|
| |
|
|
|
|
||||
| Home Help Feedback Subscriptions Archive Search Table of Contents | ||||
First published online April 18, 2006
Journal of Experimental Biology 209, 1662-1677 (2006)
Published by The Company of Biologists 2006
doi: 10.1242/jeb.02203
The significance of spiracle conductance and spatial arrangement for flight muscle function and aerodynamic performance in flying Drosophila
Department of Neurobiology, University of Ulm, Albert-Einstein-Allee 11, 89081 Ulm, Germany
* Author for correspondence (e-mail: fritz.lehmann{at}uni-ulm.de)
Accepted 7 March 2006
 |
Summary |
|---|
 TOP Summary IntroductionMaterials and methodsResultsDiscussionList of symbols and...References |
|---|
Key words: respiration, gas exchange, spiracle opening, aerodynamic force production, IFM, asynchronous flight muscle, flight power requirements, breathing, fruit fly, Drosophila melanogaster
|
Introduction |
|---|
|
TOPSummaryIntroductionMaterials and methodsResultsDiscussionList of symbols and...References |
|---|
;
Casey and Ellington, 1989;
Ellington et al., 1990;
Lehmann and Dickinson, 1997).
However, despite many years of investigation, there is still uncertainty about
how exactly oxygen enters the tracheal system and distributes within the
tracheal network, and carbon dioxide eventually leaves the respiratory system
through the open spiracles (Wigglesworth,
1972). In general, respiratory currents inside the tracheal system
are difficult to predict because they depend not only on several physiological
factors, such as the buffer capacities for respiratory gases and ventilatory
strategies, but also on the morphological architecture of the tracheal system,
including size and location of the spiracle openings
(Weis-Fogh, 1964a;
Weis-Fogh, 1964b). For
example, Drosophila may store oxygen by using haemoglobin in the
tracheal walls to reinforce oxygen partial pressure gradients
(de Sanctis et al., 2005;
Hankeln et al., 2005), and
bicarbonates in the haemolymph may buffer tracheal carbon dioxide at the
expense of changes in pH (e.g. Gulinson
and Harrison, 1996).
In the past, several researchers have attempted to determine the
significance of respiratory currents inside the tracheal system by studying
the role of individual spiracles in respiration and energetics. Bailey was one
of the first physiologists to investigate the interplay between thoracic and
abdominal spiracles by blocking individual thoracic spiracles in the honey bee
(Bailey, 1954). He found that
under CO2 stress, the resting animal produced a net tracheal air
current from the thorax to the abdomen using abdominal pumping. This tracheal
convection was, however, only present when the propodeal spiracle
(=metathoracic spiracle) was blocked, and ceased when air was prevented from
entering through the first thoracic spiracle. Similar to Bernoulli
ventilation, which was later proposed for flight respiration in large beetles
(Miller, 1966), Bailey also
suggested that in flying bees air is inhaled by the first spiracles and
exhaled via the propodeal spiracles. In locusts and cockroaches,
inspired air is routed similarly from the anterior spiracles to the segmental
tracheae and leaves the system through the abdominal spiracles
(Miller, 1982). This type of
breathing was also demonstrated in Lepidopteran pupae
(Schneiderman, 1960), which
may even control the activity of individual spiracles
(Slama, 1999). A
unidirectional air flow was reported for tethered flying hawkmoths Manduca
sexta (Wasserthal, 2001).
In this insect, there is an air stream towards the posterior spiracles that
results from a subatmospheric (negative) pressure at the mesothoracic spiracle
and a positive pressure in the mesoscutellar air sacs. By contrast, in resting
wingless dung beetles the direction of air flow is inverted (retrograde
convection) and tracheal gases flow forward from the posterior to the anterior
body (Duncan and Byrne, 2002).
These authors also reported a lateral asymmetry of air flow inside the animal
and demonstrated that the right mesothoracic spiracle is the primary route for
respiratory gas exchange. A study on running energetics in the ant
Camponotus hypothesised that asymmetries in gas exchange rate might
also occur during the discontinuous gas exchange cycle (DGC)
(Lipp et al., 2005). This
hypothesis arose from the finding of various numbers of `O'-peaks within a
single gas exchange cycle, suggesting that multiple spiracles may be involved
in tracheal gas exchange. Moreover, in the arid-adapted ant Cataglyphis
bicolor, the thoracic spiracles act as high-capacity gateways to the
tracheal system and are responsible for approximately 90% of the total gas
exchange rate during running activity
(Lighton et al., 1993a). In
this ant, the abdominal spiracles combined only have approximately half the
diffusive capacity of a single thoracic spiracle
(Lighton et al., 1993a).
Size constraints in small insects such as the fruit fly
Drosophila, make it difficult to assess tracheal air currents,
spiracle function or tracheal partial pressures. The situation is even more
complex in flying animals in which the spiracles may dynamically vary their
opening areas according to changes in flight power requirements. For example,
the results of a study on spiracle opening behaviour in flying
Drosophila are consistent with at least three basic mechanisms of
spiracle control (Lehmann,
2001). The fly may achieve an average tracheal conductance by
either (i) matching the area of each thoracic spiracle to the respiratory
needs, (ii) dynamically closing and opening the spiracles over time, or (iii)
closing some spiracles while other spiracles remain open. All mechanisms could
give similar mean tracheal conductance when estimated within a certain time
period, although the latter mechanism, in particular, might produce temporal
fluctuations in the local supply of oxygen. The importance of matching at
least the bilateral control of spiracle activity arises from the fact that the
left side of the tracheal system is virtually isolated from the right side. In
Drosophila, there are only a few tracheal commissures (pro- and
metathoracic dorsal commissure, and pro-, meso- and metathoracic ventral
commissures) that allow exchange of respiratory gases between the body sides
(Miller, 1950). In the locust
Schistocerca gregaria the thoracic system is, moreover, isolated from
the abdomen (Weis-Fogh,
1964b), while in Drosophila large tracheae connect the
thoracic parenteric air sac with the abdominal air sac
(Miller, 1950). For geometric
reasons, changes in the spatial distribution of the total spiracle opening
areas might directly hinder proper flight muscle function because the insect
flight muscle has no anaerobic capacity and thus relies on instantaneous
oxygen supply (Ziegler, 1985).
By contrast, the extensive tracheal development in Drosophila, with
broad latero-linear air sacs such as the parentic-, pleural- and lateroscutal
sac, favours the establishment of homogenous partial pressures on the
ipsilateral side even when gas exchange rates are not balanced between the
ipsilateral meso- and metathoracic spiracle.
Consequently, in this study we examine how reducing the size and deleteriously interfering with the spatial distribution of gas exchange areas impair flight muscle function and aerodynamic force production in the small fruit fly Drosophila melanogaster. This was done by selectively blocking thoracic spiracles in tethered animals flying inside a flight simulator. Flow-through respirometry and simultaneous measurements of flight force production and wing kinematics allow the determination of changes in vital flight parameters under a vast variety of breathing conditions. Incorporating energetic and aerodynamic theory, we show how (i) maximum mechanical power output of the indirect flight muscle (IFM), (ii) the efficiency with which the muscle converts chemical energy into muscle mechanical power, (iii) lift and drag coefficients for flapping wing motion and (iv) aerodynamic efficiency, all change with changing arrangements of spiracle conductance.
|
Materials and methods |
|---|
|
TOPSummaryIntroductionMaterials and methodsResultsDiscussionList of symbols and...References |
|---|
We determined the functional significance of changes in respiratory gas exchange through the four major thoracic spiracles of Drosophila by covering individual spiracle openings with small droplets of commercial two-component epoxy glue (5-min epoxy, R&G, Waldenbuch, Germany). The resin component of the glue consisted of bisphenol A- and F-epichlorhydrine, and the active component of the hardener was 2,4,6-tri(dimethylaminomethyl)phenol. The glue was selected according to its property to cure fast and in very small volumes. We tested the glue's toxicity for Drosophila by exposing 25 flies in a standard Drosophila vial to the resin, the hardener and the polymerised glue. Although the flies did physically contact the polymerised glue and its components, none of the animals died or superficially changed behaviour within the testing period of 24 h. Moreover, it seems unlikely that the small cuticle area covered by the epoxy droplets influenced the mechanical properties of the thoracic box during wing flapping, for two reasons: first, the glue droplet was only marginally larger than the opening area of the meso- or metathoracic spiracles, and second, an experiment in which we placed a glue droplet close to but not on the animal's spiracle opening did not produce any alterations in aerodynamic force production. Throughout the manuscript we use the term `unmanipulated flies' to mean tethered flying animals with none of their thoracic spiracles sealed.
