phase with shut spiracles andno external gas exchange (Bridges et al., 1980) followed by a flutterphase with the spiracles opened in close succession, and the openspiracle phase (Hetz and Bradley, 2005; Lighton, 1996). At the low-est experimental temperatures (Ta = 2.9 �C), DGC resembled aninterburst–burst pattern similar to that described by Marais andChown (2003) for Perisphaeria sp. cockroaches and Duncan andDickman (2001) for Cerotalis sp. beetles. In Vespula sp. long inter-burst (closed) phases alternated with long open burst phases con-sisting of single peaks which sometimes tended to merge at theend of the open phase (Fig. 1A), resembling to some degree ‘‘re-versed’’ flutter phases. This seems to suffice in exchanging CO2

and O2 at this overall low level of metabolic rate (Fig. 1A; Hetz,2007; Käfer et al., 2012; Moerbitz and Hetz, 2010). Nevertheless,spiracle control functioned well at this lowest experimental ambi-ent temperature. Honeybees, in comparison, fall into chill coma at

orizontally) at different ambient temperatures (Ta). Large circles indicate mean CO2

t of four individuals. The Boxplot shows Q1, Q2, Q3 and mean values (black squares).les / individuals tested. Linear fit (dotted line) is CO2 (ll g�1 cycle�1) = 40.56272 -ws honeybee data from (Kovac et al., 2007) for comparison. Wasps differ fromin slope (F-quotient = 485.64, P < 0.0001, ANOVA; data of honeybees by courtesy of

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Fig. 7. Respiration cycle frequency (f) as a function of resting metabolic rate (RMR).Comparison of wasps (1, full squares) with literature data from several insectspecies, e.g. Culiseta inornata (3, unfilled tipped squares), Glossina palpalis (19,circles, top filled), G. brevipalpis (20, circles, filled right), G. austeni (21, circles, filledleft), G. morsitans centralis (22, circles, bottom filled), Apis mellifera (25, full circles),Cratomelus armatus (27, circles), Rhodnius prolixus (28, triangles). Numbers corre-spond with those in Table 1. At low RMR data points overlap strongly; for data of allspecies see Table 1.

A

B

Fig. 8. (A) Distribution of abdominal ventilation movement (resp), ventilation andleg/antenna movement (resp&mov) and body movement possibly masking respi-ration movements (mov; see Table 2) over the phases of respiration cycles. (B)Amount of ventilation movement (resp, resp&mov, mov) in the phases ofrespiration cycles. In the right half of the bars body movement possibly maskingrespiration movements (mov) was excluded from the analysis. Flutter phases atTa > 30 �C represent respiration pattern resembling flutter phases without preced-ing closed phase (see discussion). Quantities of tested cycles were 26/13/12/16/19/14/28/74 from low to high Ta.

Fig. 9. Total duration of abdominal ventilation movements in dependence onambient temperature (Ta). Each data point represents an individual wasp (meanswith SD, numbers show tested respiration cycles). Data at 6.2 �C were not includedinto the fitting curve due to high SD (see Text, Section 3.1).

Fig. 10. Co2 emission in dependence on duration of abdominal ventilationmovements. Data points represent mean values of resting wasp individuals withtheir SDs. Numbers near the data points show number of tested respiration cycles.The inset shows data from individuals at 6.3 �C (a), 22.5 �C (b) and 26.3 �C (c).

H. Käfer et al. / Journal of Insect Physiology 59 (2013) 475–486 481

Ta � 10 �C and, losing control over their spiracles, emit CO2 contin-uously (Kovac et al., 2007; Lighton and Lovegrove, 1990; compareFree and Spencer-Booth, 1960).

With rising Ta, wasp DGC had closed phases and distinct flutterphases as found in many other resting insects (e.g. Chown and

Davis, 2003; Hadley, 1994; Hetz and Bradley, 2005; Lighton,1996; Lighton and Lovegrove, 1990; Sláma, 1999; Vogt and Appel,1999, 2000). Open phases consisted of consecutive merging and inamplitude diminishing peaks at Tas of about 6–16 �C (Figs. 1B and2A). The typical DGC pattern with closed, flutter and open phaseappeared more and more distinctly (Fig. 2B).

