|
| |
|
|
|
|
||||
| Home Help Feedback Subscriptions Archive Search Table of Contents | ||||
First published online November 28, 2008
Journal of Experimental Biology 211, 3790-3799 (2008)
Published by The Company of Biologists 2008
doi: 10.1242/jeb.018721
Haemoglobin as a buoyancy regulator and oxygen supply in the backswimmer (Notonectidae, Anisops)
Ecology and Evolutionary Biology, School of Earth and Environmental Sciences, Darling Building, University of Adelaide, Adelaide, SA 5005, Australia
* Author for correspondence (e-mail: philip.matthews{at}adelaide.edu.au)
Accepted 11 September 2008
 |
Summary |
|---|
 TOP Summary INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
Key words: buoyancy, haemoglobin, insect, respiration
|
INTRODUCTION |
|---|
|
TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
; Rahn and Paganelli,
1968). However, carrying a volume of air sufficient for underwater
respiration dramatically affects an insect's ability to stay submerged.
Insects have an overall body density slightly greater than water, because the
density of their tissues is about 1.05 g cm–3 and that of
their chitinous cuticle is about 1.3 g cm–3
(Vincent and Wegst, 2004).
This makes them slightly negatively buoyant. Thus the buoyant force of even a
comparatively small bubble of air causes an insect to float. When positively
buoyant, it must actively swim to submerge, and can only remain underwater by
continuing to swim or clinging to the substrate. But while submerged, the
volume of its bubble decreases as respiration consumes oxygen and both carbon
dioxide and nitrogen diffuse out into the surrounding water. As a result the
insect steadily becomes less buoyant, eventually to the point where it begins
to sink and must swim to the surface to refill its air-store. This constant
change in buoyancy has prevented almost all aquatic insects from successfully
occupying the mid-water zone, instead limiting them to foraging on the bottom
or floating at the surface.
The aquatic backswimmers (Hemiptera: Notonectidae, subfamily Anisopinae)
are unique among insects as they carry a bubble of air on their abdomen while
submerged and yet are capable of achieving a prolonged period of neutral
buoyancy during a dive. Backswimmers are a common sight in still, often
stagnant bodies of water, from farm dams to swimming pools. They can be found
floating motionless in the water column, occasionally moving in rapid bursts,
using their oar-like hind legs. Backswimmers are also remarkable in that they
belong to one of only three insect families known to produce substantial
quantities of haemoglobin, the other two being the larvae of Gastrophilus
intestinalis (Oestridae) and Chironomus spp. (Chironomidae)
(Weber and Vinogradov, 2001).
The backswimmer produces haemoglobin in large modified fat-body cells within
its abdomen (Bergtrom et al.,
1976). An extensive network of tracheoles pervades each
haemoglobin cell, and the air-filled tracheoles connect with the abdominal
spiracles through larger tracheae (Bare,
1929). Two parallel grooves run along the length of the abdomen's
ventral surface and contain both the abdominal spiracles and the air-store
(Figs 1 and
2). Thus the haemoglobin cells
are intimately associated with both the tracheal system and the air-store
via the abdominal spiracles. Unlike many diving insects that maintain
maximum contact between the surface of their air-stores and the surrounding
water for reasons of gas exchange [e.g. Aphelocheirus
(Thorpe and Crisp, 1947)],
backswimmers cover their air-stores with a layer of hydrophobic hairs. These
long hairs fringe the outer edge of each groove and, due to their
hydrophobicity, arrange themselves in a layer across the air–water
interface of the air-store when the backswimmer is submerged, and also stick
to the water surface when the backswimmer surfaces, thus exposing the ventral
spiracles directly to the atmosphere (Fig.
2).
|
|
).
He observed that backswimmers swam beneath the water's surface, where they
appeared to be `in perfect equilibrium', and considered that the haemoglobin
probably acted as a supplementary oxygen store to extend their time
underwater. Miller was the first to suggest that the oxygen released from the
haemoglobin during a dive was linked to the backswimmers' buoyancy
(Miller, 1964;
Miller, 1966). He observed
that the bright red oxyhaemoglobin present in Anisops pellucens at
the beginning of a dive slowly changed to a dark purple once the insect had
reached neutral buoyancy, indicating that oxygen was released from the
haemoglobin and was contributing to respiration during the neutrally buoyant
phase. In separate experiments, he fumigated them with air mixtures containing
6–25% carbon monoxide (CO), thus preventing their haemoglobin from
binding with oxygen. Backswimmers treated in this manner did not enter the
neutrally buoyant phase and passed directly from positive to negative
buoyancy. Although these experiments clearly indicate a relationship between
haemoglobin and buoyancy, they suffer from their reliance on qualitative
visual estimates (e.g. haemoglobin oxygen saturation determined by comparison
with colour standards), and estimation of buoyancy by examining the movements
of free-swimming backswimmers.