The flies were allowed to recover from the tethering and sealing procedure
for at least 30 min before being placed into the flight arena. To derive
muscle mass-specific power output, we measured the mass of each fly after each
experiment using a balance (sensitivity 0.01 mg; Sartorius MC210P,
Göttingen, Germany) and assuming a flight muscle-to-body mass ratio of
approximately 0.3 (Lehmann and Dickinson,
1997). For body mass estimation, we did not remove the glue from
the spiracle openings because its contribution to the fly's total mass was
negligible. We calculated mean lift and drag coefficients from total flight
force, wing velocity and wing size by employing quasi-steady aerodynamic
theory and assuming that the chordwise aerodynamic circulation is maximum
close to a spanwise location of 65% wing length
(Birch and Dickinson, 2001;
Lehmann et al., 2005;
Ramamurti and Sandberg, 2001).
A detailed description of this procedure is given in a previous study on
Drosophila flight (Lehmann and
Dickinson, 1998). Body mass (means ± s.d.) of the tested
animals was 0.92±0.08 mg (N=10 flies, all thoracic spiracles
left open), 0.88±0.18 mg (N=26, 1 spiracle sealed),
0.94±0.20 mg (N=43, 2 spiracles sealed), 0.96±0.17 mg
(N=23 flies, 3 spiracles sealed) and 0.92±0.08 mg
(N=5 flies, 4 spiracles sealed). A statistical test on body mass
showed no significant differences between the tested groups (ANOVA,
P>0.05). Mean temperature during the experiments was
23.5°C.
|
2745 µm2) is approximately
20% larger than the mesothoracic opening (2186 µm2),
resulting in a total maximum area for thoracic respiratory gas exchange of
approximately 9862 µm2. In comparison, a previous study
described the thoracic spiracles in Drosophila melanogaster as oval
openings of approximately 60 µmx25 µm at the surface
(Manning and Krasnow, 1993).
From these values we calculated a total thoracic exchange area of 5675
µm2, which is approximately 42% smaller than the value measured
in the present study. There could be several reasons for this discrepancy.
First, although the aspect ratio between 60 and 25 µm fairly matches the
elongated shape of the mesothoracic spiracle, the metathoracic spiracle seems
to be more circular with a diameter between 50 and 60 µm, and thus appears
to be larger than measured by Manning and Krasnow. Second, since we measured
total opening area from the size of the thick sclerite that borders the
spiracle opening at the surface, we might have overestimated the inner opening
size (Nikam and Khole, 1989).
Third and alternatively, the females that were used in the present study were
chosen according to body size and represented the largest animals from the
population. Larger females typically show a better flight endurance than their
smaller relatives. Consequently, since larger fruit flies exhibit larger
spiracle openings (F.O.L., unpublished data), it might be that the difference
between the two opening measurements is a real difference in size rather than
due to substantial differences in measurement techniques.
Compared to thoracic spiracles, the area of the 14 abdominal spiracles was
not measured directly, but was estimated by assuming that it represents 5% of
the fly's total gas exchange area or approximately 493 µm2
(Manning and Krasnow, 1993). A
simple behavioural test showed that gas exchange mediated by abdominal
spiracles is high enough to satisfy the oxygen needs during resting
metabolism, whereas in experiments in which gas exchange through all 18
spiracles was completely blocked, the flies died within approximately 10 min
after the treatment (N=5 animals). This result is similar to a study
on the regulation of carbon dioxide release from the thoracic and abdominal
spiracles in the ant Cataglyphis
(Lighton et al., 1993a). In
this animal, the abdominal spiracles have half the diffusive capacity of a
single thoracic spiracle and can fully meet the ant's oxygen uptake and carbon
dioxide release at resting metabolism.
|
Fig. 2B shows a typical trace of CO2 measurement, with periodic switching between ambient air and the supply of CO2. After this pre-check, the mesothoracic spiracle was sealed and the testing procedure performed again. Despite the high partial pressure of CO2 on the outside of the thorax, the small seal typically blocked the entire inflow of CO2 into the respirometric chamber (Fig. 2C). Afterwards, in a control experiment, we carefully removed the seal that fully restored the diffusivity of the mesothoracic spiracle for CO2 (Fig. 2D). Note that the vibrating thorax of a flying fly might cause leakage of the spiracle seal, and this effect is not simulated in the simple testing procedure. Therefore, we covered the mesothoracic spiracle of tethered flies and flew the animals for at least 15 min. We subsequently dissected those animals in a procedure similar to that described above and checked for any gas leakages. In none of the five tested flies did we measure any CO2 flux through the sealed spiracles, suggesting that thorax vibrations during flight do not harm the quality of the spiracle seals. In sum, the outcome of the pre-tests convinced us that the epoxy seal used in this study is able to sufficiently block respiratory gas exchange through the spiracle openings in a flying fruit fly.
|
;
Fig. 3A). A more detailed
description of the experimental apparatus and the procedure is given elsewhere
(Lehmann and Dickinson, 1997).
Under closed-loop feedback conditions, fruit flies actively modulated the
azimuth velocity of a vertical dark stripe displayed in the panorama using the
relative difference in stroke amplitude between the two beating wings. The
feedback conditions were set according to previous experiments on tethered
flying fruit flies (Lehmann and Dickinson,
2001). While flying in closed-loop, Drosophila changed
both kinematic and respirometric parameters in response to the vertical motion
of an open-loop striped grating. We have previously shown that under those
conditions fruit flies maximize their locomotor output, while trying to follow
the upward motion of the horizontal stripes
(Lehmann and Dickinson, 1998).
This procedure is equivalent to adding weight to the animal and elicits
maximum flight forces similar to those obtained in freely flying flies using a
weight-lifting technique (Lehmann,
1999).
Concurrently, we employed flow-through respirometry with a flow rate of
1000 ml min–1 and used a Li-cor 7000 gas analyser (Licor,
Lincoln, Nebraska, USA) to measure the rate of carbon dioxide release during
flight. The internal filter frequency of the gas analyser was set to 0.2 s.
Data sampling frequency was 125 Hz and wash-out time constant 
of the 15
ml respirometric chamber was approximately 910 ms (wash-out
time
e–x/;
Lehmann and Heymann, 2005).
Respirometric data of each animal were corrected for both temporal shift due
to wash-out and the gas delay due to the connecting tubings, and were
eventually normalized to standard temperature and pressure (STP). Further
analysis and calibration of kinematic and respiratory data were performed as
described elsewhere (Barton et al.,
2005). Metabolic data and power requirements for flight are given
as flight-specific values by subtraction of resting rates, and if not stated
otherwise also expressed as indirect flight muscle (IFM) mass-specific
units.
Experimental procedure
To vary the size of total diffusive spiracle area in the flying fruit fly,
we tested different combinations of spiracle sealing in random sequences and
subsequently pooled the data subsets (Figs
3,
4,
5). This means that in case of
a single seal, we tested flight performance under four different experimental
conditions: a blockage of the left sp.ms (mesothoracic spiracle,
N=6 flies), right sp.ms (N=7 flies), left
sp.mt (metathoracic spiracle, N=7 flies) and right
sp.mt (N=6 flies). In flies with two sealed spiracles we
investigated flight and muscle performance under six different experimental
conditions (sealed left and right sp.ms, N=8; left and right
sp.mt, N=7; left sp.ms and sp.mt, N=8; right
sp.ms and sp.mt, N=6; left sp.ms and right
sp.mt, N=6; and the right sp.ms and left sp.mt, N=8
flies). Results obtained from flies with three sealed spiracles are mean
values of four experimental conditions (sealed left sp.ms, sp.mt and
right sp.ms, N=7; left sp.ms, sp.mt and right sp.mt,
N=8; right sp.ms, sp.mt and left sp.ms, N=4; right
sp.ms, sp.mt and left sp.mt, N=4 flies). Moreover, previous
results on Drosophila flight energetics have shown that there is a
transient effect on flight performance after the initial take-off during which
flight force production and CO2 release rate peak for a short time.
