The hyperoxic switch: assessing respiratory water loss rates in tracheate arthropods with continuous gas exchange
1 University of Nevada at Las Vegas, Department of Biology, University of
Nevada at Las Vegas, 4505 Maryland Parkway, NV 89154-4004 USA
2 Section of Ecology, Behavior and Evolution, Division of Biological
Sciences, University of California at San Diego, 9500 Gilman Drive, La Jolla,
CA 92093-0116 USA
* Author for correspondence (e-mail: john{at}johnlighton.org)
Accepted 13 September 2004
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Summary |
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Key words: Drosophila melanogaster, gas exchange, water loss, cuticular permeability, Pogonomyrmex californicus, Forelius mccooki
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Introduction |
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Water loss rate (WLR), and thus other parameters such as cuticular
permeability that depend on the accurate measurement of WLR, are usually
measured gravimetrically or via radioactive isotopes. This makes
distinguishing respiratory from cuticular water loss rates difficult or
impossible. It is possible to calculate although with a wide margin of
error how much overall gas exchange is required to sustain a given
catabolic flux rate, and then calculate the amount of water lost through this
avenue by making assumptions about exchange volumes (e.g.
Lighton and Feener, 1989).
Some insects make the process easier by limiting most of their respiratory
water loss to discrete `bursts' that can be separated from intervening periods
of cuticular water loss by sensitive real-time gravimetric techniques
(Machin et al., 1991
;
Lighton, 1992
and references
therein) or by water vapor analysis
(Hadley and Quinlan, 1982
;
Lighton et al., 1993
;
Gibbs et al., 2003
;
Gibbs and Matzkin, 2001
), but
such insects may be the exception rather than the rule (review by
Lighton, 1996
; but see
Chown, 2002
). And in the case
of very small insects, this approach, even where possible in theory, is
impractical because the resolution of current instrumentation is finite. This
means, for example, that the relation between respiratory and cuticular water
loss rate in such a widely studied organism as Drosophila
melanogaster is currently uncertain because its gas exchange is generally
continuous and its water vapor signal is barely detectable
(Gibbs et al., 2003
).
In a recent paper, Lehmann
(2001) examined respiratory
water loss rates in flying Drosophila by using water vapor as a
tracer gas for tracheal ventilation, as was earlier shown in the far larger
honeybee by Joos et al.
(1997
). Lehmann demonstrated
for the first time that the degree of spiracular opening during flight is
finely modulated to provide just-sufficient oxygen delivery and carbon dioxide
release, without imposing a needless respiratory water loss penalty. This
approach worked well because of the huge respiratory gas exchange rates during
flight, coupled with the ability to modulate flight intensity at will using a
virtual-reality scenario (see also
Dickinson and Lighton, 1995
).
However this approach is not practical in resting Drosophila, which
exchange respiratory gases not only at a far lower rate than in flight, but
also at a constant rate, although some partial exceptions may occur
(Gibbs et al., 2003
;
Williams and Bradley,
1998
).
In this paper we propose a novel approach to estimating minimal rates of
respiratory water loss (RWL) even in small insects that continuously exchange
respiratory gases. Our approach exploits the fact that the degree of
spiracular opening in insects at rest is modulated by the partial pressures of
both oxygen and carbon dioxide
(Wigglesworth, 1935; Levy and
Schneiderman,
1966a
,b
;
and also see reviews by Kestler,
1985
; Lighton,
1996
). In steady state, a low and relatively constant
intra-tracheal partial pressure of oxygen is maintained to facilitate inward
diffusion of oxygen from the atmosphere. If the intra-tracheal
PO2 falls, the degree of spiracular opening
rises; if intra-tracheal PO2 rises, the degree
of spiracular opening falls. Our approach replaces the air surrounding the
insect under study with pure oxygen, briefly elevating inward oxygen diffusion
rates by fivefold. The result was a transient period of increased
intra-tracheal PO2 and thus reduced the degree
of spiracular opening accompanied by a depression in CO2 and
respiratory water vapor output. By using CO2 as the tracer gas, and
by using a sensitive flow-through water vapor analyzer to detect any
accompanying change in water vapor flux, we can then estimate rates of
respiratory water loss. As an added benefit the water vapor analysis (in
conjunction with accurately known flow rates) allows real-time water loss
rates to be measured with an accuracy approaching that of gravimetric methods,
especially in small insects, without the disruption introduced by periodic
weighing.