Despite the rarity of insect haemoglobin, there have been surprisingly few
studies on its function from a whole-animal perspective. Subsequent studies on
backswimmer haemoglobin have focused more on the biochemical aspects of
synthesis (Bergtrom et al.,
1976), structure (Bergtrom,
1977; Osmulski et al.,
1992; Vossbrinck et al.,
1993) and oxygen-binding properties
(Wells et al., 1981). These
studies examined the functioning of haemoglobin in vitro, extracting
it from large numbers of homogenised backswimmers. Only Wells and colleagues
discussed the relationship between the oxygen affinity of haemoglobin and its
physiological role (Wells et al.,
1981). Their in vitro analysis of Anisops
assimilis haemoglobin showed a very steep oxygen equilibrium curve, with
oxygen released readily only at low oxygen partial pressure
(PO2). They hypothesized that the backswimmer's
respiration would cause the PO2 and volume of
its positively buoyant air-store to decrease during a dive. With the
PO2 of the air-store reduced, the haemoglobin
could then unload its oxygen, temporarily stabilising the volume of the now
neutrally buoyant air-store. However, the insect's buoyancy, the air-store
PO2 and the relative contribution of the
haemoglobin to submerged respiration were not measured.
Progress in this area has been impeded by an inability to measure the gas
composition and volume of the air-store in these small insects. However, this
study, which expands on previously published experiments
(Matthews and Seymour, 2006),
uses new techniques to quantitatively assess the in vivo function of
haemoglobin in the backswimmer Anisops deanei (Brooks 1951). Fibre
optic oxygen probes were used to directly measure the partial pressure of
oxygen within the air-stores of submerged bugs, while measurements of buoyancy
were made on tethered backswimmers using a sensitive electronic balance. These
measurements were then used to determine the role of haemoglobin in buoyancy
control and respiration.
|
MATERIALS AND METHODS |
|---|
|
TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
), and were collected from ponds at the North Terrace campus
and Waite campus of Adelaide University, Adelaide, South Australia. They were
kept in the laboratory in a 31.5 l aquarium illuminated by a fluorescent
aquarium light on a 12 h:12 h light:dark cycle. Water temperature was
maintained at 20°C by circulating water from a temperature-controlled
water bath (Thermomix 1442 D, B. Braun, Melsungen, Germany) through a glass
coil submerged in the aquarium. The backswimmers were fed on a diet of locally
caught live mosquito larvae, cladocerans and copepods. All experiments were
carried out at 20°C.
Determination of body density
The density of Anisops deanei was determined according to
Archimedes' principle. Fifteen specimens were collected from outdoor ponds and
transported to the laboratory, where they were transferred into vials
containing cotton wool soaked in chloroform, and killed. They were gently
blotted with tissue paper to remove any water, weighed to
1x10–5 g, and then transferred into individually
numbered 5 ml vials. Each vial was filled with water, and a small square of
soft nylon mesh was inserted to prevent the insect from floating to the
surface. All air adhering to the backswimmers' bodies and in their tracheal
systems was removed by placing the uncapped vials in a desiccator jar
connected to a vacuum pump (H. I. Clements Pty, Sydney, Australia). A suction
pressure of –80 kPa was applied to the desiccator jar, which was then
sealed for 16 h. After this period any bubbles adhering to the backswimmers
were removed by gently tapping the vials. The negatively buoyant insects were
then sucked into a 1 ml disposable syringe and transferred under water to a
custom-made weighing pan hanging in a 4 cm deep Petri dish of water below an
AE 163 balance (Mettler, Greifensee, Switzerland). Their submerged weight was
determined to 1x10–5 g. Body volumes were then
determined according to the formula:
 | (1) |
H2O is the density of pure water at the
temperature used in the measurement (g cm–3). The density of
the backswimmers (b gcm–3) was then
calculated by:
 | (2) |
Determination of initial air-store volume
The volume of air carried from the surface of the water by free-swimming
backswimmers was measured by removing the gas and measuring the volume
directly (Scholander and Evans,
1947). An upright funnel with nylon mesh blocking its throat was
submerged inside a 1 l beaker filled with air-equilibrated water. A
backswimmer was dried on blotting paper, weighed on an AE 163 electronic
balance, then placed within the funnel and allowed approximately 15 min to
settle and spontaneously dive. A `capture tube' was made from a 4 ml test-tube
that was filled with kerosene and inverted in a second beaker of water, such
that the buoyant kerosene remained in the tube. After the backswimmer
submerged, the funnel was partially lifted out of the first beaker. The
suction created by the water draining from the funnel held the insect
transiently against the nylon mesh. The capture tube was transferred to the
funnel by holding a thumb over its opening, and quickly placed over the
still-submerged insect. The backswimmer's hydrophobic air-holding surfaces
were flooded with kerosene on contact, displacing the air, which floated to
the top of the test-tube. The bug was then crushed against the wall of the
capture tube with a spatula, forcing the remaining air from the tracheal
system. The trapped air bubble was drawn into a length of 0.86 mm internal
diameter polyethylene tubing and then expelled into an inverted 0.5 cm
internal diameter glass cup filled with kerosene. With a micromanipulator
under a dissecting microscope, the bubble was then transferred to a
horizontally mounted micrometre burette constructed and operated according to
the design of Scholander and Evans
(Scholander and Evans,
1947).