This transient peak was present in most of our experiments except in flies
with all four thoracic spiracles sealed. Consequently, to circumvent any
transient phenomena, we excluded the first 5 s and the last 2 s of each flight
sequence from our analysis. Throughout the manuscript all values are given as
means ± s.d.
|
|
|
Results |
|---|
|
TOPSummaryIntroductionMaterials and methodsResultsDiscussionList of symbols and...References |
|---|
|
The different regression slopes of kinematic measures (amplitude:
1.90x103 deg. µm–2, frequency:
4.70x 103 Hz µm–2 diffusive area) and
relative flight force production (0.11x103 relative force
µm–2 diffusive area;
Table 1) imply that changes in
wing velocity cannot exclusively explain the changes in aerodynamic force
production, because wing velocity is directly proportional to the product
between stroke amplitude and frequency
(Lehmann and Dickinson, 1998).
Consequently, changes in total spiracular conductance also altered the lift
coefficient for flapping wing motion that decreased by approximately 69%, from
1.21 at a diffusive area of 10355 µm2 to 0.38 at 493
µm2 (Fig. 3E).
Moreover, since flight force is the vector sum of lift and wing profile drag,
the wing's drag coefficient decreased likewise and similarly to the lift
coefficient by approximately 59%, from 0.65 at 10 355 µm2
spiracle opening area to 0.38 during breathing through abdominal spiracles
only (Fig. 3F).
Power requirements and metabolic power
As a consequence of the reductions in (i) wing kinematics, (ii) aerodynamic
lift production and (iii) drag coefficient, the maximum power requirements for
flight, such as the flight muscle mass-specific induced- and profile
requirements, decreased with the decreasing number of open thoracic spiracles
(Fig. 4A,B). Superficially,
mass-specific induced power seemed to be more affected by the respiratory
restrictions than profile power and decreased by 91% from 33.8 W
kg–1 in unmanipulated animals to 3.2 W
kg–1, when gas exchange only occurred through the abdominal
spiracles (Fig. 4A,
Table 1). We found
significantly smaller changes for mass-specific profile power, which decreased
by 69% with decreasing gas exchange area from 64.7 in unmanipulated
flying Drosophila to 20.2 W kg–1 when all thoracic
spiracles were sealed (91±22% vs 69±15%,
t-test, P<0.05, N=8;
Fig. 4B,
Table 1). According to
Ellington's energetic theory for flapping wing motion, mechanical power output
of the asynchronous flight muscle is the sum of induced and profile power
requirements, assuming 100% energy elastic storage within the flight motor
(Ellington, 1984b). Similar to
induced and profile power requirements, this measure decreased with decreasing
spiracle opening area by 7.5±1.0 W kg–1
µm–2 spiracle area (linear regression fit,
y=13.1+7.5x; Fig.
4C, Table 1).
Flight-specific metabolic power was calculated from the instantaneous
measurements of CO2 release during flight and was 68% lower in
flies that only breathed through abdominal spiracles (334 W
kg–1, Fig. 4D)
compared to the unmanipulated control group (1031 W
kg–1; Fig. 4D,
Table 1).
Changes in muscle and flight efficiency
Due to the changes in flight muscle mass-specific mechanical power output
and metabolic power, muscle efficiency changed likewise.
Fig. 5A shows that muscle
efficiency, defined as the ratio between metabolic and muscle mechanical power
output, decreased from 9.8±1.2% in unmanipulated flies to
5.6±1.5% in flies that only breathed through a single thoracic
spiracle. Interestingly, muscle efficiency apparently recovered
(7.2±1.8%) when Drosophila only breathed through abdominal
spiracles compared to gas exchange through a combined diffusive area of
abdominal and 1 (5.6±1.5%) and 2 (6.3±1.6%) thoracic spiracles
(Fig. 5A). However, this result
might be partly ascribed to the delayed release of CO2 (see
paragraph below) after flight initiation that produced a temporal mismatch
between steady-state power requirements for flight and respiratory activity of
the animal. In this sense, the apparent recovery of muscle efficiency during
breathing through the abdominal spiracles does not reflect changes in the
physiological state of the flight musculature but results from the measuring
method.
In contrast, aerodynamic efficiency of wing motion, defined as the ratio
between minimum power requirements for flight (i.e. Rankine–Froude
power) and muscle mechanical power, decreased linearly with decreasing gas
exchange area from a maximum of 26.8±0.92% to
11.2±5.3% during abdominal breathing
(Fig. 5B,
Table 1)
(Ellington, 1984b). Total
flight efficiency is the product of muscle and aerodynamic efficiency and a
measure for the overall performance of the chemo-aerodynamic conversion
process of Drosophila's flight apparatus. During flight of the tiny
fruit fly, total flight efficiency decreased from 2.61±0.35% in
unmanipulated animals to a value well below 1% (0.77±0.30%) during
pure abdominal breathing (Fig.
5C).
Spatial distribution of spiracle opening areas and flight performance
Most of the changes in wing kinematics, flight power requirements,
metabolic rates and flight efficiencies shown in Figs
3,
4,
5 (grey data) can be explained
by the changes in flight force production. It has previously been demonstrated
that wing kinematics including muscle-mechanical and aerodynamic efficiency
co-vary with alterations in lift production – and most of the measured
alterations may be attributed to this effect
(Lehmann, 2002;
Lehmann and Dickinson, 1997).
Thus, we separated the changes due to variations in flight force production
from those caused by variations in the arrangement of spiracle conductance.
This was achieved by comparing the various measures (grey bars) in Figs
3,
4,
5 with measures from
unmanipulated animals at flight forces that matched (within ±2%
accuracy) the maximum force of those flies whose spiracles had been
manipulated. According to Fig.
3D, these body weight-specific forces are: 1.01 (1 thoracic
spiracle blocked), 0.74 (2 thoracic spiracles blocked), 0.48 (3 thoracic
spiracles blocked) and 0.29 relative flight force (four thoracic spiracles
blocked). In other words, while the grey bars in Figs
3,
4,
5 represent measures at 1%
maximum flight force production of manipulated flies (0–3 open
spiracles) and the unmanipulated control group (4 spiracles open), the red
data (means ± s.d., Figs
3,
4,
5) were measured in
unmanipulated animals at times when the flies produced flight forces equal to
one of the five force values shown in Fig.
3D. As mentioned before in the Materials and methods, the fruit
flies varied flight force production between maximum (1.36 force/weight) and
minimum (0.29 force/weight) values in response to the vertically oscillating
horizontal background stripe grating displayed inside the virtual-reality
background flight arena. To further highlight the effect of changes in local
oxygen supply to the flight muscles, we subtracted these data (red) from the
results obtained during spiracle manipulation (grey data, Figs
3,
4,
5) and then plotted the most
essential differences as a function of spiracle opening area in
Fig. 6.
|
64% higher muscle mechanical power
output than breathing through all 18 spiracles in unmanipulated flies (at 0.29
relative flight force). The relative changes in muscle mechanical power output
at 493, 3780 and 5424 µm2 spiracle opening areas were
significantly different from those at 7889 and 10 355 µm2
(t-test, P<0.05, Fig.
6A). (2) Compared to unmanipulated animals, relative muscle
efficiency did not change significantly when oxygen supply was restricted to 3
(7889 µm2) and 2 (5424 µm2 opening area) thoracic
spiracles (t-test, P>0.05,
Fig. 6B). However, muscle
efficiency appeared to be significantly higher during pure abdominal breathing
(5% of total spiracle opening area) when compared with unmanipulated animals
(t-test, P<0.001, Fig.
