Journal of Insect Physiology 56 (2010) 522–528
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Journal of Insect Physiology
journal homepage: www.elsevier.com/locate/jinsphys
Transitions in insect respiratory patterns are controlled by changes
in metabolic rate
H.L. Contreras *, T.J. Bradley
University of California, Irvine, Ecology and Evolutionary Biology Department, 321 Steinhaus Hall, Irvine, CA 92697-2525, United States
A R T I C L E I N F O
A B S T R A C T
Article history:
Received 30 January 2009
Received in revised form 20 May 2009
Accepted 28 May 2009
We examined the respiratory patterns of Rhodnius prolixus and Gromphadorhina portentosa as metabolic
rates varied with temperature to determine whether insects transition from discontinuous (DGC),
cyclical and continuous respiration as a response to increasing aerobic demand. Using flow through
respirometry we: (1) determined the effects of temperature on metabolic rate; (2) objectively defined
periods of spiracular closure; (3) observed whether there was a correlation between metabolic rate and
length of spiracular closure. At low temperatures both species exhibit lengthy periods of spiracular
closure reflecting a discontinuous respiratory pattern. As metabolic rate increased, periods of spiracular
closure decreased and insects displayed a more cyclical pattern of respiration. As metabolic rates
increased even further under the highest experimental temperatures, periods of spiracular closure
decreased even more and a continuous respiratory pattern was employed by both species. Our results
suggest that the three described respiratory patterns in insects are not distinct but are instead a
continuum of respiratory responses driven by the metabolic demand experienced by the insect.
Published by Elsevier Ltd.
Keywords:
Insect respiration
DGC
Temperature
Metabolism
Respiratory pattern
1. Introduction
In insects, gas exchange between the atmosphere and
metabolically active tissues is accomplished via an air-filled,
tracheal system (Buck, 1962; Nation, 2002) that consists of
spiracles, tracheal trunks, tracheae and tracheoles. The spiracles
are external openings that act as valves and are found on the
abdomen and thorax. Internal to the spiracles are large, longitudinal tracheal trunks that subdivide into the smaller tracheae.
Due to repeated branching, tracheal diameter decreases as
distance from the external cuticle increases (Wigglesworth,
1970). The tracheae terminate in the tracheoles, which act as
the principal site of gas exchange with metabolically active cells
(Schmitz and Perry, 1999). Gasses can diffuse very rapidly along
the tracheae since the majority of the tracheal system is filled with
air (Krogh, 1936; Kestler, 1985).
Insects have been found to employ a variety of respiratory
patterns (Lighton, 1996; Williams et al., 1997; Gibbs and Johnson,
2004). The pattern most studied by insect physiologists is the
discontinuous gas exchange cycle (DGC) (Lighton, 1996, 1998;
Chown et al., 2006). During DGC, periods of rapid CO2 release
alternate with interburst periods exhibiting reduced gas exchange
(Levy and Schneiderman, 1966a). Specifically, a classical descrip-
* Corresponding author. Tel.: +1 949 824 7038; fax: +1 949 824 2181.
E-mail address: [email protected] (H.L. Contreras).
0022-1910/$ – see front matter . Published by Elsevier Ltd.
doi:10.1016/j.jinsphys.2009.05.018
tion of discontinuous gas exchange involves three distinct phases:
open, flutter, and closed. During the open phase of the DGC,
spiracular muscles are relaxed and the spiracles are opened
allowing O2 to diffuse into the tracheal system while CO2 diffuses
out into the external environment (Krogh, 1920; Levy and
Schneiderman, 1966a,b). In addition many insects also actively
ventilate the tracheae during the open phase by contracting
abdominal muscles (Buck, 1962; Kestler, 1985; Slama, 1988; Sibul
et al., 2004). These pulsating movements can flush air from one end
of the body to the other through the longitudinal tracheal trunks
and can facilitate the exchange of air between the tracheae and the
external atmosphere (Weis-Fogh, 1967; Slama, 1988, 1999).
Recently it has also been suggested that CO2 may be released by
bulk flow during the open phase (Slama et al., 2007).