|
Pieces of 0.1 mm diameter iron wire, approximately 16 mm long, were bent at right angles half-way along their length. One arm of the wire was then doubled back to form a U shape. Backswimmers were narcotised with CO2, gently blotted on tissue paper, and weighed. The U part of the wire was then dipped in a drop of cyanomethacrylate adhesive (Selleys Pty) and fixed lengthwise along the back of the immobilised backswimmer, with the unbent arm oriented posteriorly and perpendicularly away from it (Fig. 3). The unbent arm of the wire was then inserted into a sheet of foam, thus holding the backswimmer on its back for 10 min while the adhesive set. During this time the backswimmer was placed either in air or in a 60 ml glass syringe ventilated with 15% CO, 20% O2 and 65% N2 standard temperature and pressure (STP) at a rate of 63 ml min–1 from a gas-mixing apparatus custom-built from mass flowmeters (model GFC171; Aalborg Instruments and Controls, Orangeburg, NY, USA) regulated by a PC running control software through a D/A converter (ProfessorDAQTM and PowerDAQTM PD2-AO, United Electronic Industries, Walpole, MA, USA). Backswimmers treated with CO were submerged in 20°C water that had been aerated with the same gas mixture; otherwise the water used was air equilibrated.
While CO binds readily to haemoglobin, it also has the potential to inhibit
aerobic metabolism by binding to cytochrome oxidase
(Ball et al., 1951). However, a
pilot study demonstrated that oxygen partial pressure in the air-stores of
CO-treated backswimmers initially declined in the same manner as in control
insects, thus demonstrating aerobic respiration. For CO to depress aerobic
respiration it must reduce the number of available cytochrome oxidase enzymes
below that required to support a particular level of aerobic activity. So
while CO may have depressed the backswimmers' maximum possible rate of oxygen
consumption, it would not have depressed their resting metabolic rate. This is
in agreement with previous studies that show insects are remarkably tolerant
of CO. For example, in Miller's study on Anisops, the dive durations
of backswimmers exposed to 6%, 12% and 25% CO were not significantly different
as would be expected if aerobic respiration was depressed
(Miller, 1966). Studies on
stick insects and beetles show they can survive for over 30 days in 20% CO and
for up to 10 days in atmospheres containing 80% CO
(Baker and Wright, 1977). And
at the extreme, adult silkworm hearts exposed to 4 atm (
404 kPa) of CO
pressure continue to show aerobic function, with complete aerobic inhibition
achieved at 5 atm (505 kPa) (Harvey
and Williams, 1958). We therefore conclude that 15% CO did not
inhibit aerobic metabolism.
Using the unbent wire arm as a handle, the backswimmer was moved to the water-filled container beneath the balance with a pair of anti-magnetic pointed tweezers. It was half-submerged head first at approximately 25 deg. until the hairs fringing the abdominal air-store caught on the surface tension of the water. It was then fully submerged. While still submerged, the backswimmer was released 1 cm away from, and slightly above, the magnet. The magnet readily attracted and held the insect, orienting it in the normal swimming position (horizontally, with its ventral surface uppermost). The combined weight of the submerged backswimmer, its air-store, and the wire and glue, was recorded every 2 s. Once the backswimmer attempted to surface, it was recovered from the magnet, and the recording stopped. The balance was left to settle and then re-tared. The glue and wire were then easily peeled off the backswimmer, allowing it to be weighed separately underwater.
Air-store volume calculation
The air-store volume at each 2 s interval was calculated according to the
formula:
 | (3) |
 | (4) |
Air-store PO2 measurement
Changes in air-store PO2 during a dive were
measured before and after a backswimmer had been exposed to CO. The bug was
narcotised in pure CO2 gas for 3 min and then weighed. While still
immobilised it was then affixed to a glass microscope slide on its back, using
a drop of cyanomethacrylate adhesive. The insect was then left in air to
recover for 10 min before measurements began.
A 2 cm deep Petri dish filled with air-equilibrated water at 20°C was placed on the stage of a dissecting microscope. The bug was half-submerged in the water until the long hydrophobic hairs fringing the abdominal grooves were caught on the surface tension of the water. The bug was then completely submerged, with a bubble of air held over the abdomen beneath the hairs. A syringe-mounted optical oxygen probe with a <50 µm diameter tip attached to an oxygen meter (TX3, PreSens GmbH, Regensburg, Germany) was then manoeuvred into the air-store using a micromanipulator. PO2 was then logged at 1 s intervals until the backswimmer attempted to surface, at which point recording was stopped and the animal was removed from the Petri dish. For the CO treatment, this procedure was repeated using the same backswimmer, but with the 10 min recovery period performed in a 60 ml glass syringe ventilated with 15% CO, 20% O2 and 65% N2 STP at a rate of 63 ml min–1 from the gas-mixing apparatus. The CO-treated bug was then submerged in water that had been aerated with the same gas mixture, and measurement of air-store PO2 was carried out during the dive as described above.