Measurement of RWL during steady-state CO2 output is of
particular interest in species that are also capable of engaging in a
discontinuous gas exchange cycle or DGC. The selective advantage of the
stringent spiracular control required for the DGC is usually claimed to be a
reduction in RWL relative to continuous, usually primarily diffusion-based,
gas exchange, but this argument has recently been challenged
(Lighton and Berrigan, 1995;
Lighton, 1998
and references
therein). Direct comparisons of RWL between members of the same species in the
presence or absence of the DGC have not, to our knowledge, been made to date
and offer a means to evaluate the classic `hygric' or RWL-reduction hypothesis
(Lighton, 1998
) for the
evolution of the DGC in insects and certain other tracheate arthropods.
Respiratory water loss at rest is an important parameter. Perhaps less
important, but still interesting, is the maximal rate of respiratory water
loss. This is a function of total tracheal system area and maximal degree of
spiracular opening, and it is reasonable to assume that it will scale with
maximal aerobic capacity. By replacing the oxygen, in turn, with nitrogen, the
maximal rate of respiratory water loss through diffusion can also be measured.
(It is possible that during activities, such as locomotion, that the maximal
rate of respiratory water loss may be enhanced by active mechanisms, such as
convection.) Measurement of this parameter is facilitated because the response
to anoxia of an insect continuously exchanging respiratory gases is a rapid
and maximal increase in spiracular opening (see
Lighton and Fielden, 1996;
Klok et al., 2002
). The
resulting efflux of water vapor can then be directly measured and compared to
resting respiratory and cuticular water loss rates.
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Materials and methods |
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Foragers of Forelius mccooki and Pogonomyrmex
californicus were collected from the University of California Elliot
Chaparral Reserve and the Cuyacama Mountains, San Diego County, California,
respectively. They were maintained in round polyethylene containers containing
test tubes with water reservoirs backed by cotton wool 13 weeks before
testing. The test tubes were wrapped in aluminum foil to block ambient light.
Workers were maintained at ambient indoor temperatures (23±3°C) and
ambient photoperiod (Northern Hemisphere fall). They were fed on crickets and
sugar water ad libitum. Drosophila melanogaster were obtained from
wild-type stocks in the laboratory of Stephen Roberts at the University of
Nevada at Las Vegas. On arrival, the fly vials were opened and the flies were
liberated. After 24 h the newly eclosed flies were decanted into a new vial,
stored as above, and used for measurement on days 34 after eclosion.
Flies were not sexed prior to measurement to minimize handling stress. The fly
groups were approximately 50% of each sex (the ants, of course, were all
female). Sex does not appear to significantly influence mass- or area-specific
water loss rates in D. melanogaster
(Gibbs and Matzkin, 2001 and
references therein). Flies were reared with media until experiments on water
loss were conducted.
Respirometry
Most metabolic and water-loss studies are conducted separately by
stopflow (constant volume) and gravimetric techniques, respectively.
Both techniques have poor temporal resolution. Stopflow techniques
yield integrated averages of catabolic flux rates over periods of an hour or
more, during which bursts of activity may lead to serious measurement
overestimates (Lighton et al.,
2001 and references therein). Likewise, gravimetric studies cannot
isolate mass-loss events unrelated to cuticular or respiratory water loss,
such as excretion or salivation during grooming, which is a particular problem
with Drosophila. Regular weighing also disturbs animals. By contrast,
we used flow-through respirometry for all measurements. Although more
demanding of instrumentation stability and resolution, flow-through
respirometry minimizes these errors
(Lighton, 1991
), and also
allows the evaluation of hypotheses that require high temporal resolution for
experimental testing.