6B). The tendency towards higher relative muscle efficiencies at
smaller respiratory exchange areas suggests that even the worst condition for
tracheal oxygen supply (abdominal breathing) does not impair the IFM
chemo-mechanical conversion efficiency (linear regression fit,
y=63.2–7.6x10–3x,
R2=0.41, N=5, P=0.24). (3) Relative lift
coefficient significantly increased with increasing diffusive area (linear
regression fit, y=–68.6+6.84x10–3x,
R2=0.97, N=5, P=0.002,
Fig. 6C). During pure abdominal
breathing, the lift coefficient was 54% of the lift coefficient
determined in unmanipulated animals producing similar aerodynamic force. (4)
With increasing spiracle opening area, aerodynamic efficiency increases
significantly with a slope of 5.37x10–3
µm–2 (linear regression fit,
y=–55.7+5.37x10–3x,
R2=0.97, N=5, P=0.0026,
Fig. 6D), suggesting that
oxygen supply through all thoracic and abdominal spiracles produces the best
aerial performance score in Drosophila
(Fig. 6D). Aerodynamic
efficiency during pure abdominal breathing was 48% below the maximum
value obtained in unmanipulated flies. In conclusion, the latter results
suggest that manipulations of the arrangement of spiracle conductance are more
likely to cause subtle alterations in wing motion that alter aerodynamic
efficiency, than significantly changing the chemo-mechanical conversion
efficiency of the asynchronous flight musculature.
Difference between meso- and metathoracic oxygen supply
To evaluate the functional differences between meso- and metathoracic
spiracle-mediated respiration, we compared kinematic, aerodynamic and
energetic variables in experiments in which we sealed either the two anterior
meso- or the two caudal metathoracic spiracles.
Fig. 7 shows the relative
performance differences (`meso- – meta-') normalized to the absolute
performance measured in unmanipulated flies. The metathoracic spiracle opening
area is 26% larger than the mesothoracic spiracle opening area
(Fig. 1), so we expected an
26% higher contribution of the metathoracic spiracle to the overall
flight performance score. Although the data in
Fig. 7 confirmed this
hypothesis, yielding a mean relative difference of –23.6±12.6%
(N=12 measures), the strength in reduction slightly differed between
the various flight measures. Except for stroke amplitude, stroke frequency and
metabolic power, all values were significantly different from zero
(t-test, P<0.01, Fig.
7).
|
).
In our experiments total flight force diminished after this initial peak to a
steady state value that was typically equal to hovering flight force and thus
equal to the animal's body weight (Fig.
8A, black trace). The force peak, including the corresponding peak
in CO2 release rate, ceased completely when respiratory flux was
limited to 50% total thoracic spiracle opening area
(Fig. 8, red trace). During
pure abdominal breathing, we found a transient mismatch between force
production and CO2 release rate, suggesting that CO2 is
partly buffered in the tracheae and haemolymph at this early stage of the
flight sequence (Fig. 8, blue
trace). At flight stop, the buffered CO2 was then apparently
released during the subsequent resting period (Figs
8,
9). Moreover, at the instant
after flight stop, we characteristically found a transient steep decrease in
CO2 release (Fig.
9D, arrow) followed by a more moderate decrease during which flies
released the remaining tracheal CO2 over a time period of up to 40
s (Fig. 10A).
|
|
|
Carbon dioxide buffering and delayed release produced by nine flight starts
and stops of a single fly are shown in Fig.
9A,B. Averaged data for six flies are plotted in
Fig. 9C,D. The temporal
dynamics of CO2 release rate after flight stop suggests a first
order exponential decay with a mean time constant of 14.6 s
(y=–2.3+e–x/14.6,
R2=0.92, 
2/d.f.=1.49;
Fig. 10A, red line). This is
consistent with the assumption of partial pressure gradient-driven diffusive
respiratory processes in the fruit fly. For comparison, in unmanipulated flies
CO2 release rate after a flight sequence returned back to 67% of
the resting rate within 1.89±1.08 s (time constant of first order
exponential fit, y=–0.11+e–x/1.89,
R2=0.79±0.15,
2/d.f.=4.86±3.0, N=25 fits, 9 unmanipulated
animals). Approximately half of this time, however, is due to the wash-out
time constant of the respirometric chamber (0.9 s; see Materials and
methods).
The temporal integral of post-flight CO2 release rate allowed us to estimate the combined in vivo haemolymph–tracheal buffer capacity for CO2 in Drosophila. We plotted these values in Fig. 10B as a function of mean flight force that was measured within the last 2 s prior to rest (pre-resting force). According to our analysis, mean CO2 buffer capacity of Drosophila amounted to approximately 33.5±13.9 µl g–1 body mass (N=35 sequences, blue line, Fig. 10B). Interestingly, this buffer capacity did not significantly change with increasing pre-resting flight force production or power requirements for flight as shown by linear regression analysis (linear regression fit on pre-resting force, P=0.46, N=35 flight sequences, Fig. 10B).
|
Discussion |
|---|
|
TOPSummaryIntroductionMaterials and methodsResultsDiscussionList of symbols and...References |
|---|
33.5 µl g–1
body mass.
Spiracle conductance and tracheal partial pressure for CO2
Although tethered flying Drosophila sporadically employ
ventilation to manipulate tracheal gas flow, diffusion is still to be
considered the main type of respiration in fruit flies, and diffusive theory
may be applied (Kestler, 1985;
Lehmann, 2001;
Lehmann and Heymann, 2005;
Weis-Fogh, 1964a). Together
with our experimental data, the analytical framework conveniently permits
estimations of spiracle conductance and partial pressures for CO2
inside the tracheal system, and thus estimations of the animal's physiological
stress resistance to high tracheal CO2 concentration during flight.
Tracheal partial pressure for a gas, PT, that diffuses
through a spiracle opening can be expressed as:
 | (1) |
is the rate of gas flux, G
is the conductance, and PA is the partial pressure of the
gas in the ambient air (Kestler,
1985). The conductance for gas flow in turn depends on tracheal
geometry, the diffusion coefficient D and the capacitance coefficient
ß for that gas. This relationship can be expressed as:
 | (2) |
). The diffusion
coefficient for CO2 is 0.165 cm2 s–1,
and the capacitance coefficient is 410.5 nmol cm–3
kPa–1 (Kestler,
1985). From the values and equations above, we calculated the
maximum spiracle conductance for our various experimental conditions as: 27.9
(unmanipulated animal), 21.2 (1 thoracic spiracle sealed), 14.6 (2 thoracic
spiracles sealed), 10.2 (3 thoracic spiracles sealed) and 1.32 ng gas
kPa–1 s–1 (four thoracic spiracles
sealed).
Previous analyses of tracheal partial pressure in diffusion-based
respiratory systems relied on the assumption that maximum spiracle opening
area in an insect matches the instantaneous gas exchange rate at maximum
locomotor performance (Lehmann,
2001). Originally, this assumption was reinforced by the finding
that most insects have developed strategies to avoid respiratory water loss
through the open spiracles, i.e. the discontinuous gas exchange cycle, DGC
(Harrison and Roberts, 2000;
Lighton, 1994;
Lighton, 1996;
Miller, 1981;
Slama, 1994;
Snyder et al., 1995). Gas
exchange areas larger than needed would reinforce tracheal water loss at
maximum locomotor performance and thus increase the risk of desiccation in
xeric environments (Lighton,
1994). The data in Fig.
3D provide support for this hypothesis and our experiments thus
permit derivation of maximum tracheal partial pressure for CO2
(PT,CO2) under the various breathing conditions. Employing
the equations above and setting spiracle conductance according to the various
maximum spiracle opening areas, we estimated the following mean values for
PT,CO2 inside the tracheal system during flight:
0.98±0.23 (unmanipulated animal), 0.83±0.17 (1 thoracic spiracle
sealed), 1.20±0.25 (2 thoracic spiracles sealed), 1.47±0.24 (3
thoracic spiracles sealed) and 5.52±0.36 kPa (4 thoracic spiracles
sealed). A slightly higher value for tracheal partial pressure of
CO2 (1.4±0.2 kPa) at maximum flight performance in
unmanipulated Drosophila was calculated in a study on spiracle
control strategies (Lehmann,
2001).