Whether gas exchange occurs by diffusion or by bulk flow, the
open phase of DGC is followed by the closed phase. During this
phase, the spiracles are tightly shut preventing any gas exchange
with the external environment. Metabolically active tissues use O2,
which leads to a decrease in intra-tracheal pO2. In addition CO2
accumulates in the insect. Since CO2 is more soluble than O2 in
water, a net negative pressure in the tracheae develops during the
closed phase. When pO2 reaches a critically low level, the spiracles
begin to open and close repeatedly (Schneiderman, 1960). This is
referred to as the flutter phase. During this phase, small amounts of
air enter the tracheal system due to the negative pressure created
in the tracheae. This bulk inflow of air assures O2 enters the system.
However, CO2 continues to accumulate in the tracheae until pCO2
H.L. Contreras, T.J. Bradley / Journal of Insect Physiology 56 (2010) 522–528
reaches a critical level (4–6 kPa) (Levy and Schneiderman,
1966a). Once this level is reached the spiracles are opened
completely and the whole cycle begins once more.
In addition to the DGC, two other respiratory patterns (termed
cyclic and continuous) have been described in insects (Gibbs and
Johnson, 2004). In cyclic respiration, CO2 bursts also alternate with
interburst periods. However, the spiracles are never completely
closed. In continuous respiration there are no bursts of CO2 release.
Instead, the spiracles are maintained open and gas exchange is
relatively constant (Williams et al., 1997; Gibbs and Johnson,
2004). An insect’s neuroendocrine system may be responsible for
controlling the type of respiratory pattern employed (discontinuous, cyclical or continuous) at any time to fit the demands of
external environment or internal physiological conditions (Bustami and Hustert, 2000; Bustami et al., 2002; Slama et al., 2007;
Woodman et al., 2008).
Although the respiratory pattern employed by insects has been
extensively described, explanations for their occurrence in specific
insects or environments remain controversial. Several hypotheses
have been set forth to try and explain why these different gas
exchange patterns may have evolved (Lighton, 1998; Bradley,
2006; Chown et al., 2006; Slama et al., 2007). Traditionally,
discontinuous gas exchange is thought to be a physiological
adaptation to arid environments which reduces respiratory water
loss. This hypothesis suggests that modulation of the spiracles
prevents high rates of evaporative water loss through the tracheal
system which directly supplies O2 to the tissues. Hetz and Bradley
(2005) proposed that discontinuous gas exchange evolved in
insects to reduce oxidative toxicity since oxygen, even at low
levels, causes oxidative damage to tissues (Yan et al., 1997; Wei
et al., 1998). Since the tracheal system of insects has a high
conductance for gases (Krogh, 1920) an increase in trans-spiracular
resistance is important to maintain pO2 at low levels that prevent
oxidative damage (Hetz and Bradley, 2005). The oxidative damage
hypothesis posits that insects use DGC during periods of low
metabolic activity to reduce, and then regulate, the pO2 at low
levels near the tissues.
Recently one of us has proposed that an interaction between
metabolic rate and the capacity of the respiratory system to deliver
O2 to active tissues may influence the type of gas exchange pattern
observed (Bradley, 2008). The reasoning is as follows. In insects
showing DGC, an increase in metabolic rate shortens the closed
phase since the enhanced rate of O2 consumption causes the
critical level of pO2 to be reached more rapidly. If metabolic rates
increase, the closed phase becomes shorter and shorter until it
completely disappears. Therefore, insects would transition from a
DGC pattern to a cyclical pattern where only the flutter and open
phases are observed. CO2 bursts would still be observed intermittently but levels of CO2 release would never reach zero. If
metabolic rates continued to rise, the flutter phase would shorten
until eventually metabolic rates would be so high that the flutter
phase also completely disappears. The reduction in closed and
flutter phases allows more O2 to enter in support of aerobic
metabolism. This allows CO2 to be released as it is produced,
eliminating bursts of CO2 release.