Effect of temperature and aquatic PO2 on voluntary dive duration
The dive durations of backswimmers in an aquarium maintained at different
temperatures and PO2 were measured.
Backswimmers naturally seek out conspecifics and form aggregations in the wild
(Bailey, 1987). It was observed
that backswimmers placed individually into the aquarium took far longer to
settle than when at least one other backswimmer was present. Thus in order to
obtain the most consistent and repeatable measurements, dive durations were
measured when the backswimmers were kept in pairs.
A 15.8l aquarium was vertically partitioned with a black sheet of plastic perforated at the top and bottom. A coil of glass tubing flushed with water from a temperature-controlled water bath was placed in the small compartment between the end wall of the aquarium and the plastic sheet, to maintain the aquarium water at the desired temperature. Compressed air or nitrogen was blown through an air stone next to the coil to control the PO2 of the water. Aquatic PO2 was monitored using an optical oxygen probe submerged in the aquarium connected to the TX3 oxygen meter. The back and sides of the dive chamber were covered with black plastic sheeting, leaving the front of the aquarium open for observation. Backswimmers were introduced to the large compartment in pairs and given 30 min to acclimatise, after which six consecutive dive events performed by each backswimmer were observed and timed using a stopwatch. Dive times were recorded in water PO2 treatments of 100%, 80%, 40% and 25% air saturation at 20°C, as well as in normoxic water at 10, 15, 20, 25 and 30°C.
Statistical analysis
All statistics were performed using the Microsoft ExcelTM add-in
StatistiXL version 1.6
(www.statistixl.com).
Data are given as the mean of N samples ±95% confidence
interval (CI).
|
RESULTS |
|---|
|
TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
b) of A.
deanei was determined to be 1.0784±0.0074 g cm–3.
This value was used in all calculations. Submerged backswimmer weight
calculated using this value is expected to lie within ±8.5% of the
actual submerged weight (i.e. within 95% CI limits).
Mechanisms of air-store volume regulation
The volume of air collected at the surface of the water is regulated
largely by the abdominal grooves on the backswimmer's abdomen and the long
hairs that fringe them (Fig.
2). When submerging the backswimmers for experimentation, it was
necessary to lower the bug into the water until its abdominal hairs spread
onto the surface tension of the water before submergence. Over-large bubbles
occurred if the backswimmer was submerged too quickly, and flooded grooves
occurred if it was submerged too slowly. Observation of backswimmers diving in
aquaria also revealed the importance of the abdominal hairs when surfacing.
Backswimmers refilled their air-stores in two distinct ways. The first
involved reversing, abdomen first, towards the surface, touching the tip of
the abdomen to the surface for a fraction of a second, and then diving
rapidly. This behaviour, which was the most frequently observed, allowed a
small quantity of fresh air into the air-store and a quick return to buoyant
equilibrium. The second surfacing behaviour involved the backswimmers placing
the entire ventral surface of their abdomen against the water's surface. This
caused the hairs to stick to the surface tension of the water, forming a dark
border resembling eyelashes on each side of the abdomen, while exposing the
abdominal grooves to the atmosphere (Fig.
2). The backswimmers rested against the water's surface for a
little over 1 s before rapidly diving with quick strokes of their oar-like
hind legs. This resulted in the hairs snapping back into place over the
air-filled grooves. Once submerged, it took approximately 20 s before the
insects again approached neutral buoyancy. Occasionally, after refilling its
air-store in this manner, a backswimmer submerged carrying too much air.
Inbetween swimming to keep submerged, the backswimmer would vigorously attempt
to wipe the excess air from its abdomen using its hind legs. Dislodging minute
bubbles of air reduced the insect's buoyancy and allowed it to assume its
normal position in the water column. If it was unsuccessful, it returned to
the surface to obtain a more suitable air volume.
|
b) of A.
deanei.
Air-store volume and buoyancy in experimental dives
The initial volume of air carried by each backswimmer was calculated from
the first stable weight once the balance had recovered from the disturbance of
having the backswimmer attached. To verify that the initial air-store volume
was determined accurately, the percentage decrease in initial volume that had
occurred by the time the backswimmer began vigorous surfacing attempts
(observed in the data from the electronic balance as rapid spikes in weight)
was calculated. This volume decrease was due primarily to consumption of
oxygen, which constitutes approximately 20% of the initial volume of air, and
a smaller amount of nitrogen diffusion into the water. Thus a backswimmer
attempting to surface and replenish its oxygen stores must do so after a
decrease in air-store volume of just over 20%. In 10 control and six
CO-treated backswimmers, decreases in air-store volume were 24.8±3.6%
and 26.8±5.7%, respectively. A two-tailed t-test found no
difference between the initial air-store volumes of control and CO-treated
backswimmers, and so these data were pooled. There were a few records where
volume appeared to drop more than 35%, and these were considered to represent
an underestimation of initial air volume, resulting from a shift in the tared
weight of the balance, a change in the meniscus effect on the support shaft or
loss of part of the bubble, and these data were excluded from the analysis.