The above results demonstrate that according to total spiracle opening
area, mean PT,CO2 may increase approximately 1.5-fold from
0.98 kPa in unmanipulated flies (10 355 µm2 opening area)
to 1.47 kPa in flies in which total gas exchange area was limited to 3780
µm2. A gas exchange area of 493 µm2 even produces
a PT,CO2 that is 6.6 times higher than in unmanipulated
flies. Although statistical analysis reveals that all estimations of
PT,CO2 are significantly different from each other
(t-test, P<0.05), a linear regression fit suggests that
the slope between PT,CO2 and gas exchange area is not
significantly different from zero (linear regression fit,
y=4.38–4.26x10–4x,
R2=0.66, P=0.09, N=5). Nevertheless, the
small trend in the data set suggests that Drosophila is able to
partially cope with the reduction in spiracle opening area by increasing the
partial pressure gradient for CO2 between tracheal system and
ambient air of up to 5–6 kPa.
In comparison, since ambient partial pressure for oxygen is constant,
oxygen uptake rate in Drosophila can only be reinforced by
ventilation or by actively lowering the tracheal partial pressure for oxygen
(Wigglesworth, 1972). For
example, it has recently been shown that during hovering flight force
production, tethered Drosophila sporadically employ the proboscis as
a pump to actively ventilate their tracheal system
(Lehmann and Heymann, 2005). A
possible mechanism for active buffering of tracheal oxygen is haemoglobin
storage. This phenomenon is discussed in the paragraph on `the significance of
CO2 buffer capacity'.
There are comparatively few data on tracheal partial pressure estimates of
respiratory gases in flying insects, and most of the available data refer to
the DGC. For example, Harrison et al.
(Harrison et al., 1995)
reported for grasshoppers that during the DGC interburst period, haemolypmph
PCO2 rises from 1.8 to 2.26 kPa with minimal acidification
of the haemolymph. The authors concluded that spiracle opening is induced at
internal threshold levels between 2 and 2.9 kPa
(Gulinson and Harrison, 1996).
Moreover, it has previously been shown that in many insects an endotracheal
partial pressure of 4–6 kPa triggers the peripherally mediated
inactivation of the spiracle closer muscle (for reviews, see
Lighton, 1996;
Krogh, 1913). The highest
values reported for PT,CO2 during the DGC was for
Cecropia pupae of 7 kPa that did not fall below a minimum
threshold of about 3.6 kPa (Burkett and
Schneiderman, 1974). The overall shift of
PT,CO2 in Lepidopteran pupae towards higher tracheal
partial pressures has been interpreted as a consequence of the high
vulnerability of the pupae to desiccation
(Harrison et al., 1995).
If we consider that in an insect a high PT,CO2, due to
prolonged DGC interburst intervals, indicates a strategy to avoid respiratory
water loss, the comparatively low PT,CO2 measures in
unmanipulated flying Drosophila would in turn suggest that in this
insect respiratory water loss is of minor importance for total water balance.
This interpretation is driven by the finding that in several insects,
respiratory water loss is relatively small compared to cuticular transpiration
so that the DGC does not predominantly influence the water balance of the
animal. For example, the ratio between cuticular and respiratory transpiration
is 98.1:1.9 in Camponotus vicina and 92.0:8.0 in the ant
Cataglyphis bicolor (Lighton,
1988), 87.0:13.0 in the ant Pogonomyrmex rugosus
(Lighton et al., 1993b) and
the cockroach Periplaneta americana
(Machin et al., 1991),
97.0:3.0 in the grasshopper Romalea guttata
(Hadley and Quinlan, 1993) and
95.4:4.6 in the grasshopper Taeniopoda eques
(Quinlan and Hadley, 1993).
However, in flying Drosophila melanogaster this ratio appears to be
almost inverted and amounts to 17.4:82.6
(Lehmann, 2001). From these
values we conclude that the spiracles represent a significant route for
tracheal water loss in the fruit fly, and with respect to the hypothesis
above, this cannot easily account for the low PT,CO2 in
the unmanipulated flying animal.
The significance of CO2 buffer capacity
The dead volume of the tracheal system serves as a buffer space for
respiratory gases. In addition, Drosophila may store oxygen using
haemoglobin that is mainly synthesized in the tracheal walls and the fat body
of the animal (de Sanctis et al.,
2005; Hankeln et al.,
2005). During transient locomotor activity, the indirect flight
musculature might benefit from haemoglobin-mediated oxygen transport and
storage that produces a temporal mismatch between the uptake rate of oxygen
through the spiracles and flight force production. To largely circumvent this
problem in our respiratory measurements, we excluded the first 5 s in each
flight sequence from our analysis, expecting to achieve rather steady-state
flight metabolism of 10–15 times the resting rate
(Casey, 1989;
Casey and Ellington, 1989;
Lehmann and Dickinson, 1997).
By contrast, tracheal CO2 concentration is correlated with the
haemolymph bicarbonate level that buffers CO2 at the expense of
changes in pH (Harrison et al.,
1995). Gulinson and Harrison investigated CO2 buffering
in the grasshoppers Romalea guttata and Schistocerca
americana by injections of NaHCO3, HCl and NaOH into the
haemolymph (Gulinson and Harrison,
1996). In our experiments, by contrast, we derived CO2
buffer capacity from the delay in gas release after flight stop
(Fig. 10). Since the amount of
total flight-specific CO2 released after flight was independent of
pre-resting flight force production and thus of metabolic activity, we suggest
that the value of 33.5 µl CO2 g–1 body mass may
represent a maximum estimation of Drosophila's total CO2
buffer capacity (tracheal and haemolymph buffer).
To assess the flight time during which an unmanipulated fruit fly may rely
on CO2 buffering instead of CO2 release through the
spiracles, we converted the value of 33.5 µl CO2
g–1 body mass into units of time that yielded
2.30±0.95 µl s–1 g–1 body mass.
Subsequently, we compared this measure with the mean CO2 release
rate produced during hovering flight conditions, that is 2.8 µl
s–1 g–1 body mass
(Lehmann et al., 2000). The
ratio between both values (2.3/2.8) suggests that CO2 buffer
capacity ensures flight for only 0.82 s and thus for a relatively short
flight time. Moreover, this value might explain why the large thoracic
spiracles open immediately after flight initiation in this insect
(Fig. 8)
(Lehmann, 2001). In a resting
fruit fly that exclusively breathes through the abdominal spiracles, a
tracheal CO2 partial pressure threshold of 5.52 kPa (see previous
paragraphs) would be reached within 3.1 s at the given CO2 buffer
capacity (resting metabolism=0.74±0.33 µl s–1
g–1 body mass) (Lehmann
et al., 2000). Ignoring all potential errors associated with these
findings, the latter result might explain why Drosophila melanogaster
rarely employs a clear DGC pattern during rest compared to many other insects
(Harrison et al., 1995;
Lighton, 1994;
Lighton, 1996;
Williams and Bradley, 1998).
During discontinuous breathing the spiracles open only sporadically with
typical time periods in the range of several minutes. Within a DGC interburst
interval of several minutes, however, Drosophila's
PT,CO2 would exceed the critical threshold value of
4–6 kPa several-fold, suggesting that the fruit fly must allow
continuous gas exchange through the thoracic spiracles even during rest.
Nevertheless, we should keep in mind that this conclusion, among the other results in this study, may critically depend on the simple assumption that respiratory gas exchange in Drosophila relies on diffusion alone and can be described by the simple analytical model above. This approach neglects any dynamics resulting from both buffering of respiratory gases and tracheal ventilation. In addition, there are potential errors associated with our measurement technique including the difficulty to temporally match locomotor performance and CO2 release rate of the flying animal. The outcome of this study should thus be regarded with care and direct measurements of tracheal partial pressures at the various experimental conditions have still to verify our predictions.