In this study, we sought to experimentally manipulate
metabolic rates, using temperature, to test whether a clear
transition in respiratory patterns could be observed as metabolic
rates varied. It is well known that an increase in temperature will
increase respiratory rate (Keister and Buck, 1964). However, how
this increase in respiratory rate affects gas exchange pattern has
not been well established. We used two insect species: (1)
Rhodnius prolixus (Triatomid bug) and (2) Gromphadorhina
portentosa (hissing cockroaches). Rhodnius prolixus is a wellstudied species which has been shown to have a pattern of gas
exchange that is categorized by regular periods of rapid CO2 release
523
(CO2 bursts) at rest (Bradley et al., 2003). Metabolic rates in R.
prolixus have been shown to increase after a blood meal, but during
fasting, metabolic rates are relatively low and constant. Therefore,
unfed R. prolixus seem to show constant metabolic rates that could
be directly manipulated via temperature changes. Likewise G.
portentosa seem to have low and constant metabolic rates (due to
their sedentary lifestyle) which could be directly manipulated via
temperature changes. Many species of cockroaches have been
shown to use DGC and to actively ventilate the respiratory system
during the open phase (Slama, 1999; Marais and Chown, 2003;
Dingha et al., 2005; Woodman et al., 2007). During this phase CO2
is released in rapid pulses which are associated with the abdominal
pumping of active ventilation. Using these two species of insects
we aimed to: (1) determine the effect of temperature on metabolic
rate; (2) objectively determine periods of spiracular closure; (3)
observe if changes in metabolic rates are correlated with length of
spiracular closure.
2. Methods
2.1. Insects
2.1.1. Rhodnius prolixus
Insects from Midwestern University were used to found a
colony at the University of California, Irvine. The colony was
maintained at 27 8C and 80% RH on a 12:12 day/night cycle. Each
adult male used for this study was kept in a separate 15 ml vial for
easy identification. Insects were not fed for at least 1 week prior to
the experiment.
2.1.2. Gromphadorhina portentosa
Cockroaches, with approximately equal sizes and masses, were
collected from a colony maintained at the University of California,
Irvine. Two individuals were maintained in one plastic container at
a time, such that a specific roach could be easily identified. All
roaches were maintained in the laboratory at 25 8C on a 12:12
day/night cycle and fed dog food (Holistic Natural Canine Formula,
Bench and Field Pet Foods. Mishawaka, Indiana) and apples, and
given water ad libitum. Food was removed from the containers at
least 1 day prior to date of measurements.
2.2. Effect of temperature on spiracular closure
Individuals were weighed prior to each experiment. Rhodnius
were placed in the experimental chamber and left undisturbed for
55 min (roaches for 135 min) before the measurement period
commenced.
Flow through respirometry was used to measure CO2 release
with Sable Systems (Las Vegas, NV, USA) data acquisition software
controlling an 8-channel multiplexor and logging data from an
infrared CO2 analyzer (Li-Cor model 6262 infrared; Lincoln, NE,
USA). A total of two chambers were attached to the multiplexor: a
baseline (empty) and an experimental (containing the insect)
chamber. During a normoxic experiment, room air was pumped
through two silica and one ascarite/drierite column to be scrubbed
of water and CO2. Rhodnius measurements were conducted in 2 ml
chambers with airflow of 200 ml/min. Gromphadorhina portentosa
were measured in 60 ml chambers with a flow rate of 1000 ml/min.
Flow rate was adjusted using a mass flow controller (Sierra
Instruments, Monterey, CA, USA). As a result, the time constants for
volume exchange were 0.6 s for the Rhodnius measurements and
3.6 s for G. portentosa measurements. Therefore, it took 3 s and 18 s
respectively for 99% of the air to be replaced in the experimental
chambers. Air leaving the experimental chamber passed into the
CO2 analyzer. When an insect was not being measured, its chamber
was still perfused with H2O- and CO2-free air at a rate equal to the
H.L. Contreras, T.J. Bradley / Journal of Insect Physiology 56 (2010) 522–528
524
regulated flow entering the measured chamber. VCO2 was
measured in a random block design at three different temperatures. For each species the three temperatures were bracketed
around the rearing temperature. Rhodnius prolixus, which is
maintained in our laboratory at 25 8C, was therefore measured
at 15 8C, 25 8C or 35 8C while roaches, reared at 20 8C, were
measured at 10 8C, 20 8C or 30 8C. All 10 individuals from each
species were exposed to the three temperature treatments in
random order. Temperature measurements were made on
separate days with the insects being returned to colony
temperature between measurement days.