We used a Sable Systems International
(www.sablesystems.com)
TR-2 system (SSI; Las Vegas, Nevada, USA) for flow-through respirometry with a
LiCor CO2 analyzer (Lincoln, NE, USA; resolution <0.1
p.p.m. CO2), supplemented by an SSI RH-100 water vapor analyzer
with direct readout in Pa water vapor pressure (1 Pa of resolution or better
and 1% accuracy), SSI's Datacan V data acquisition software with UI-2 16-bit
measurement interface (basic accuracy 0.03%), and SSI ExpeData data analysis
software. A Tylan FC-260 mass flow control valve attached to SSI's two-channel
mass flow controller (TR-MFC1) controlled flow rates. Specimen temperatures
were controlled to ±0.1°C by an SSI Pelt-4 temperature controller
coupled to a SSI PTC-1 Peltier Effect cabinet with an internal volume of 8 l.
The CO2 analyzer was zeroed with CO2-scrubbed outside
air and spanned at 976 p.p.m. with a certified span gas. The water vapor
analyzer was zeroed with nitrogen and spanned by bubbling air through pure
water at an accurately known temperature 5°C lower than ambient,
setting the RH-100 to its dewpoint mode, and adjusting it to read the correct
water temperature.
Our system is diagramed in Fig. 1. Bev-A-Line low-permeability tubing was used throughout to minimize water vapor adsorbance errors (Thermoplastic Processes Inc., Georgetown, DE, USA). Briefly, air from outside the lab building was pulled by an SSI TR-SS1 sub-sampler through a Drierite/Ascarite/Drierite drying column (Ascarite; Thomas Scientific, Swedesboro, NJ, USA) to remove CO2 and H2O, then into the mass flow control valve, which was set to an STP-corrected flow rate appropriate to the organism (20 ml min1 for Forelius and Pogonomyrmex, or 40 ml min1 for Drosophila). After the mass flow control valve, the gas flow passed through a second column containing Ascarite and magnesium perchlorate, which dries air to a greater extent than Drierite and helped to stabilize the water vapor baseline. The prepared air then entered the temperature cabinet, which was set to 20 or 40°C, passed through a helical aluminum temperature equilibration coil, and then entered the respirometry chamber. Because of the small size of Forelius mccooki, we employed a 4 cm length of 3 mm I.D. low-permeability Bev-A-Line tubing as the chamber. For the other animals we employed a sealed glass/metal chamber with an interior volume of about 3 ml (SSI TRRM). Finally, air left the respirometry chamber (having gathered CO2 and H2O from the insects on its way), entered the RH-100 water vapor analyzer, and then traveled to the CO2 analyzer.
|
As outlined in the Introduction, we could at any time switch pure nitrogen
or oxygen for the dry air that normally passed through the system. This was
accomplished by disconnecting the normal drying column, and instead allowing
the system's pump to pull gas from a manifold into which either dry oxygen or
dry nitrogen flowed at a rate of approximately 100 ml min1.
During a typical hyperoxic switch run, a recording was started that
established the baselines for the carbon dioxide and water vapor analyzers.
Meanwhile a group of 10 Forelius or a single Pogonomyrmex
was gathered and weighed to 0.01 mg with a Mettler AG245 balance
(Mettler-Toledo Inc., Columbus, OH, USA). After weighing the recording was
paused and the animals were placed in the respirometry chamber. A built-in
stainless steel filter disk prevented the smaller animals from creeping from
the chamber into the system's tubing. The flow rate through the system was
increased to 100200 ml min1 for 35 min to
speed equilibration, and then lowered to the appropriate flow rate (see above)
for a further 510 min before re-starting the recording. The recording
continued until a plateau in water loss rate was reached. At that point the
gas flowing through the system was changed to pure oxygen. After a further
20 min, the oxygen was changed to nitrogen, and the recording continued
until the animals' carbon dioxide production fell to near baseline levels.