With rising Ta, the DGC patterns changed in a way that theclosed and flutter phases diminished in duration and then succes-sively vanished entirely (Fig 3). This result was in accordance to thefindings of Contreras and Bradley (2010) in Rhodnius prolixus andGromphadorhina portentosa, which showed that metabolic rate af-fects spiracle activity, which may be an explanation for the differ-ent patterns of gas exchange in one species at differenttemperatures. At Ta � 27.5 �C, 50% of the cycles showed flutterand closed phases (see Supplementary material, Table & Fig. S5).Closed phases ceased between 26.2 and 31.1 �C (i.e. atTa = 31.1 �C no closed phases were detectable; see Fig. 3;

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Table 1Mass specific resting metabolic rate (RMR), respiration frequency (f) and cycle duration (s) data from this study and literature data. Ta = experimental ambient temperature; Nind. = number of individuals; n = number of respiration cycles (where available).

No. Species Ta (�C) N ind. n f SD Duration SD RMR CO2 SD References

(mHz) (s) (ll g�1 min�1)

1 Vespula sp. 2.9 2 130 0.99 0.99 1062.22 826.24 2.31 1.71 This study and5.8 4 80 1.21 1.12 887.60 587.04 3.79 3.16 Käfer et al. (2012)7.7 4 59 2.40 2.28 538.57 466.40 5.62 3.92

10.0 6 87 3.03 3.14 440.86 381.05 8.38 5.4412.0 3 62 2.31 1.36 402.33 289.80 7.87 3.4215.1 4 60 4.53 2.66 194.58 141.87 9.96 2.7816.2 6 79 6.08 2.55 173.24 121.11 13.28 2.9819.8 5 40 6.08 3.77 237.67 120.97 16.06 5.7222.6 1 41 4.19 1.79 91.78 63.07 18.37 3.7126.3 7 74 11.40 5.09 37.29 29.17 27.62 5.9831.1 7 56 28.66 12.87 25.66 14.63 38.86 5.1735.8 6 43 38.91 14.90 16.15 10.71 60.93 5.8139.7 4 31 62.69 15.39 25.19 36.24 101.30 10.17

2 Blattella germanica 10.0 19 1.48 0.06 675.68 15.60 8.20 1.37 Dingha et al. (2005)2a Blattella germanica 10.0 12 1.20 0.04 833.33 26.40 6.48 1.233 Culiseta inornata 10.0 4 10.00 0.50 100.00 ⁄ 6.48 0.53 Gray and Bradley (2006)

20.0 3 18.33 1.17 54.56 ⁄ 30.10 2.2130.0 3 41.67 2.50 24.00 ⁄ 59.70 1.55

4 Zophosis complanata 25.0 9 4–90 1.59 0.41 676.20 207.60 2.87 0.82 Duncan and Byrne (2002)5 Zophosis punctata 25.0 10 4–90 1.86 0.77 621.00 231.60 3.25 1.276 Pimelia canescens 25.0 2 4–90 1.20 ⁄ 878.40 ⁄ 1.90 0.007 Pimelia grandis 25.0 6 5–17 1.24 0.29 14.10 3.23 1.62 0.15 Duncan and Byrne (2002)8 Onymacris multistriata 23.0 8 20–55 1.79 0.46 587.40 126.00 4.25 0.96 Duncan (2003)9 Pachylomerus femoralis 25.0 5 0.73 0.30 1384.62 ⁄ 1.64 0.22 Duncan and Byrne (2005)10 Scarabaeus gariepinus 25.0 7 0.33 0.20 3000.00 ⁄ 1.34 0.6611 Scarabaeus striatum 25.0 6 0.42 0.20 2400.00 ⁄ 1.34 0.4212 Circellium bacchus 23.5 6 63 0.26 0.05 3829.79 ⁄ 0.86 0.30 Duncan and Byrne (2002)13 Cerotalis sp. 20.0 7 2to10 0.63 0.42 2103.60 1084.20 1.30 0.83 Duncan and Dickman (2001)

25.0 11 2to10 1.54 0.49 712.20 246.60 2.01 0.7530.0 5 2to10 2.11 0.84 537.00 211.20 2.10 0.8735.0 8 2to10 2.72 0.63 381.60 70.20 2.48 0.5540.0 6 2to10 4.90 1.42 228.00 99.60 3.51 0.68