A total experimental run lasted 55 min for R. prolixus
experiments and 135 min for G. portentosa experiments. During
a run, three 5-min baselines were recorded (where an empty
chamber was read) at the beginning, middle and end of the total
run. Baseline values were used to provide accurate zero values and
to correct for instrumental drift. Insects were observed during the
experiments for signs of activity.
Cor model 6262 infrared; Lincoln, NE, USA) and data were recorded
using Sable Systems (Las Vegas, NV, USA) data acquisition
software.
Before a trial commenced, a single male Rhodnius was placed in
the experimental chamber. This chamber was then placed inside a
plastic bag and submerged into a water bath (Brinkmann RM6
LAUDA Circulator; Delran, NJ, USA) set initially at 15 8C. Rhodnius
were kept submerged at this temperature for at least 30 min before
the trial started. Once the trial had begun, temperature was
increased 2 8C every 30 min. A thermocouple (Multilogger
Thermometer HH506RA, OmegaAnet; Stanford, CT, USA) was
placed adjacent to the insect to determine the exact temperature of
the submerged experimental chamber throughout the experiment.
Baselines values, where an empty chamber was measured, were
taken every 2 h to provide accurate zero values and to correct for
instrumental drift. Insects were observed during the experiment
for activity.
2.5. Data analysis
2.3. Objectively defining spiracular closure
It is very difficult to objectively, and unequivocally, determine
when an insect’s spiracles are fully closed. Due to the difficulty of
defining periods of spiracular closure Chappel and Rogowitz (2000)
combined their data from closed and flutter phases into one value.
We wished to employ an objective, repeatable measure for
quantitatively determining the degree to which the spiracles
were closed during a run.
Previous studies have shown that under hyperoxic conditions
the closed phase is prolonged (Lighton et al., 2004). Therefore, to
determine the closed phase of DGC during a normoxic trial, we first
measured VCO2 during hyperoxic conditions at 15 8C for R. prolixus
and 10 8C for G. portentosa (Table 1). During these trials, instead of
ambient air, 100% O2 (Airgas, Los Angeles, CA, USA) gas was
pumped directly into the chamber being measured at 200 ml/min
for R. prolixus and 1000 ml/min for G. portentosa. The lowest VCO2
values (minimum of 150 s of continuous data) for each individual
during these trials were averaged and multiplied by a factor of four
to raise the cutoff threshold above instrumental noise. All data
points below this ‘‘zero’’ value were considered to be associated
with a period of spiracular closure. These criteria were used to
determine the proportion of time that the spiracles were closed
during a normoxic trial at the three different temperatures being
measured.
2.4. Effect of gradual temperature change on metabolic rate and
spiracular closure
To determine whether the effects of temperature on metabolic
rate and respiratory gas exchange patterns were gradual, as
opposed to occurring at specific temperatures, we measured VCO2
from eight male Rhodnius (different individuals from the ones used
in the above experiment) as temperature was slowly increased
from 15 8C to 39 8C. Room air was pumped through silica/ascarite
to be scrubbed of CO2 and water before it passed through a 2 ml
experimental chamber. Airflow was set at 200 ml/min. Air leaving
the experimental chamber entered an infrared CO2 analyzer (LiTable 1
Minimum rate of CO2 release (VCO2) for R. prolixus and G. portentosa during
hyperoxic conditions (column 2). Column 3 shows the same value multiplied by 4.
This value was used to define periods of closure.
R. prolixus
G. portentosa
VCO2 SE (ml/min)
VCO2 4 (ml/min)
N
0.0464 0.0058
0.362 0.0243
0.1854
1.4463
10
8
Sable Systems (Las Vegas, NV, USA) Expedata analysis software
was used to process VCO2 measurements. CO2 levels were
recorded in parts per million and, after data were zeroed using
baseline values, converted to microliters/min. Data were then
exported into Excel. To determine the effects of temperature
(metabolic rate) on length of spiracular closure, estimates of mass
specific metabolic rates (VCO2, ml/min) were determined for all
individuals at three experimental temperatures (15 8C, 25 8C,
35 8C for R. prolixus and 10 8C, 20 8C, 30 8C for G. portentosa).