Finally, the recording was paused, the animals were removed from the chamber,
the chamber was replaced, and after equilibration another baseline taken.
We weighed Drosophila and recorded their water vapor and CO2 output in the same manner as that used for the ants except for the following. First, we cooled flies for easier handling, and then placed groups of 411 individuals in the respirometric chamber to record CO2 and water vapor emission. After that, flies were weighed and the mass of water lost during the recording (as determined from the respirometry data) was added to their post-recording weight.
Each recording consisted of a variable number of data points taken at 2 s
intervals, using finite impulse response digital filtration to reduce
short-term noise (Lighton,
1991). Recordings typically lasted for 1.52.0 h including
initial and final baselines.
Analysis and statistics
Hyperoxic switch recordings were analyzed using Sable Systems ExpeData
software (ß release 3.1). For each hyperoxic switch recording:
During these operations a master log file was recorded that spanned the
analysis of all of the data files. If required (for example, to add an
additional variable to the spreadsheet for each file), the master log file
could be edited and then played back, allowing all of the data files to be
re-analyzed very rapidly and without operator error. When complete, the data
in the spreadsheet were saved as a delimited ASCII file with column headings.
Once the data were in the spreadsheet, further data manipulations could be
performed. These included the calculation of water vapor pressure saturation
deficit from chamber temperature (formula in
Lighton and Feener, 1989) and
the calculation of ant surface area (ibid) and, thence, cuticular
permeability.
Recordings of animals engaging in the DGC (i.e. P. californicus at
20°C) were analyzed as described elsewhere
(Lighton et al., 1993). Means
are accompanied by N and S.E. unless otherwise noted.
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Results |
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Calculation of total vs respiratory water loss rates
The hyperoxic switch technique we used in this study did, as predicted,
caused a transient drop in carbon dioxide output (see Figs
2,
3,
4 and
Table 1). The decline in carbon
dioxide emission also caused a temporary decrease in water loss rate. Thus at
steady-state, assuming diffusive gas exchange with fixed trans-spiracular
partial pressure gradients:
![]() | (1) |
![]() | (2) |
Exposure to pure oxygen produced short-term decreases in
CO2 of
4688% (Table 1). If the
partial pressure gradients for water and carbon dioxide did not change during
these periods, then these decreases in
CO2 were caused
by decreases in A/L of 4688%. Thus, the simplest estimate of
trans-spiracular
H2O would be
![]() | (3) |
It is, however, possible to correct the
H2O data for the
limited temporal resolution of the system and to assign an upper limit to
respiratory water loss rates. This correction requires three assumptions.
First, it is certainly reasonable to assume a ratiometric relation between the
hyperoxic declines in
CO2 and
H2O, as shown in
Equations 1 and
2, bearing in mind that
CO2 comes only from the spiracles while H2O comes from
both the spiracles and the cuticle. The second assumption is that for a short
time, the degree of spiracular opening is reduced to very low levels after
exposure to hyperoxia in fact P. californicus shows a decline
of nearly 90%. We will assume a transient decline to zero. By this reasoning,
we should increase the hyperoxic
H2O decline by
the ratio of (100%:hyperoxic
CO2 decline%).
If this assumption is reasonable, then we can assign an upper limit to
respiratory water loss rates, as is shown in
Table 1. So, assuming no
changes in internal PCO2 or
PH2O:
![]() | (4) |
However, as noted above, this estimate of maximal trans-spiracular
H2O assumes no
changes in the partial pressure gradients for water or carbon dioxide. This is
the third assumption of this model. While it seems likely that tracheal air
stays saturated with water within the tracheae during hyperoxia, it is
possible that in these insects, as in grasshoppers
(Gulinson and Harrison, 1996
),
that hyperoxia induces a rise in tracheal PCO2.