Significance of spatial distribution of tracheal gas exchange areas on muscle function and lift production
One of the most unexpected results of the present study is the finding that
muscle efficiency changes only slightly in response to manipulations of the
spatial distribution of spiracle exchange areas. The pronounced relative
difference in muscle efficiency of 143±60.3% at 0.29 relative flight
force production appears to be an exception and probably is partly due to the
fly's CO2 buffering capacity (Figs
6B,
10). Nevertheless, we found a
trend in the data set suggesting that relative muscle efficiency is inversely
correlated with spiracle diffusive area
(Fig. 6B). The same trend is
also visible in the second muscle physiological parameter, i.e. mechanical
power output (Fig. 6A). Note
that these results run counter to the results on the relative lift coefficient
(Fig. 6C), relative drag
coefficient (data not shown) and relative aerodynamic efficiency
(Fig. 6D), which taken together
show a significant decrease in magnitude with decreasing spiracle exchange
area. In sum, the findings above suggest that changes in the spatial
distribution of tracheal oxygen supply due to local blocking of individual
spiracles negatively affect the ability of the animal to produce flight force
– but reinforce the production of mechanical power and the efficiency of
the mechano-chemical conversion process of the indirect flight
musculature.
The results above are rather surprising because in unmanipulated fruit
flies, muscle efficiency decreases with decreasing flight force production
(Lehmann, 2002). It has been
suggested that low muscle efficiency either reflects a decreased crossbridge
activation between actin and myosin filaments or an unfavourable strain regime
of the asynchronous flight muscle
(Josephson, 1999;
Josephson et al., 2001;
Lehmann and Dickinson, 1997).
Therefore, we originally hypothesized that at flight forces below maximum
performance, an inhomogeneous supply of oxygen to the IFM would even reinforce
this attenuation in crossbridge activation. Instead, our data apparently show
that the geometry of the tracheal system and the location of gas exchange
areas (spiracles) are of minor importance for IFM overall efficiency. A
possible explanation for this finding might be the fact that the large air
sacs of the dorsal and lateral tracheal system [pleural-, notopleural-,
lateroscutal- and medioscutal sacs
(Demerec, 1965)] homogenize
oxygen concentration within the fly body. However, if this explanation is
true, it still remains puzzling why muscle mechanical power output so strongly
depends on the changes in the arrangement of spiracle conductance.
We also unexpectedly found that relative mean lift coefficient and
aerodynamic efficiency differ up to approximately 50% between flies facing
alterations of oxygen supply distribution and unmanipulated animals
(Fig. 6C,D). By contrast,
kinematic variables such as stroke amplitude were affected relatively little,
and stroke frequency was even widely indistinguishable between unmanipulated
and spiracle-manipulated flies at the various flight forces (1.44±11.7
Hz, mean difference ± s.d., N=5 forces,
Fig. 3,
Table 1). Previous studies on
the mechanisms of unsteady aerodynamics in flapping insect wings have shown
that flight force production linearly decreases with decreasing wing velocity,
thus following conventional aerodynamic laws
(Ellington, 1984a;
Lehmann and Dickinson, 1998).
Apparently, wing velocity (the product between stroke amplitude and frequency)
in unmanipulated animals decreases to a greater extent than in flies in which
flight force reduction is forced by a reduction in diffusive exchange area. As
a consequence, mean lift coefficient in unmanipulated flies varies less
dramatically with changing flight forces than in the spiracle-manipulated
animals (Fig. 3E). From these
findings, we hypothesize that alterations in the spatial distribution of gas
exchange areas ultimately alter the fine structure of wing motion such as the
angle of attack, the velocity profile during wing translation or the wing's
rotational speed and timing during the stroke reversals. For example, it has
been shown that in a robotic model of Drosophila, changes in wing
rotation may alter both flight costs and lift production
(Dickinson et al., 1999;
Sane and Dickinson, 2001). An
8% advanced rotational timing, during which the wing rotates prior to the
stroke reversal, may reinforce aerodynamic force production by more than 70%
of total force, compared to a delayed rotation that occurs at the beginning of
each half stroke (Dickinson et al.,
1999). Somewhat smaller increases in lift production have been
reported for increases in the wing's angular velocity during rotation
(Sane and Dickinson, 2002).
Changes in rotational speed and timing, moreover, may alter the benefit of the
wake capture mechanism and clap-and-fling lift enhancement that also
contribute to Drosophila's high lift coefficient
(Lehmann et al., 2005).
Besides the changes in relative lift coefficient, the findings in Figs
3F,
4B,
5B suggest that changes in the
arrangement of spiracle conductance may also decrease the ratio between lift
and drag coefficients and thus increase the relative profile power
requirements for wing flapping. However, since lift and drag forces are
vectors of total flight force, an attenuation in lift would likely cause a
relative decrease in drag rather than a relative increase. The same
3-dimensional robotic model wing of Drosophila mentioned above has
demonstrated a tremendous increase in wing drag towards higher angles of
attack (Dickinson et al.,
1999; Usherwood and Ellington,
2002). While at angles of attack <45° lift and drag
coefficients are positively correlated, lift and drag coefficient are
inversely correlated at angles >45°. Increases in angle of attack above
this threshold could thus explain, for example, why the mean drag coefficient
may relatively increase while the mean lift coefficient decreases during
flight (Lehmann, 2004). A
detailed reconstruction of the wing kinematic pattern by means of high-speed
video technique should allow us to tackle this hypothesis for the underlying
aerodynamic mechanisms in the future.
The clap-and-fling mechanism for lift enhancement supposedly may not
contribute to the relative change of lift-to-drag ratio in
Drosophila, because its occurrence changes the lift-to-drag ratio
only slightly, from 0.57 to 0.58
(Ellington, 1975;
Lehmann et al., 2005;
Weis-Fogh, 1973). Altogether,
the results suggest that respiratory gas exchange based on the usage of
multiple thoracic and abdominal spiracles appears to be beneficial for
maintaining an elevated efficacy of aerodynamic force production in the fruit
fly. The ultimate explanation for this finding might be that changes in the
arrangement of spiracle conductance alter the contraction dynamics or muscle
stiffness of the IFM and thus the movements of the mechanical thoracic
oscillator (Josephson, 1999;
Josephson et al., 2001;
Josephson and Stokes, 1999;
Vigoreaux, 2001;
Vigoreaux et al., 2000). These
parameters were not covered by our measurement technique.
An alternative explanation
Alternatively, we should consider whether the above results are simply due
to the different mechanisms used by the flies for modifying flight force
production. While unmanipulated flies `voluntarily' altered flight forces in
response to the vertical motion of the visual lift stimulus, the
spiracle-blocked flies probably reduced flight force due to the restriction in
mechanical power output of the flight musculature. Consequently, our findings
can be interpreted such that changes in wing kinematics mediated by
active control of flight steering muscles produce more favourable
stroke kinematics than a passive change via mechanical power
limits (Götz, 1983;
Heide and Götz, 1996).
Numerous kinematic and electrophysiogical studies have shown that 17 flight
control muscles (steering muscles) tune several aspects of wing motion during
manoeuvring flight, such as stroke amplitude, stroke frequency, the timing of
wing rotation, angle of attack or the wing trajectory in Drosophila
(Götz, 1983;
Heide and Götz, 1996;
Lehmann and Götz, 1996)
(for a review, see Dickinson and Tu,
1997). In this case, the data plotted in
Fig. 6 would highlight the
energetic and aerodynamic consequences of wing kinematic alterations due to
different flight control strategies rather than reflect the significance of
respiratory constraints. It appears to be difficult to distinguish
unambiguously between both interpretations in our experiments because changes
in the arrangement of spiracle conductance also involve changes in total
diffusive area. However, the comparison between gas flux through the meso- and
metathoracic spiracles should be noted as an exception
(Fig. 7). As already mentioned,
the mean difference in the performance measures of 26% approximately matches
the difference in spiracle opening area between meso- and metathoracic
spiracles, which suggests a negligible 2.4% difference in oxygen supply rate
between the meso- and or metathoracic spiracle. However, due to the large
ipsilateral tracheal trunks and air sacs, this ipsilateral effect was expected
to be small a priori (Miller,
1950).