14 Carenum sp. 20.0 3 2to11 0.56 0.10 1816.80 351.00 1.06 0.32 Duncan and Dickman (2001)25.0 11 2to12 1.22 0.52 928.80 310.20 1.36 0.2530.0 6 2to13 2.14 0.40 478.80 77.40 1.92 0.5335.0 3 2to14 2.89 0.62 357.00 76.80 2.92 1.53

15 Sysiphus fasiculatus 20.0 6 2.22 0.80 503.40 180.00 4.60 0.98 Duncan and Byrne (2000)16 Scarabaeus rusticus 20.0 6 5.00 3.10 288.00 186.00 2.53 1.5317 Anachalcos convexus 20.0 5 6.45 3.60 211.80 132.00 1.85 0.5318 Scarabaeus flavicornis 20.0 12 1.86 1.40 778.80 420.00 2.62 0.6819 Glossina palpalis 15.0 13 26.29 3.91 38.04 ⁄ 5.82 8.86 Basson and Terblanche (2011)

20.0 36.69 10.15 27.26 ⁄ 9.94 18.4725.0 52.19 4.24 19.16 ⁄ 21.20 35.4530.0 52.12 32.36 19.19 ⁄ 51.38 181.8635.0 36.04 0.00 27.75 ⁄ 40.92 80.04

20 Glossina brevipalpis 15.0 21 37.45 13.15 26.70 ⁄ 7.40 13.2720.0 39.09 15.15 25.58 ⁄ 15.44 18.24

21 Glossina austeni 15.0 14 44.88 22.28 6.30 9.7720.0 62.20 12.68 16.08 ⁄ 12.52 16.9725.0 91.67 10.91 19.84 23.5135.0 71.42 14.00 28.55 41.03

22 Glossina morsitans 20.0 56 56.76 4.03 17.62 ⁄ 9.90 1.05 Terblanche and Chown (2010)centralis 24.0 56 62.66 4.80 15.96 ⁄ 13.61 1.65

28.0 56 67.47 5.88 14.82 ⁄ 18.87 2.0532.0 56 70.85 6.31 14.11 ⁄ 24.30 2.07

23 Aphodius fossor 15.0 10 1.15 0.14 999.40 115.40 2.75 0.12 Chown and Holter (2000)24 Bombus terrestris 24.0 6 1.44 0.16 694.44 ⁄ 6.17 0.70 Karise et al. (2010)25 Apis mellifera 14.1 5 166 8.69 6.03 115.01 165.90 7.94 7.19 Kovac et al. (2007)

18.2 5 180 11.72 5.14 85.31 194.48 11.01 6.5822.4 5 101 13.83 3.97 72.32 251.96 12.27 2.7826.5 4 167 22.27 4.81 44.91 207.76 18.10 3.1430.4 4 127 33.54 11.27 29.81 88.73 25.00 3.6234.5 6 217 31.84 10.55 31.41 94.80 35.36 4.6038.1 6 177 50.97 10.23 19.62 97.76 41.37 4.96

26 Aquarius remigis 10.0 1 0.83 ⁄ 1233.80 406.30 2.44 0.49 Contreras and Bradley (2011)20.0 1 2.78 ⁄ 295.77 121.22 6.80 0.86

27 Cratomelus armatus 15.0 35 8.33 120.05 35.38 4.64 Nespolo et al. (2007)20.0 35 11.11 90.01 110.26 14.0625.0 35 25.00 40.00 92.16 18.00

28 Rhodnius prolixus 15.0 10 1.25 0.59 76.23 74.70 1.08 0.07 Contreras and Bradley (2010)25.0 10 4.17 ⁄ 23.84 5.11 2.50 0.0335.0 10 20.83 ⁄ 4.88 1.44 5.64 0.29

Cells marked with ⁄ indicate missing SD due to calculation of mean duration from mean frequency literature data or vice versa, or because data was measured from figurespublished in literature.

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Table 2Occurrence of individual wasps’ abdominal movement during respiration cycles in per cent. Cycles were split in the three phases of discontinuous gas exchange (see Fig. 2B). Alltypes of movement may occur in the same phase. Example: All (100%) flutter phases at Ta = 16.5 �C show abdominal movement (resp) as well as phases with abdominal andantenna/leg/wing movement (resp & mov) and 33.3% showed body movement (mov). Ta indicates the ambient temperature, n is the number of cycles tested. Abdominalmovement in closed phases – though resembling movement in flutter phases – did not concur with CO2 release.