Significant differences between metabolic rates for each temperature and proportion of time spiracles were closed were tested
using a repeated measures ANOVA. A repeated measures ANOVA
was also used to determine differences in the proportion of time R.
prolixus maintained their spiracles closed when temperature was
slowly altered. To compare the effects of method used to alter
temperature (fast vs. slow), a univariate ANOVA was used to
compare mass specific metabolic rates at 15 8C, 25 8C and 35 8C in
R. prolixus.
3. Results
3.1. Effect of temperature changes on spiracular closure
The rate of CO2 emission varied with temperature in R. prolixus
and G. portentosa (Table 2). In R. prolixus the temperature
coefficient (Q10) of CO2 release between 15 8C and 25 8C was
2.33 and between 25 8C and 35 8C it was 2.25. In G. portentosa, the
rate of CO2 release was greatly reduced at 10 8C when compared to
20 8C leading to a very high Q10 of 5.34. The Q10 between 20 8C and
30 8C was 2.11.
Table 2
Values for mass and average rate of CO2 release (VCO2) for R. prolixus and G.
portentosa when placed under three distinct temperatures. Values are STP
corrected. N equals sample size.
R. prolixus
Mass SE (g)
VCO2 SE
(ml/min)
Mass specific
VCO2 SE (ml/g/min)
N
15 8C
25 8C
35 8C
0.073 0.007
0.078 0.002
0.073 0.006
0.078 0.003
0.196 0.006
0.401 0.008
1.08 0.067
2.50 0.032
5.64 0.291
10
10
10
G. portentosa
Mass SE (g)
VCO2 SE
(ml/min)
Mass specific
VCO2 SE (ml/g/min)
N
10 8C
20 8C
30 8C
6.08 0.37
6.24 0.40
6.16 0.29
1.19 0.14
6.39 1.40
13.75 1.60
0.19 0.020
1.06 0.165
2.23 0.228
10
10
10
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525
Gas exchange patterns in these two species, also varied greatly
with temperature. At the coldest temperature (15 8C) R. prolixus
showed DGC, where bursts of CO2 release above the calculated
threshold line were interchanged with longer periods of CO2 release
below the threshold (Fig. 1A). When temperature was increased to
25 8C a pattern emerged in which CO2 was released in rapid bursts,
but, during the interburst periods CO2 values oscillated about the
threshold line (Fig. 1B). At 35 8C rapid bursts of CO2 were still
observed. However, using the criterion described in the methods for
determining periods of spiracular closure, about 60% of R. prolixus at
35 8C showed no periods of closure (continuous gas exchange)
(Fig. 1C). G. portentosa also exhibited a transition in gas exchange
pattern as a response to temperature. At the coldest temperature
(10 8C), roaches showed DGC and spent most (87%) of the
experimental period in the closed phase (Fig. 2A). As temperature
increased to 20 8C, DGC was still observed but the closed phase was
much shorter (Fig. 2B). When roaches were placed in 30 8C, the closed
phase completely disappeared in most individuals (Fig. 2C) although
40% still showed very limited periods of spiracular closure.
Using the criterion described in Section 2, we determined the
proportion of time that spiracles were closed in both species at
each temperature (Fig. 3). In R. prolixus, significant (p < 0.05)
differences were observed across all temperature levels in the
proportion of time the spiracles were closed. At 15 8C, the spiracles
were closed for most of the experimental period (90% of the total
observation time) and at 35 8C they were closed for less than 2% of
the time (Fig. 3A). Similarly, in G. portentosa the spiracles were
Fig. 1. Representative gas exchange patterns of R. prolixus at 15 8C (A), 25 8C (B) and
35 8C (C). Data are shown for one individual. Chambers containing an insect were
read for two 20 min periods (black) and an empty chamber was read for three
periods (grey) at 0–5 min, 25–30 min and 50–55 min for baselining. The dashed line
depicts four times the CO2 levels during spiracular closure as calculated from
hyperoxic experiments at 15 8C. Any points below this line are considered periods of
spiracular closure and points above the line are considered periods when spiracles
are not closed.