Mean initial air-store volume was 0.11401±0.01259 ml
g–1 and 31.5±7.3% larger than the calculated volume
required for neutral buoyancy (Fig.
5).
|
CO significantly reduced the time taken for backswimmers to begin surfacing behaviour, with CO-treated backswimmers attempting to surface after only 372±57 s compared with 490±58 s in control bugs (one-tailed t-test: control dive duration > CO dive duration, P=0.006). As well as reducing dive duration, the manner in which air-store volume decreased during a dive was also affected. Both control and CO-treated backswimmers began a dive with a steep initial decline in air-store volume. In control treatments, this was then followed by a period of relative stability, with only a minor decrease in volume over several minutes, until a final drop triggered a burst of activity in the backswimmers as they attempted to surface (Fig. 6). In CO-treated backswimmers, this extended plateau was eliminated and air-store volume dropped continuously.
|
|
The progressive changes in air-store PO2
were divided into three phases (P1, P2 and P3) according to their relative
rates of decline. P1 corresponds to the initial linear
PO2 decrease, followed by the plateau (P2),
which terminates in a final PO2 drop (P3). The
precise change point between P1 and P2 was defined as the point on the
PO2 trace where the initial rate of air-store
PO2 change
(
O2) decreased
by half (Fig. 8, point a). The
transition from P2 to P3 was then defined as the point where the rapid
increase in O2
at the end of the plateau began to slow
(Fig. 8, point b). Using these
definitions, P1 and P2 were of nearly equal duration (243±48 and
271±55 s, respectively), with P2 beginning at 5 kPa and ending at 2 kPa
(Table 1). P2 showed a marked
decrease in O2
relative to P1, with the PO2 declining 12 times
more slowly during the plateau period (the middle two-thirds of P2) than at
the beginning of the dive. Oxygen released by the haemoglobin maintained the
PO2 at a mean value of 3.79 kPa during this
plateau (Table 1).
|
|
Effect of temperature and aquatic PO2 on voluntary dive duration
Aquatic PO2 was found to have no significant
effect (nested ANOVA P=0.260) on dive time
(Fig. 9) with the dive time at
100% air-saturation (mean 275±36 s) only slightly greater than that at
25% (248±19 s). Water temperature showed a significant effect (nested
ANOVA P=0.000) on dive duration. Dive time increased with decreasing
water temperature, with the line of least-squares regression having a slope of
–12.453 s °C–1 and R2=0.969
(Fig. 10).
|
|
|
DISCUSSION |
|---|
|
TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
).
Measurements of air-store PO2 showed the same,
three-phase pattern, where a steep, constant decline in air-store oxygen is
followed by an extended stable period before a rapid, final drop. This pattern
can be explained by the oxygen-binding characteristics of the backswimmer's
haemoglobin. At the beginning of every dive the insect's respiration consumes
oxygen largely from within the air-store alone, causing a constant decline in
PO2 (P1). Eventually, decreasing
PO2 causes the haemoglobin to unload its
oxygen. This oxygen supplements the oxygen being consumed from the air-store,
thus reducing the rate at which PO2 declines
(P2). But once bound oxygen becomes depleted, the
PO2 drops for a final time and the air-store
becomes severely hypoxic (P3). The PO2 traces
from CO-exposed bugs had none of these phases and instead showed a continuous
asymptotic decline in PO2
(Fig. 7).
There are important differences between voluntary dives of free backswimmers and the pattern of changes in buoyancy and PO2 in tethered dives, when the insects were inactive. The relative durations of the three phases of a dive must be affected by changes in the backswimmer's activity during a dive. Under natural conditions, insects must swim to remain submerged during the positive buoyancy of P1. This activity would increase their respiration rate and reduce the duration of this phase. However, the duration of P2 would not be expected to change, as swimming activity is greatly reduced. The third phase involves activity in both cases, but free insects are able to reach the surface within a few seconds.
Dive durations of forcibly submerged backswimmers were affected both by the method of submergence and by the method used to define surfacing activity. Animals glued to wire and held submerged on a magnet began actively attempting to surface after 490±58 s (control) and 372±57 s (CO treated). In comparison, bugs glued to microscope slides were considered to `surface' once the PO2 in their air-stores reached 2 kPa. By this determination control dives lasted 510±94 s and only 330±120 s following exposure to CO. Both methods showed reduced dive durations following CO exposure. However, as shown by the large confidence intervals, the variability of the air volumes carried by forcibly submerged insects resulted in variable surfacing times.