Ta (�C) Flutter (%) Open (%) Closed (%) n

resp resp & mov mov resp resp & mov mov resp resp & mov mov

6.2 84.2 5.3 15.8 68.2 9.1 59.1 5.0 – 10.0 2611.8 – – 87.5 61.5 7.7 76.9 – – – 1314.4 – 11.1 22.2 75.0 33.3 83.3 9.1 9.1 27.3 1216.5 100 100 33.3 81.3 93.8 43.8 23.1 30.8 15.4 1619.8 87.5 43.8 81.3 94.7 26.3 84.2 46.2 – 23.1 1922.5 – – 25.0 85.7 – 14.3 – –– 9.1 1431.1 75 – – 82.6 – 47.8 – –– – 2835.8 – – – 68.9 45.9 48.6 – – – 74

resp = abdominal (respiration) pumping movement.resp & mov = abdominal (respiration) movement and concurrent movement of e.g. antennae, legs, wings.mov = movement of the whole wasp, possibly masking abdominal (respiration) movements.Flutter data at 31.1 �C is marked grey because flutter phase at this Ta merged with open phase (see Section 4).

H. Käfer et al. / Journal of Insect Physiology 59 (2013) 475–486 483

Supplementary material, Table & Fig. S5). In R. prolixus, Contrerasand Bradley (2010) still observed closed phases at Ta = 35 �C. Ithas to be kept in mind that they determined this relationship ina different experimental procedure, exposing insects to a tempera-ture ramp while our insects were exposed to constant tempera-tures. A rough estimation of the cease temperature of closedphases can be done by determining the quotient of cycle to openphase duration (QC/O). We calculated a best fit curve of the QC/O

from the quotients of the original cycle and open-phase durationvalues. At a QC/O of 1, the open phase was as long as the respirationcycle, and the closed phase had vanished. This occurred at a tem-perature of 36.8 �C. This value corresponded almost exactly withthe one determined from the best-fit curves for cycle and openphase duration in Fig. 3, which was 36.7 �C. Flutter phases ceasedbetween 35.8 and 39.7 �C (see Fig. 3, Supplementary materialS6). The fusion frequency of cycles should depend to a considerabledegree on the relation between (basal) metabolic rate and CO2 buf-fer capacity of an insect. A prediction of Hetz (2007) suggests thatDGCs should mainly occur in insects with large differences in met-abolic rate due to changing temperatures or in insect species withhuge spiracular conductance due to short-time high metabolic de-mands (e.g. during endothermy or flight). This applies to wasps(Käfer et al., 2012; and own unpublished measurements). Theirrather high fusion frequency (despite a high RMR), therefore, sug-gests a high CO2 buffer capacity.

As RMR increases with Ta, the curve progression of cycle dura-tion vs. Ta (Fig. 3) seems similar to that in cycle duration vs. RMR(Fig. 4). However, while in the former case the curves are best de-scribed by the mentioned exponential functions, analysis of the lat-ter revealed a higher order of dependence than a simpleexponential growth. Good linear regression in dual logarithmicscaling (Fig. 4, inset) backs this finding. Due to high intra- and in-ter-individual variation in gas exchange pattern, neither switchedall wasps from one pattern to another at the same experimentaltemperature, nor did they always show the same pattern at thesame Ta. Such variation was also observed in the cockroach Perisp-haeria sp. by Marais and Chown (2003) and in several beetle spe-cies of southern Africa by Chown (2001).

It is discussed that opening an insect’s spiracles for extendedperiods leads to critical tracheal water loss in dry environments(Chown et al., 2006a; Dingha et al., 2005; Duncan and Byrne,2000; Duncan et al., 2002a,b; Hadley, 1994; Kivimägi et al., 2011;Williams et al., 1998, 2010; Williams and Bradley, 1998). Contraryfindings question this hypothesis (Contreras and Bradley, 2009;Gibbs and Johnson, 2004). An alternative model suggests that pos-sible O2 intoxication caused by high partial O2 pressure in the tra-