Fig. 2. Representative gas exchange pattern of G. portentosa at 10 8C (A), 20 8C (B)
and 30 8C (C). Data are shown for one individual. One experimental trial lasted
135 min. Chambers containing a roach were read for two 60 min periods (black) and
an empty chamber was read for three periods (grey) at 0–5 min, 65–70 min and
130–135 min for baselining. The dashed line depicts four times the CO2 levels
during spiracular closure as calculated from hyperoxic experiments at 10 8C. Any
points below this line are considered periods of spiracular closure and points above
the line are considered periods when spiracles are not closed.
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526
Fig. 4. Gas exchange pattern of R. prolixus at temperatures ranging from 15 8C to
39 8C during slow temperature ramping (2 8C per 30 min). Data shown for one
individual. Chambers containing an insect were read continuously for at least 2 h
(black) and an empty chamber was read for four periods (grey) at 0–5 min, 125–
135 min, and 445–450 min for baselining. The dashed line depicts CO2 levels during
spiracular closure calculated from hyperoxic experiments at 15 8C. Any points
below this line are considered periods of spiracular closure and points above the
line are considered periods when spiracles are not closed.
Fig. 3. Proportion of total experimental time that the spiracles were closed in R.
prolixus (A) and G. portentosa (B) at three different temperatures. Data shown as
mean SE for N = 10. Significantly (p < 0.05) different closure times are indicated by
different letters above the column.
closed for the highest proportion of time when roaches were
placed in the coldest temperature (10 8C) and significant (p < 0.05)
decreases in the proportion of spiracular closure occurred with
each increase in temperature (Fig. 3B).
3.2. Effect of gradual temperature change on metabolic rate and
spiracular closure
Having established that respiratory pattern changed with
temperature, we wished to determine if these changes occurred
abruptly at a certain temperature or were gradual over the tested
temperature range. We therefore tested the effects of gradual,
step-wise temperature increases in R. prolixus. As in the previous
experiment, increases in temperature had the effect of increasing
the rate of CO2 emission (Table 3). The temperature coefficient
(Q10) of CO2 release between 15 8C and 25 8C was 1.54 and between
25 8C and 35 8C it was 2.48. Overall, as temperature was increased
slowly (2 8C every 30 min) the rate at which CO2 was released
gradually increased.
Gas exchange patterns also showed gradual change as
temperature was slowly increased. A representative example of
the results obtained is shown in Fig. 4. Fig. 5 shows cumulative data
indicating the proportion of time the spiracles were closed as
metabolic rate increased due to temperature increase. We see that
as mass specific metabolic rate increased the proportion of time
the spiracles were closed decreased. At 33 8C the proportion of time
the spiracles are closed significantly differs from R. prolixus found
at 15 8C (p < 0.05). However, it is not until the water bath
temperature reached 39 8C that the spiracles were closed for a
period of time statistically indistinguishable from zero.
4. Discussion
Our results support the hypothesis that changes in metabolic
rates (related to changes in temperature) influence the type of gas
exchange pattern observed at a given time in insects. Bradley
(2008) has proposed that there are two main factors affecting gas
exchange patterns in insects: metabolic rate and the capacity of the
respiratory system to deliver oxygen to metabolically active tissue.
Many insects show discontinuous gas exchange (DGC) at rest with
closed, flutter and open spiracular phases. However, as metabolic
rates increase, the proportion of time during which the spiracles
Table 3
Values for mass and average rate of CO2 release (VCO2) for R. prolixus when
temperature is slowly increased. Values are STP corrected. N equals sample size.
R. prolixus
Mass SE (g)
VCO2 SE
(ml/min)
Mass specific
VCO2 SE (ml/g/min)
N
15 8C
25 8C
35 8C
0.053 0.007
0.053 0.007
0.053 0.007
0.056 0.008
0.091 0.015
0.219 0.024
1.12 1.68
1.73 0.180
4.29 0.434
9
9
9
Fig. 5. Proportion of time that spiracles were closed vs. mass specific metabolic rate
in R. prolixus as temperature was slowly ramped (2 8C per 30 min) from 15 8C to
39 8C. Data shown for N = 9 (at 39 8C data is for N = 4). Closure declines significantly
(p < 0.05) with temperature. Note that the change is gradual over the temperature
range tested.