As can be seen from Equation 2,
if
PCO2 doubled in response to
hyperoxia, a 50% drop in
CO2 would be
associated with a 100% reduction in A/L. We attempted to minimize this problem
by using measurements of
H2O and
CO2 from only
the initial minutes after exposure to hyperoxia.
It is trivial to modify Equation 4 to allow for a less than complete reduction in the degree of spiracular opening across the hyperoxic switch; this assumption simply allows us to set an upper limit to the contribution of respiratory water loss to total water loss.
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Discussion |
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The hyperoxic switch
We are not aware of any previously published report of a reduction in
CO2 following
exposure to hyperoxia in any insect. However, a senior study by Wigglesworth
(1935
) in the flea, showed the
modulation of spiracular opening by gas composition. Particularly, in his
Fig. 4H
(Wigglesworth, 1935
), it is
clear that the closed phase of the last abdominal spiracle lasts for a longer
time in pure oxygen; however, the transient effects of hyperoxia on
CO2 have not
previously been reported. Levy and Schneiderman
(1966a
) did not cite the
Wigglesworth's 1935
study, but
they demonstrated a similar phenomenon. With silkworm pupae, they observed an
increase in the period of spiracular constriction with increasing ambient
O2 (PO2=60%), and hypothesized a
theoretical spiracular response in pure O2. We suggest that the
hyperoxic switch effect is widespread and could be a useful tool for arthropod
respiratory physiologists. This method may be especially applicable for small
arthropods with continuous gas exchange and for which gas exchange occurs
primarily by diffusion, which obviously appears to be the case in our study
from the kinetics of gas exchange shown in our graphs.
As alluded to in the Results, our raw measurements may underestimate the effects of the hyperoxic switch on the degree of spiracular opening. This is because of the limited temporal resolution of our respirometry system, chiefly in the analyzers rather than in the flow characteristics of our respirometer chambers at the flow rates we used. We propose (see Results) a simple correction to mitigate this effect and to allow calculation of an upper limit to respiratory water loss rates. Direct visual observation of spiracular activity during the hyperoxic switch would provide valuable information regarding the validity of our correction technique.
Total water loss rates
Our respirometrically measured rates of total water loss in D.
melanogaster are similar to those of Gibbs and Matzkin
(2001;
42 µg
h1 fly1, their
Fig. 3), and the gravimetric
measurements of Lehmann et al.
(2000
; 30 µg
h1 fly1, their Table 2). The overall water
loss rate of D. melanogaster divided by estimated surface area and
water vapor pressure deficit yields its gross cuticular permeability (i.e.
including the respiratory component). This value, again, is similar to the
value that can be calculated from published data (ibid.).
The WLR of F. mccooki has not been previously described but is
typical for a xeric ant (Duncan and
Lighton, 1994). That of P. californicus is far higher on
an area-specific basis than that of the other two insects, but this is to be
expected in view of its higher measurement temperature (40°C vs
20°C). When corrected for water vapor pressure saturation deficit, its
gross cuticular permeability is lower than, but not dissimilar to, that of two
congeners, P. rugosus and P. occidentalis (mean 28.4 µg
h1 cm2 Torr1; where Torr
133.3 Pa; Quinlan and Lighton,
1999
).
Respiratory water loss rates during continuous CO2 emission
Among the ants we measured, respiratory water loss rates varied from
57% of total water loss rates, which is similar to values measured
in ants that express a DGC; 2% (Camponotus vicinus;
Lighton, 1992
); 8%
(Cataglyphis bicolor; Lighton,
1992
); 4% (Pogonomyrmex occidentalis, which is very
similar to P. californicus in body size and morphology;
Quinlan and Lighton, 1999
) and
2% (Pogonomyrmex rugosus; Quinlan
and Lighton, 1999
). They are well within the range for insects in
a variety of other orders for which estimates exist in the literature (see
especially Table 1 in Chown,
2002
).