Air-store volume and buoyancy
The air-store carried by backswimmers serves as both an oxygen reserve and
a buoyancy device. For a bubble of air to allow a backswimmer to become
neutrally buoyant, (a) it must have a constant volume to confer a stable
buoyancy, and (b) the stable volume must produce a buoyant force that is
matched to the backswimmer's weight. These conditions cannot occur in a simple
air bubble, as the constant decline in PO2
caused by the insect's respiration would cause the bubble's volume and
buoyancy to drop continuously. Backswimmers temporarily overcome this problem
as oxygen is released from their haemoglobin cells in response to declining
PO2. This stabilises the
PO2 and, therefore, the volume of their
air-store. But because haemoglobin only responds to
PO2 and not to the total volume of the
air-store, it alone cannot guarantee that the stable volume will confer
neutral buoyancy. For this to occur, the insect must collect a specific volume
of air so that neutral buoyancy coincides with the
PO2 necessary to cause the haemoglobin to
unload its oxygen. This constrains the volume of air that a backswimmer can
collect from the surface.
The water-saturated air above the surface of a pond at 20°C is 20.5% oxygen, corresponding to a partial pressure of 20.77 kPa at 1 atm (101.3 kPa). Therefore, an insect's respiration can reduce the volume of a newly refilled air-store by a maximum of 20.5%, assuming all oxygen is consumed. Consequently, for a positively buoyant air-store to be reduced to neutral buoyancy requires that the initial air-store volume is less than 20.5% larger than that required for neutral buoyancy, as this reduction in volume would reduce the PO2 of the air-store to 0, thus making the dive unsustainable. The exact volume of air collected at the surface must be matched to the oxygen-binding properties of the backswimmer's haemoglobin, as the haemoglobin stabilises the air-store's volume over a specific range of PO2 values. The plateau phase (P2) of the air-store begins at a PO2 of 5 kPa and ends at 2 kPa. During this phase the haemoglobin readily unloads its oxygen into the air-store, maintaining the PO2 of the middle two-thirds of P2 at 3.79±0.16 kPa. Assuming that the initial air-store began with a PO2 of 20.77 kPa, and occupied 20.5% of the initial air-store volume, then these PO2 changes at the beginning, middle and end of the plateau correspond to decreases in initial air-store volume of 16.0%, 17.2% and 19.0%, respectively. From this it can be seen that an initial air-store volume about 17% larger than that required for neutral buoyancy would be necessary to result in neutral buoyancy at the PO2 levels at which haemoglobin releases its oxygen. Because of the relatively high oxygen affinity of the haemoglobin, it is therefore necessary for the bug to be positively buoyant for a considerable period at the beginning of a dive.
The mean volume of air collected by free-diving bugs was 6.7±4.2% larger than that required for neutral buoyancy (Fig. 4), compared with the forcibly submerged insects, which were calculated to begin the dive with a mean air-store volume 31.5±7.3% larger than that required for neutral buoyancy (Fig. 5). The air volumes collected by free-swimming backswimmers provide an insight into the behavioural mechanisms of buoyancy control used by these insects. The measured volume of air did not always equate to the initial volume, as the backswimmers usually tried to escape being captured in kerosene after diving and consequently must have decreased the volume of their air-stores by consuming oxygen. Since this was unavoidable, the largest air volumes (i.e. 16%–19%) are the most representative of the initially collected volumes of air. This supports the assertion that backswimmers should select initial air volumes approximately 17% larger than neutral buoyancy. This is further supported by the measured range of air-store volumes collected from free-diving backswimmers. Assuming all backswimmers collected a volume of air 17% larger than that required for neutral buoyancy and were then caught after consuming varying proportions of their oxygen stores (from none to all), then the volumes should vary around neutral buoyancy over a range equivalent to the initial 20.5% volume of oxygen in the air-store, i.e. from 17% above neutral buoyancy to 3.5% below it. This is comparable to the actual range of air volumes, which varied from 19.4% above neutral buoyancy to 4.9% below (Fig. 4).
Gas exchange with the surrounding water
A bubble of air held under water dissolves due to the hydrostatic pressure
of the water increasing the partial pressures of the gases it contains above
those in the surrounding water. If respiration is reducing the
PO2 within this bubble, then the partial
pressure of nitrogen (PN2) increases according
to Dalton's law, further increasing the gradient driving nitrogen loss. As
backswimmers control their buoyancy by regulating the volume of their
air-store, any volume change due to this nitrogen diffusion must be minimised.
Unlike other diving insects that can use their air-stores as gills (e.g.
Ege, 1915;
Vlasblom, 1970), backswimmers
achieve this by limiting the area of their air-store in contact with the
surrounding water. Not only are the grooves containing the air-store narrow,
but also they are fringed with hydrophobic hairs, which arrange themselves
across the air–water interface (Fig.