Conclusions
The in-depth evaluation of the significance of tracheal gas exchange in
Drosophila potentially provides several new insights onto how the
spatial distribution and the size of spiracle exchange areas determine the
function of the flight motor in a flying insect. The present results provide
direct evidence for the general assumption in respiratory research that the
tracheal development of a simple diffusion-based system matches the
respiratory need at maximum metabolic activity of the animal. Under those
conditions, respiratory water loss would be minimal, which in turn prevents
water stress on animals living in xeric environments
(Lighton, 1994;
Lighton, 1996). Moreover, our
findings apparently show that changes in the arrangement of spiracle
conductance primarily effect aerodynamic phenomena in addition to flight
muscle mechanical power output, but not predominantly muscle efficiency. The
exact reason why relative muscle mechanical power output depends more strongly
on the spatial distribution of spiracle areas than muscle efficiency remains
unknown and will require further research on the indirect flight musculature
in the behaving animal. Since it has been assumed that insects have no
anaerobic capacity, the magnitude of oxygen and CO2 buffer capacity
might play a crucial role in breathing behaviour and spiracle control in the
fruit fly (Ziegler, 1985). On
the one hand, CO2 buffer capacity may explain why spiracles have to
open immediately after flight initiation and match their opening area to
metabolic need (Lehmann,
2001). On the other hand, the small CO2 buffer capacity
might also partly explain why inbred lines of Drosophila reared on
commercial food do not exhibit a clear DGC pattern during resting metabolism
(Williams and Bradley, 1998).
Eventually, in conjunction with the small safety margin for tracheal
respiration at maximum locomotor capacity, we conclude that
Drosophila may apparently maximize the efficiency of its locomotor
system for flight by well-balancing respiratory gas flow between the four
large spiracles in the fly's thorax.
|
List of symbols and abbreviations |
|---|
|
TOPSummaryIntroductionMaterials and methodsResultsDiscussionList of symbols and...References |
|---|
A
M
T
|
Acknowledgments |
|---|
|
References |
|---|
|
TOPSummaryIntroductionMaterials and methodsResultsDiscussionList of symbols and...References |
|---|
Bailey, L. (1954). The respiratory currents in
the tracheal system of the adult honey bee. J. Exp.
Biol. 31,589
-593.
Barton, B., Ayer, G., Heymann, N., Maughan, D. W., Lehmann,
F.-O. and Vigoreaux, J. O. (2005). Flight muscle
properties and aerodynamic performance of Drosophila expressing a
flightin gene. J. Exp. Biol.
208,549
-560.
Birch, J. M. and Dickinson, M. H. (2001). Spanwise flow and the attachment of the leading-edge vortex on insect wings. Nature 412,729 -733.[CrossRef][Medline]
Burkett, B. and Schneiderman, H. A. (1974).
Role of oxygen and carbon dioxide in the control of spiracular function in
Cecropia pupae. Biol. Bull. Mar. Biol. Lab. Woods
Hole 147,274
-293.
Casey, T. M. (1989). Oxygen consumption during flight. In Insect Flight (ed. G. J. Goldsworthy and C. H. Wheeler), pp. 257-272. Boca Raton: CRC Press.
Casey, T. M. and Ellington, C. P. (1989). Energetics of insect flight. In Energy Transformations in Cells and Organisms (ed. W. Wieser and E. Gnaiger), pp.200 -210. Stuttgart: Thieme.
de Sanctis, D., Dewilde, S., Vornhein, C., Pesce, A., Moenz, L.,
Ascenzi, P., Hankeln, T., Burmester, T., Ponassi, M., Nardini, M. et
al. (2005). Bis-histidyl heme hexacoordination, a key
structural property in Drosophila melanogaster hemoglobin.
J. Biol. Chem. 280,27222
-27229.
Demerec, M. (1965). Biology of Drosophila. New York, London: Hafner Publishing Company.
Dickinson, M. H. and Lighton, J. R. B. (1995).
Muscle efficiency and elastic storage in the flight motor of Drosophila.Science 268,87
-89.
Dickinson, M. H. and Tu, M. S. (1997). The function of Dipteran flight muscle. Comp. Biochem. Physiol. 116A,223 -238.[Medline]
Dickinson, M. H., Lehmann, F.-O. and Sane, S.
(1999). Wing rotation and the aerodynamic basis of insect flight.
Science 284,1954
-1960.
Duncan, F. D. and Byrne, M. J. (2002).
Respiratory airflow in a wingless dung beetle. J. Exp.
Biol. 205,2489
-2497.
Ellington, C. P. (1975). Non-steady-state aerodynamics of the flight of Encarsia formosa. In Swimming and Flying in Nature, vol.2 (ed. T. Y. Wu, C. J. Brokaw and C. Brennan), pp.783 -796. New York: Plenum Press.
Ellington, C. P. (1984a). The aerodynamics of insect flight. IV. Aerodynamic mechanisms. Philos. Trans. R. Soc. Lond. B Biol. Sci. 305,79 -113.
Ellington, C. P. (1984b). The aerodynamics of insect flight. VI. Lift and power requirements. Philos. Trans. R. Soc. Lond. B Biol. Sci. 305,145 -181.
Ellington, C. P., Machin, K. E. and Casey, T. M. (1990). Oxygen consumption of bumblebees in forward flight. Nature 347,472 -473.[CrossRef]
Götz, K. G. (1983). Bewegungssehen and flugsteuerung bei der fliege Drosophila. In BIONA-report 2 (ed. W. Nachtigall), pp. 21-34. Stuttgart: Fischer.
Gulinson, S. L. and Harrison, J. F. (1996). Control of resting ventilation rate in grasshoppers. J. Exp. Biol. 199,379 -389.[Abstract]
Hadley, N. F. and Quinlan, M. C. (1993). Discontinuous carbon dioxide release in the Eastern lubber grasshopper Romalea guttata and its effect of respiratory transpiration. J. Exp. Biol. 177,169 -180.[Abstract]
Hankeln, T., Jaenicke, V., Kiger, L., Dewilde, S., Ungerechts, G., Schmidt, M., Urban, J., Marden, M. C., Moens, L. and Burmester, T. (2005). Characterization of Drosophila hemoglobin. Evidence for hemoglobin-mediated respiration in insects. J. Biol. Chem. 277,29012 -29017.
Harrison, J. F. and Roberts, S. P. (2000). Flight respiration and energetics. Annu. Rev. Physiol. 62,179 -205.[CrossRef][Medline]
Harrison, J. F., Hadley, N. F. and Quinlan, M. C. (1995). Acid–base status and spiracular control during discontinuous ventilation in grasshoppers. J. Exp. Biol. 198,1755 -1763.[Medline]
Heide, G. and Götz, K. G. (1996). Optomotor control of course and altitude in Drosophila is achieved by at least three pairs of flight steering muscles. J. Exp. Biol. 199,1711 -1726.[Abstract]
Josephson, R. K. (1999). Dissecting muscle power output. J. Exp. Biol. 202,3369 -3375.[Abstract]
Josephson, R. K. and Stokes, D. R. (1999). The force–velocity properties of a crustacean muscle during lengthening. J. Exp. Biol. 202,593 -607.[Abstract]
Josephson, R. K., Malamud, J. G. and Stokes, D. R.
(2001). The efficiency of an asynchronous flight muscle from a
beetle. J. Exp. Biol.
204,4125
-4139.
Kestler, P. (1985). Respiration and respiratory water loss. In Environmental Physiology and Biochemistry of Insects (ed. K. H. Hoffmann), pp.137 -186. Berlin: Springer.
Krogh, A. (1913). On the composition of air in the tracheal system of some insects. Skand. Arch. Physiol. 29,29 -36.
Lehmann, F.-O. (1999). Ambient temperature affects free-flight performance in the fruit fly Drosophila melanogaster.J. Comp. Physiol. B 169,165 -171.[CrossRef][Medline]
Lehmann, F.-O. (2001). Matching spiracle
opening to metabolic need during flight in Drosophila.Science 294,1926
-1929.
Lehmann, F.-O. (2002). The constraints of body size on aerodynamics and energetics in flying fruit flies: an integrative view. Zoology 105,287 -295.[Medline]
Lehmann, F.-O. (2004). Aerial locomotion in flies and robots: kinematic control and aerodynamics of oscillating wings. Arthropod Struct. Dev. 33,331 -345.[CrossRef]
Lehmann, F.-O. and Dickinson, M. H. (1997). The changes in power requirements and muscle efficiency during elevated force production in the fruit fly, Drosophila melanogaster. J. Exp. Biol. 200,1133 -1143.[Abstract]
Lehmann, F.-O. and Dickinson, M. H. (1998). The
control of wing kinematics and flight forces in fruit flies
(Drosophila spp.). J. Exp. Biol.