H.L. Contreras, T.J. Bradley / Journal of Insect Physiology 56 (2010) 522–528
are closed will become shorter and shorter. Eventually the closed
phase will be eliminated. This is in keeping with the suggestion
that the purpose of the closed phase is a lowering of pO2 inside the
insect (Hetz and Bradley, 2005). By decreasing the time that the
spiracles are closed, the volume of oxygen entry and CO2 release
are adjusted to metabolic demands. This results in a transition in
respiratory pattern as a function of metabolic rate.
The likelihood that an insect will show one respiratory pattern
or another depends on the balance between oxygen content inside
the insect at time of spiracular closure and the rate of oxygen use
through aerobic metabolism. The oxygen content at closure
depends on the architecture (for example volume) of the
respiratory system and the external oxygen concentration. The
rate of metabolism is of course a function of various parameters
including activity, feeding and temperature. In Aphodius fossor,
changes in respiratory gas exchange pattern were observed as a
response to a reduction in ambient oxygen concentrations (Chown
and Holter, 2000). In this study the transitions occurred in
response to increasing temperature.
Determination of the gas exchange pattern being employed by
an insect is not straightforward. The time constants for respiratory
chambers and flow rates can have a profound effect on observed
gas exchange patterns in small insects (Gray and Bradley, 2006). If
these are not appropriate, the patterns described can be artifactual
and not representative of the actual gas exchange pattern
employed by the insect. In this study, the rate of air flow and
chamber volumes produced a time constant of 0.6 s for Rhodnius
and 3.6 s for cockroaches, assuring that 99% of the air in the
chamber turned over every 3 s in Rhodnius experiments and every
18 s in roach experiments. These time constants were appropriate
for our experimental analysis.
Designating the start and end of the closed and flutter phases of
DGC can also be very difficult. Some studies have therefore
combined the closed and flutter phases when describing the
phases of DGC (Chappel and Rogowitz, 2000). In this study we
sought to define each phase of the DGC as objectively as possible.
We therefore characterized the closed phase of DGC using data for
the average release of CO2 during hyperoxic conditions (100% O2)
at the coldest experimental temperatures for each species (R.
prolixus = 15 8C; G. portentosa = 10 8C). Lighton et al. (2004)
reported that an increase in O2 leads to an increase in spiracular
closure. In the present study, using low temperatures and a
hyperoxia, we were able to increase and lengthen the periods of
spiracular closure, allowing us to more accurately determine the
rate of CO2 release observed during a closed phase. Under these
conditions we identified the lowest CO2 release rate from each of
our individual insects. These were never as low as the baseline
values from an empty chamber since CO2 was still given off by the
insect (i.e. small amounts of CO2 could have been released via the
cuticle even when all spiracles were closed). This lowest value was
multiplied by a factor of four to bring the criterion of closure above
instrument noise (Table 1). If CO2 release, during an experimental
trial, fell below this adjusted value then that period was defined as
a period of spiracular closure. If rate of CO2 release was above this
value then that period was defined as being a period when the
spiracles were open.
Using the above guidelines, R. prolixus showed discontinuous
gas exchange when placed in the coldest temperature. During an
interburst period CO2 values fell well below the threshold line and
were close to zero (Figs. 1A and 4). When R. prolixus was placed at
25 8C (Fig. 1B) we observed cyclic bursts with but the interburst
periods were associated with a substantial increase in CO2 release.
Interburst values varied about the threshold line. When placed at
35 8C (Fig. 1C) very few periods of spiracular closure were observed
(Fig. 3A) and the pattern varied between cyclic and continuous.
Similarly, when temperature was slowly ramped, R. prolixus
527
showed DGC at 15 8C and slowly transitioned toward a cyclic gas
exchange pattern (>31 8C) and finally to continuous gas exchange
when temperature was above 37 8C (Fig. 4).