2), further reducing the area available for diffusion. The
efficiency of this arrangement in limiting diffusion is evident when examining
the effect of aquatic PO2 on dive duration.
Dive duration was not significantly affected by
PO2 (Fig.
9), indicating minimal exchange of oxygen between the air-store
and water. Similar experiments by Miller, Vlasblom, and Wells and colleagues
support this finding (Miller,
1966; Vlasblom,
1970; Wells et al.,
1981). As the Krogh's diffusion coefficient for nitrogen in water
at 20°C is 2.19 times smaller than that for oxygen
(Rahn and Paganelli, 1968), it
is concluded that the volume of nitrogen lost during an average 4 min dive in
air-equilibrated water is inconsequential.
Oxygen store volume
The only oxygen available to a submerged backswimmer is what it carries
from the surface in its air-store and that bound to its haemoglobin. The
volume of oxygen contributed by each of these two stores, as well as the
backswimmer's oxygen consumption rate
(
O2), can be
determined from the PO2 traces. This requires
that the respiration rates of the inactive backswimmers are constant during
measurement and any diffusion of oxygen into the air-store from the
surrounding water is insignificant. Because the volume of air held by the
submerged backswimmers was unknown during PO2
measurement, it was approximated from the mass-specific mean initial air-store
volume of forcibly submerged insects (114.01µlg–1). Under
these conditions the constant respiration of the insect would cause the
PO2 of the air-store to drop steadily to 0 kPa.
The time (t, h) taken to reach 0 kPa is:
 | (5) |
O2 is the rate
of PO2 change (kPa h–1)
obtained from the first 20 s of PO2
measurement. The initial air-store volume (Va, µl)
includes an amount of oxygen (VO2) equivalent to:
 | (6) |
O2 (µl
h–1) is:
 | (7) |
O2 of 3.84
µlh–1. This is in close agreement with the mean predicted
metabolic rate of a typical arthropod at rest (2.89 µlh–1)
calculated as
O2=127.7M
0.853b at 20°C, where Mb is body
mass (g) (Lighton et al.,
2001). The greatest potential error in these calculations arises
from the approximation of the initial volume of air carried by the
backswimmer.
The oxygen carried by a backswimmer within its air-store and haemoglobin is
never completely exhausted during a dive, because low
PO2 stimulates the insect to surface before its
air-store becomes completely anoxic. The amount of oxygen consumed before the
air-store PO2 drops below this hypoxia
threshold is the effective oxygen supply. The threshold was designated as 2
kPa because this PO2 coincided with an increase
in activity of the tethered backswimmers and is also the critical
PO2 of many insects – the point at which
the resistance of their tracheal system begins to limit the rate of oxygen
diffusion and they can no longer maintain a constant
O2
(Greenlee and Harrison, 2004).
The time taken for the PO2 trace to reach 2 kPa
(Fig. 8, point b) is the sum of
the time taken to consume the effective oxygen supply of both the air-store
and haemoglobin individually. In the absence of haemoglobin the time taken to
consume the air-store's oxygen alone would be proportional to the
backswimmer's
O2. Therefore,
the time taken to consume the air-store's effective oxygen supply was found by
extrapolating the initial rate of PO2 decline
to 2 kPa (Fig. 8, point c).
This time was then multiplied by the previously calculated
O2 to determine
the effective oxygen content of the air-store
(VO2a; Table
2). The difference in the time taken to reach point b and C
(t) is equivalent to the length of time the dive is sustained by
oxygen released from the haemoglobin. Thus the effective oxygen contribution
of haemoglobin (VO2Hb;
Table 2) is found by:
 | (8) |
|
Thus the air-store and haemoglobin possess virtually equal oxygen stores of
0.26±0.02 and 0.25±0.06 µl, respectively. A two-tailed
t-test revealed no significant difference (P=0.814) between
them. However, the oxygen contribution of the air-store is likely to be an
over-estimation because the measurements were made on insects that were
forcibly submerged, causing them to carry larger volumes of air than if they
had been free to surface and fill their air-store. The mean air volume
collected from freely submerging backswimmers was 32% smaller than the volume
carried by forcefully submerged insects. An air-store of this size would
contribute only 42% of the total oxygen respired during a dive, the remaining
58% being supplied by the haemoglobin. This suggests that under natural
conditions the backswimmers rely more on the oxygen supplied by their
haemoglobin to sustain their respiration while submerged and less on their
air-stores. Miller estimated that Anisops pellucens derived up to 75%
of the oxygen consumed during a dive from haemoglobin, 25% from the air-store,
and a negligible amount from the surrounding water
(Miller, 1966).
LIST OF ABBREVIATIONS
O2
O2
H2O
b
|
References |
|---|
|
TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
Andersen, N. M. and Weir, T. A. (2004). Australian Water Bugs: Their Biology and Identification, 344pp. Collingwood, Victoria: CSIRO Publishing.