cheal system is a key parameter which forced development ofdiscontinuous gas exchange (Hetz and Bradley, 2005). In any case,the amount of accumulated CO2 is the trigger for the opening ofspiracles (Lighton, 1996; Schneiderman and Williams, 1955). Withrising Ta, and resulting increase in RMR, yellow jackets have tobalance spiracle opening, O2 ingress and CO2 emission. Short, fastopenings (i.e. flutter) accompanied by single, small-scale abdomi-nal ventilation movements could maintain a sufficient PO2 insidethe wasp for longer periods (see Förster and Hetz, 2010), until ithas to get rid of CO2 in a comparably short, huge burst, concur-rently inhaling O2. This allows for the following closed phase withno or little O2 uptake and CO2 emission and tracheal water loss.When the CO2 level reaches a certain threshold, the cycle startsanew. However, this works only up to a certain temperature andtherefore metabolic rate. As reported by Chown and Nicolson(2004) and Contreras and Bradley (2010), with increasing ambienttemperature, duration of the closed phase becomes shorter andshorter first, and in succession the flutter phase vanishes. In Vesp-ula sp., above experimental temperatures of about 30 �C, with ris-ing temperature the CO2 trace increasingly often did not reachzero, which is said to be a criterion of a DGC (Chown et al.,2006b). However, right at the beginning of the open phase, CO2 in-creased in steps at a rate clearly distinguishable from that of themain peak (Fig. 2B, e.g. at 7, 12 and 18 min). Probably, these aresingle peaks of a flutter phase, below the temporal resolution ofour measurement setup and therefore forming a graduated slope.In our opinion these graduated slopes are flutter phases mergingwith the consecutive open phases (Fig. 3, large triangles; Table 2,marked data). We suppose that this represents DGC on the vergeof cyclic respiration. This resembles findings of Contreras and Brad-ley (2009) on R. prolixus. At temperatures higher than 36 �C, openphases of wasps occurred in such close succession that the peaksmerged at the base and the CO2 signal never reached baseline lev-els. Their metabolic rate was so high that the produced and emit-ted CO2 could not be entirely removed from the measurementchamber before the next pulse was generated. The respiration pat-tern became entirely cyclic (compare Gray and Bradley, 2003).

4.2. Respiration frequency and respiration movements

The wasps’ RMR increases exponentially with rising Ta (seeKäfer et al., 2012)). They respond to the according demand of in-creased gas exchange with a likewise exponential increase in res-piration frequency (Fig. 5) but not with an increasing CO2

emission per respiration cycle (Fig. 6). This was also reported forhoneybees (Kovac et al., 2007) and fire ants (Vogt and Appel,

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484 H. Käfer et al. / Journal of Insect Physiology 59 (2013) 475–486

2000). A comparison over flying and non-flying insect species re-veals a positive correlation of respiration frequency and RMR(Fig. 7, Table 1). In spite of a high variation in level as well as inslope of the single species data, a trend is obvious in insects to in-crease CO2 emission with an increase in respiration frequencyrather than in ‘‘depth of breath’’ or other measures.

In the lower to medium temperature range (Ta = 10–27 �C), rest-ing yellow jackets’ respiration frequency did not differ much fromthat of honeybees (see Fig. 5). The increasing deviation of thecurves above 27.5 �C could result from the exceptional steep in-crease in RMR in yellow jackets compared to honeybees (see Käferet al., 2012). Regarding CO2 emission per respiration cycle, yellowjackets show a slight decrease with Ta similar to honeybees (Kovacet al., 2007; Fig. 6). Because of virtually identical testing arrange-ments in Vespula sp. and Apis mellifera, a straight comparison ofthese two species is possible. At similar respiration frequencies(Fig. 5), resting yellow jackets have a much higher energetic turn-over (see Käfer et al., 2012) and emit CO2 on average in much high-er amounts per cycle (Figs. 6 and 7) than honeybees at similarambient temperatures. Wasps seem to breathe more efficientlywith respect to gas exchange volume per cycle than honeybees.This might base on anatomical (compare Snelling et al., 2011 onLocusta migratoria tracheae), physiological or behavioral differ-ences between the two species. Both are known to have thoracicand abdominal air sacs serving as buffering reservoirs for CO2 la-den exhalation air. These air sacs are documented in anatomicaldrawings by Snodgrass (1985) for honeybees, but to our knowl-edge there is no detailed information for yellow jackets available,and volume data of the tracheal system including the air sacs isavailable neither for honeybees nor for wasps. The insect hemo-lymph serves as a CO2 buffer (Buck and Keister, 1958; Buck andFriedman, 1958; Kaiser, 2002). However, there is also no reportof differences in the buffer capacity of wasp and honeybee hemo-lymph available. Future investigations will have to elucidate thesetopics.