Where Drosophila melanogaster is concerned, there are very few
published estimates of respiratory water loss in animals at rest. As Gibbs et
al. (2003) bemoan,
`Unfortunately, water loss from individual flies in our experiments was so low
that we could not measure it reliably... Carbon dioxide readings never reached
zero, indicating that at least one spiracle remained open at all times, which
would also reduce our ability to distinguish respiratory losses. Thus, we were
unable to reliably detect increases in water loss caused by spiracular
opening...' Williams and Bradley
(1998
) were fortunate enough,
however, to find of the 50 flies they examined two that
exhibited sufficient discontinuity in CO2 emission to allow them to
assess respiratory water loss, which they estimated at
25% of total water
loss rate. This is very similar to our figure, which can therefore be
provisionally accepted as a reasonable figure for non-flying D.
melanogaster.
We were somewhat struck by the fact that the
H2O trace often
appeared to stay below pre-hyperoxic levels, whereas the
CO2 trace
invariably returned to pre-hyperoxic levels. It is possible that this may
reflect a lasting reduction in the degree of spiracular opening caused by
hyperoxia, in conjunction with a higher internal
PCO2 that establishes itself at a level
sufficient to maintain normal rates of CO2 emission through the
constricted spiracles. Then again, most of the
H2O traces
showed a long-term downward trend, making firm statements about long-term
changes in
H2O
problematic. This was another reason why we restricted our measurements to the
section of the
H2O trace
immediately coincident with the
CO2 hyperoxic
decline (time difference only 5 min or less on average). For D.
melanogaster, which showed the strongest long-term downward trend in
H2O, we
restricted our measurements to an interval of only 3.19±1.05 min (mean
± S.E.; N=9) immediately following the hyperoxic
switch. This was because water excretion events often took place shortly after
the effects of hyperoxia manifested themselves (see
Fig. 3). We are at a loss to
explain this apparent behavioral effect either of hyperoxia or, conceivably,
of moderate endotracheal hypercapnia caused by spiracular constriction.
In any event we conclude that the hyperoxic switch technique has potential utility to students of the respiratory physiology of tracheate arthropods.
Respiratory water loss rates during the DGC
The RWL of P. californicus, expressed as a percentage of total
WLR, did not change whether or not it expressed a DGC
(Table 1). This is an
interesting result, and serves to caution that simplistic evolutionary
hypotheses regarding the evolution of the DGC, e.g. the long-held `hygric
hypothesis', may not hold water (Lighton,
1998 and references therein; with respectful apologies to Allen
Gibbs). It should be mentioned here that hypotheses regarding the
RWL-reduction benefits of the DGC should be exclusively fielded under the
banner of the hygric hypothesis; the chthonic hypothesis
(Lighton, 1998
) is exclusively
concerned with gas exchange under severe hypoxic or hypercapnic conditions,
not with what may (or may not) occur in conditions where partial pressures of
respiratory gases are more quotidian, as in CO2-scrubbed air.
We have no reason to believe that our sample of ants was expressing the DGC
abnormally. The DGCs and catabolic flux rates of our sample of P.
californicus were almost identical to those reported by Quinlan and
Lighton (1999), compensating
where necessary for the 5°C higher temperature of the ants in that study.
In addition their catabolic flux rates are very similar to those predicted for
motionless ants of that mass and temperature by the allometric equation of
Lighton et al. (2001
);
P=0.25, see Table
1.