Bailey, P. C. E. (1987). Abundance and age-specific spatial and temporal distribution in two waterbug species, Anisops deanei (Notonectidae) and Ranatra dispar (Nepidae) in three farm dams in South Australia. Oikos 49, 83-90.[CrossRef]
Baker, G. M. and Wright, E. A. (1977). Effects of carbon monoxide on insects. Bull. Environ. Contam. Toxicol. 17,98 -104.[CrossRef][Medline]
Ball, E. G., Strittmatter, C. F. and Cooper, O.
(1951). The reaction of cytochrome oxidase with carbon monoxide.
J. Biol. Chem. 193,635
-647.
Bare, C. O. (1929). Haemoglobin cells and other studies of the genus Buenoa (Hiemiptera, Notonectidae). Univ. Kansas Sci. Bull. 18,265 -349.
Bergtrom, G. (1977). Partial characterisation of haemoglobin of the bug, Buenoa confusa. Insect Biochem. 7,313 -316.[CrossRef]
Bergtrom, G., Gittelman, S., Laufer, H. and Ovitt, C. (1976). Haemoglobin synthesis in Buenoa confusa (Hemiptera). Insect Biochem. 6, 595-600.[CrossRef]
Ege, R. (1915). On the respiratory function of the air-stores carried by some aquatic insects (Corixidae, Dytiscidae, Notonecta). Z. Allg. Physiol. 17, 81-125.
Greenlee, K. J. and Harrison, J. F. (2004).
Development of respiratory function in the American locust Schistocerca
americana I. across-instar effects. J. Exp. Biol.
207,497
-508.
Harvey, W. R. and Williams, C. M. (1958).
Physiology of insect diapause. XII. The mechanism of carbon
monoxide-sensitivity and-insensitivity during the pupal diapause of the
Cecropia silkworm. Biol. Bull.
114, 36-53.
Hungerford, H. B. (1922). Oxyhaemoglobin present in the backswimmer, Buenoa margaritacea Bueno. Can. Entomol. 54,262 -263.
Lighton, J. R. B., Brownell, P. H., Joos, B. and Turner, R. J. (2001). Low metabolic rate in scorpions: implications for population biomass and cannibalism. J. Exp. Biol. 204,607 -613.[Abstract]
Matthews, P. G. D. and Seymour, R. S. (2006). Diving insects boost their buoyancy bubbles. Nature 441, 171.
Miller, P. L. (1964). Possible function of haemoglobin in Anisops. Nature 201, 1052.
Miller, P. L. (1966). The function of
haemoglobin in relation to the maintenance of neutral buoyancy in Anisops
pellucens (Notonectidae, Hemiptera). J. Exp.
Biol. 44,529
-543.
Osmulski, P. A., Vossbrinck, C. R., Sampath, V., Caughey, W. S. and Debrunner, P. G. (1992). Spectroscopic studies of an insect hemoglobin from the backswimmer Buenoa margaritacea (Hemiptera, Notonectidae). Biochem. Biophys. Res. Comm. 187,570 -576.[CrossRef][Medline]
Rahn, H. and Paganelli, C. V. (1968). Gas exchange in gas gills of diving insects. Respir. Physiol. 5,145 -164.[CrossRef][Medline]
Scholander, P. F. and Evans, H. J. (1947).
Microanalysis of fractions of a cubic millimeter of gas. J. Biol.
Chem. 169,551
-560.
Thorpe, W. H. and Crisp, D. J. (1947). Studies
on plastron respiration. II. the respiratory efficiency of the plastron in
Aphelocheirus. J. Exp. Biol.
24,270
-303.
Vincent, J. F. V. and Wegst, U. G. K. (2004). Design and mechanical properties of insect cuticle. Arthropod Struct. Dev. 33,187 -199.[CrossRef][Medline]
Vlasblom, A. G. (1970). The respiratory significance of the physical gill in some adult insects. Comp. Biochem. Physiol. 36,377 -385.
Vossbrinck, C. R., Osmulski, P. A. and Debrunner, P. G. (1993). Characterization of hemoglobin from the backswimmer Buenoa margaritacea (Hemiptera: Notonectidae) reveals high electrophoretic heterogeneity. Insect Biochem. Mol. Biol. 23,421 -429.[CrossRef]
Weber, R. E. and Vinogradov, S. N. (2001).
Nonvertebrate hemoglobins: functions and molecular adaptations.
Physiol. Rev. 81,569
-628.
Wells, R. M. G., Hudson, M. J. and Brittain, T. (1981). Function of the hemoglobin and the gas bubble in the backswimmer Anisops assimilis (Hemiptera: Notonectidae). J. Comp. Physiol. 142,515 -522.
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Reddit 
Technorati 
Twitter What's this?
Related articles in JEB:
This article has been cited by other articles:
|
|
 |
|
  K. Phillips BACKSWIMMERS REGULATE BUOYANCY WITH HAEMOGLOBIN J. Exp. Biol., December 15, 2008; 211(24): i - ii. [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