Another explanation might lie in differences in the respirationmovements between yellow jackets and honeybees. Other thanin honeybees, the wasps’ abdominal ventilation movements werenot of a uniform pumping pattern, but often consisted of lateralflipping of the abdomen or single pumps accompanied by wingor leg movement (spasm-like; see Supplementary material, IR vi-deo S4). These body movements might contribute to the abdominalpumping in discharging tracheas and air sacs of CO2 laden air. Weobserved abdominal ventilation movements in 100% of the openphases. The wasps showed ventilation movements also in 71.4%of the flutter phases (66.7% if the distinct increase in the CO2 signalbefore an open phase above 26.3 �C was not counted as a flutterphase), whereas in honeybees no distinct pumping movementscould be observed (Kovac et al., 2007). For a sufficiently effectivegas exchange of adult insects diffusion is not enough (Hadley,1994). The wasps seem to rely on active ventilation during the flut-ter phase in addition to the open phase (Table 2, Fig. 8). Someabdominal movements did also occur in closed phases (see alsoGroenewald et al., 2012; Hetz et al., 1994). Passive gas influx dur-ing micro openings in the closed phase leads to a gradual abdom-inal elongation in Attacus atlas pupae (Hetz and Bradley, 2005;Hetz, 2007) and Pieris brassicae pupae (Jõgar et al., 2011). Theclosed phase movements observed in yellow jackets resembledthe single small abdominal pumping movements observed in flut-ter phases but were clearly not of the passive type (see Brockwayand Schneiderman, 1967).

Vespula sp. has a high energetic turnover at rest compared to A.mellifera (Käfer et al., 2012). However, the yellow jackets’ respira-tion frequencies are similar to that of honeybees up to ambienttemperatures of about 27.5 �C (see Fig. 5), but with overall higherCO2 emission per cycle (see Fig. 6). Despite their high resting met-

abolic rate (Käfer et al., 2012), wasps are among the insects with arather low respiratory frequency at a given RMR. Variation in databetween insect species is so high that no meticulous conclusionscan be drawn from one species to another. However, a generaltrend to raise CO2 emission with an increase in respiration fre-quency can be seen (Fig. 7).

The amount of CO2 emission per cycle correlated positively withthe duration of abdominal respiration movements (Fig. 10). Waspsreduced the duration of ventilation movements at higher temper-atures (Fig. 9). Total duration of respiration movement events wasup to tenfold longer than in honeybees (42.2 vs. 4.8 s at 20 �C, 27.8vs. 2.3 s at 25 �C; mean values, honeybee data from Kovac et al.,2007). It seems that resting yellow jackets gain their efficient gasexchange to a considerable extent via the length of respirationmovements per respiratory cycle. Therefore, they manage a consid-erably higher RMR (see Käfer et al., 2012) with a similar respirationfrequency as honeybees (see Fig. 4). The high respiration volumeand efficiency might be responsible for the rather high transitiontemperature from discontinuous to cyclic respiration.

5. Conclusion

Despite an overall high level and a steep increase of restingmetabolism with increasing ambient temperature (high Q10), rest-ing yellow jackets maintain DGC at comparably high ambient tem-peratures. They breathe more ‘efficiently’ than other insects,achieving more CO2 emission per respiration cycle at comparablerespiration frequencies.

Abdominal ventilation movements at rest were not uniformpumping movements but also included movements of legs anten-nae and wings, and lateral flipping of the abdomen. Results suggestthat respiration efficiency was increased by long duration of theseventilation movements.

Acknowledgements

The research was funded by the Austrian Science Fund (FWF):P20802-B16, P25042-B16. We greatly appreciate the help withelectronics by G. Stabentheiner and with data evaluation by M.Bodner, M. Brunnhofer, M. Fink, P. Kirchberger, A. Lienhard, L. Mir-wald and A. Settari. We also thank W. Schappacher for his help inclarifying some quirks with data conversion, two anonymousreviewers for helpful comments and the editor D.L. Denlinger.

Appendix A. Supplementary data

Supplementary data associated with this article can be found, inthe online version, at http://dx.doi.org/10.1016/j.jinsphys.2013.01012.

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