201,385
-401.
Lehmann, F.-O. and Dickinson, M. H. (2001). The production of elevated flight force compromises flight stability in the fruit fly Drosophila. J. Exp. Biol. 204,627 -635.[Abstract]
Lehmann, F.-O. and Götz, K. G. (1996). Activation phase ensures kinematic efficacy in flight-steering muscles of Drosophila melanogaster. J. Comp. Physiol. 179,311 -322.
Lehmann, F.-O. and Heymann, N. (2005).
Unconventional mechanisms control cyclic respiratory gas release in flying
Drosophila. J. Exp. Biol.
208,3645
-3654.
Lehmann, F.-O., Dickinson, M. H. and Staunton, J. (2000). The scaling of carbon dioxide release and respiratory water loss in flying fruit flies (Drosophila spp.). J. Exp. Biol. 203,1613 -1624.[Abstract]
Lehmann, F.-O., Sane, S. P. and Dickinson, M. H. (2005). The aerodynamic effects of wing–wing interaction in flapping insect wings. J. Exp. Biol. 208,2075 -3092.
Lighton, J. R. B. (1988). Discontinuous
CO2 emission in a small insect, the formicine ant Camponotus
vicinus. J. Exp. Biol. 134,363
-376.
Lighton, J. R. B. (1994). Discontinuous ventilation in terrestrial insects. Physiol. Zool. 67,142 -162.
Lighton, J. R. B. (1996). Discontinuous gas exchange in insects. Annu. Rev. Entomol. 41,309 -324.[CrossRef][Medline]
Lighton, J. R. B., Fukushi, T. and Wehner, R. (1993a). Ventilation in Cataglyphis bicolor: regulation of carbon dioxide release from the thoracic and abdominal spiracles. J. Insect Physiol. 39,687 -699.[CrossRef]
Lighton, J. R. B., Garrigan, D. A., Duncan, F. D. and Johnson, R. A. (1993b). Spiracular control of respiratory water loss in female alates of the harvester ant Pogonomyrmex rugosus. J. Exp. Biol. 179,233 -244.[Abstract]
Lipp, A., Wolf, H. and Lehmann, F.-O. (2005).
Walking on inclines: energetics of locomotion in the ant Camponotus.J. Exp. Biol. 208,707
-719.
Machin, J., Kestler, P. and Lampert, G. J.
(1991). Simultaneous measurements of spiracular and cuticular
water loss in Periplaneta americana: implications for whole-animal
mass loss studies. J. Exp. Biol.
161,439
-453.
Manning, G. and Krasnow, M. A. (1993). Development of the Drosophila tracheal system. In The Development of Drosophila Melanogaster (ed. M. Bate and A. M. Arias), pp. 609-686. Cold Spring Harbor: Cold Spring Harbor Laboratory Press.
Miller, A. (1950). The internal anatomy and histology of the imago of Drosophila melanogaster. In Biology of Drosophila (ed. M. Demerec), pp.420 -534. New York: John Wiley.
Miller, P. L. (1966). The supply of oxygen to
the active flight muscles of some large beetles. J. Exp.
Biol. 45,285
-304.
Miller, P. L. (1981). Ventilation in active and in inactive insects. In Locomotion and Energetics in Arthropods (ed. C. F. Herreid II), pp.367 -390. New York: Plenum.
Miller, P. L. (1982). Respiration. In The American Cockroach (ed. H. J. Bell and K. G. Adiyodi), pp. 87-116. London: Chapman & Hall.
Nikam, T. B. and Khole, V. V. (1989). Insect Spiracular Systems. New York, Chichester, Brisbane, Toronto: Ellis Horwood.
Quinlan, M. C. and Hadley, N. F. (1993). Gas exchange, ventilatory patterns, and water loss in two lubber grasshoppers: quantifying cuticular and respiratory transpiration. Physiol. Zool. 66,628 -642.
Ramamurti, R. and Sandberg, W. C. (2001). Computational study of 3-D flapping foil flows. In 39th Aerospace Sciences Meeting and Exhibit, pp.605 . Reno, NV: AIAA.
Sane, S. and Dickinson, M. H. (2001). The
control of flight force by a flapping wing: lift and drag production.
J. Exp. Biol. 204,2607
-2626.
Sane, S. and Dickinson, M. H. (2002). The
aerodynamic effects of wing rotation and a revised quasi-steady model of
flapping flight. J. Exp. Biol.
205,1087
-1096.
Schneiderman, H. A. (1960). Discontinuous
respiration in insects: role of the spiracles. Biol.
Bull. 119,494
-528.
Slama, K. (1994). Regulation of respiratory acidemia by the autonomic nervous system (coelopulse) in insects and ticks. Physiol. Zool. 67,163 -174.
Slama, K. (1999). Active regulation of insect respiration. Ann. Entomol. Soc. Am. 92,916 -929.
Snyder, G. K., Sheafor, B., Scholnick, D. and Farrelly, C. (1995). Gas exchange in the insect tracheal system. J. Theor. Biol. 172,199 -207.[CrossRef][Medline]
Usherwood, J. R. and Ellington, C. P. (2002).
The aerodynamics of revolving wings II. Propeller force coefficients from
mayfly to quail. J. Exp. Biol.
205,1565
-1576.
Vigoreaux, J. O. (2001). Genetics of the Drosophila flight muscle myofibril: a window into the biology of complex systems. BioEssays 23,1047 -1063.[CrossRef][Medline]
Vigoreaux, J. O., Moore, J. R. and Maughan, D. W. (2000). Role of the elastic protein projection in stretch activation and work output of Drosophila flight muscles. In Elastic Filaments of the Cell (ed. Granzier and Pollack), pp. 237-250. London, Amsterdam: Kluwer Academic/Plenum Publishers.
Wasserthal, L. T. (2001). Flight-motor-driven
respiratory air flow in the hawkmoth Manduca sexta. J. Exp.
Biol. 204,2209
-2220.
Weis-Fogh, T. (1964a). Diffusion in insect wing
muscle, the most active tissue known. J. Exp. Biol.
41,229
-256.
Weis-Fogh, T. (1964b). Functional design of the tracheal system of flying insects as compared with the avian lung. J. Exp. Biol. 41,207 -227.[Abstract]
Weis-Fogh, T. (1973). Quick estimates of flight
fitness in hovering animals, including novel mechanisms for lift production.
J. Exp. Biol. 59,169
-230.
Wigglesworth, V. B. (1972). The Principles of Insect Physiology. London: Methuen (Chapman & Hall).
Williams, A. E. and Bradley, T. J. (1998). The effect of respiratory pattern on water loss in desiccation-resistant Drosophila melanogaster. J. Exp. Biol. 201,2953 -2959.[Abstract]
Ziegler, R. (1985). Metabolic energy expenditure and its hormonal regulation. In Environmental Physiology and Biochemistry of Insects (ed. K. H. Hoffmann), pp.95 -118. Berlin, Heidelberg, New York, Tokyo: Springer.
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Reddit 
Technorati 
Twitter What's this?
This article has been cited by other articles:
|
|
 |
|
  W. A. Van Voorhies Metabolic function in Drosophila melanogaster in response to hypoxia and pure oxygen J. Exp. Biol., October 1, 2009; 212(19): 3132 - 3141. [Abstract] [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
) in performance between unmanipulated flies and animals in which up
to 4 thoracic spiracles have been sealed during flight. The differences are
scaled to the performance of the unmanipulated control group. Performance
scores are plotted against total diffusive area of the animal's abdominal and
thoracic spiracles that may participate in tracheal gas exchange. Due to the
reduction of maximum flight force production with decreasing total spiracle
opening area, data are calculated at 0.29 (493 µm2 total
spiracle area), 0.48 (3780 µm2), 0.74 (5424 µm2),
1.01 (7889 µm2) and 1.36 (10 355 µm2) relative
flight force production for 0–4 thoracic spiracles open, respectively. A
value of 1.0 normalized force means that the fly produces a flight force equal
to body weight. Grey areas in the pictograms indicate maximum total spiracle
opening area available for respiratory gas exchange and red lines indicate
linear regression fits. See legend of 