Length of spiracular closure at 25 8C and 35 8C differed between
R. prolixus exposed to thermal ramping and those exposed to an
acute temperature change (Figs. 3A and 5). In fact, it is not until
temperature reaches 39 8C during ramping that the spiracles are
closed to the same level as they were at 35 8C during an acute
exposure. These results are related to differences in the rate of CO2
release between these two methods (Tables 1 and 2). When
temperature was slowly but steadily increased, R. prolixus showed
a significantly (p < 0.05) lower rate of CO2 release at 25 8C and
35 8C than did insects that were held at these temperatures
(Tables 2 and 3). Rapid temperature ramping may cause an
animal’s body temperature to lag behind ambient temperature
(Lutterschmidt and Hutchinson, 1997). However, temperature
ramping in this study was very slow (2 8C every 30 min) and
therefore this phenomenon could not have affected our results.
Studies on Melanoplus nymphs have also shown an overshoot in
oxygen consumption in nymphs placed in any one temperature,
other than the rearing temperature, compared to nymphs exposed
to a slow temperature increase (Roger, 1929). Clearly, respiratory
pattern is adjusted in insects as metabolic rate increases. The time
course of these changes is apparently sensitive to rate of change.
Overall, we found a strong correlation between temperature
and the proportion of time spent in spiracular closure, as defined
by our criteria using the rate of CO2 release (Figs. 3A and 5).
Similarly, Chappel and Rogowitz (2000) saw that at temperatures
above 30 8C, the Eucalyptus boring beetle spent most of the
experimental period in the open phase of the DGC. As temperature
decreased, time spent in the open phase decreased. They also
observed that at 20 8C the interburst phases could not be clearly
separated into a closed or flutter phase and VCO2 never fell to zero,
as was expected in the classic definition of DGC. The absence of a
closed phase has also been seen in grasshoppers (Hadley and
Quinlan, 1993) and termites (Shelton and Appel, 2000, 2001;
Shelton, 2001). We suggest that the absence of a closed phase is
due to an increase in metabolic rate. As metabolic rates increase
further, the flutter phase completely disappears since spiracular
closure is not needed to reduce internal partial pressures of O2.
In our study, G. portentosa also showed a change in gas exchange
pattern as temperature increased (Fig. 2A). Discontinuous gas
exchange has been observed in other cockroach species at low
rates of oxygen consumption (Bartholomew and Lighton, 1985;
Marais et al., 2005; Dingha et al., 2005; Woodman et al., 2007).
When placed at 20 8C a transition to reduced spiracular closure was
observed (Fig. 2B), however the pattern was not one of cyclic bursts
with intermediate lower levels of release as observed in Rhodnius.
Instead, DGC was still employed but the closed phase was much
shorter than that observed at colder temperatures. When roaches
were placed at 30 8C, a continuous gas exchange pattern was
observed with much reduced periods of spiracular closure (Figs. 2C
and 3B). Dingha et al. (2005) observed that the German cockroach
(Blatella germanica) showed a transition from a discontinuous to a
more cyclic gas exchange pattern when temperature was gradually
increased from 10 8C to 35 8C. The giant burrowing cockroach
(Macropanesthia rhinoceros) has been shown to transition from a
discontinuous pattern to a more continuous gas exchange pattern
once oxygen demand increases (Woodman et al., 2007).
In conclusion, we propose that the gas exchange patterns of
insects are not discrete respiratory forms but instead are a
continuum that reflects a balance between oxygen demand and
oxygen supply. As metabolic rates increase, insects must
accommodate the increasing oxygen demand by increasing
spiracular conductance. It is these factors, and not phylogenetic
position, ecological niche, or environmental humidity that
528
H.L. Contreras, T.J. Bradley / Journal of Insect Physiology 56 (2010) 522–528
determines the respiratory pattern employed. For this reason the
respiratory pattern employed by insects is labile and varies with
metabolic demand.
Acknowledgements
We would like to thank Dr. Michael Quinlan for providing
Rhodnius prolixus used to found our insect colony. This study was
supported by Sigma Xi Grants in Aid of Research. One of us is a
fellow of the Alliance for the Graduate and Equity Program.
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