However, the WLRs of P. californicus during continuous and
discontinuous CO2 emission are not as directly comparable as one
might wish, calling for a note of interpretative caution. This is because the
temperatures of the ants expressing the two gas exchange strategies differed
by 20°C. This temperature disparity should not affect the ratio of
respiratory to total water loss rates, especially because WLR (as expressed in
Table 1) is also standardized
by the water vapor pressure saturation deficit at each temperature. WLR per
unit water vapor pressure saturation deficit is generally independent of
temperature (Edney, 1977;
Lighton and Feener, 1989
and
references therein). Catabolic flux rate is strongly temperature dependent,
increasing some fourfold from 20 to 40°C, and should elevate RWL at the
higher temperature. However, this effect (of uncertain magnitude with regard
to RWL, accompanied by unknown changes in the degree of spiracular opening)
would increase RWL to total WLR ratios at the higher temperature, so it would
make the present comparison more conservative. This is because the RWL to
total WLR ratio would be expected to be higher at 40°C, exaggerating any
putative reductions in RWL caused by the DGC at 20°C, a temperature at
which RWL should decrease anyway because of decreased catabolism. This makes
the parity reported here of RWL as a percentage of total WLR, during
continuous and discontinuous CO2 emission, all the more
interesting, although it would be preferable to turn the DGC on or off at a
single temperature and within a single individual while monitoring WLR. Such
studies could go some way towards resolving the controversies surrounding the
selective correlates of the DGC (Lighton,
1998
; Chown, 2002
;
and references in both).
Response to anoxia
A large increase in CO2 emission followed exposure to nitrogen
(Figs 2,
3,
4;
Table 1), which was caused by
the opening of the insects' spiracles after detection of anoxia. However, the
biological meaning of this peak is difficult to interpret in this context
because it consists of material (CO2) that requires O2
for its production, but O2 was something that the insects, in their
case, no longer had. (For this reason, convective gas exchange caused, for
example, by abdominal pumping, would rapidly cease during anoxia.) The
CO2 peak therefore consisted largely of carbon dioxide escaping
from the tracheal system and hemolymph, steadily decreasing after the initial
peak with classic first-order-decay kinetics. This contrasts with water vapor
output, which in the short term is independent of oxygen availability and is,
therefore, a better indicator, for present purposes, of total respiratory
throughput capacity. Not surprisingly, D. melanogaster, as an insect
capable of flight, showed the largest anoxic factorial increase in
H2O, but F.
mccooki was not far behind. To anyone who has watched F. mccooki
in the field this comes as no surprise, because this tiny ant forages at high
speed at high substrate temperatures
(Hölldobler and Wilson,
1990
; J.R.B.L., personal observations) and is therefore presumably
capable of high aerobic throughput. Ants have elsewhere been shown to be
capable of attaining factorial aerobic scopes close to those required for
flight (Roces and Lighton,
1995
).
The episode of peak water loss following exposure to nitrogen is brief,
because, we hypothesize, the terminal tracheoles rapidly lose their small
reservoirs of free water in the absence of metabolic activity
(Wigglesworth, 1983). After
the reservoirs evaporate, water diffuses into the tracheoles from the
hemolymph and then diffuses through the tracheal system and out through the
spiracles at a much lower, and more constant, rate. Other possible explanation
for the fall in water vapor rates after a period of time in anoxia could be
the cessation of some active process that is facilitating the high rates of
water loss.
It is reasonable to infer from the far higher maximal rate of water vapor loss through the spiracles immediately after exposure to nitrogen (see above), that the rate-limiting step in this phase of the anoxia response is the diffusion of water across the tracheolar membranes, rather than spiracular and tracheal resistance. Thus, following the initial brief peak of water loss, there follows a lower plateau at an approximate steady state composed of cuticular plus tracheal water loss (not strictly respiratory water loss because respiration has ceased). We assume this steady state to represent the steady flux of water vapor from the tracheoles, following the wholesale loss of water-vapor-saturated air from the endotracheal space during the initial stages of anoxia.
Measuring H2O
and
CO2 kinetics
after exposure of tracheate arthropods to anoxia may therefore yield
non-invasive and, if of short duration, non-lethal comparative information on
maximal degree of spiracular opening, tracheal volume, tracheal conductance
and summed tracheolar surface areas.
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Acknowledgments |
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References |
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