Can Oxygen Set Thermal Limits in an Insect and Drive
Gigantism?
Wilco C. E. P. Verberk*, David T. Bilton
Marine Biology and Ecology Research Centre, University of Plymouth, Plymouth, Devon, United Kingdom
Abstract
Background: Thermal limits may arise through a mismatch between oxygen supply and demand in a range of animal taxa.
Whilst this oxygen limitation hypothesis is supported by data from a range of marine fish and invertebrates, its generality
remains contentious. In particular, it is unclear whether oxygen limitation determines thermal extremes in tracheated
arthropods, where oxygen limitation may be unlikely due to the efficiency and plasticity of tracheal systems in supplying
oxygen directly to metabolically active tissues. Although terrestrial taxa with open tracheal systems may not be prone to
oxygen limitation, species may be affected during other life-history stages, particularly if these rely on diffusion into closed
tracheal systems. Furthermore, a central role for oxygen limitation in insects is envisaged within a parallel line of research
focussing on insect gigantism in the late Palaeozoic.
Methodology/Principal Findings: Here we examine thermal maxima in the aquatic life stages of an insect at normoxia,
hypoxia (14 kPa) and hyperoxia (36 kPa). We demonstrate that upper thermal limits do indeed respond to external oxygen
supply in the aquatic life stages of the stonefly Dinocras cephalotes, suggesting that the critical thermal limits of such
aquatic larvae are set by oxygen limitation. This could result from impeded oxygen delivery, or limited oxygen regulatory
capacity, both of which have implications for our understanding of the limits to insect body size and how these are
influenced by atmospheric oxygen levels.
Conclusions/Significance: These findings extend the generality of the hypothesis of oxygen limitation of thermal tolerance,
suggest that oxygen constraints on body size may be stronger in aquatic environments, and that oxygen toxicity may have
actively selected for gigantism in the aquatic stages of Carboniferous arthropods.
Citation: Verberk WCEP, Bilton DT (2011) Can Oxygen Set Thermal Limits in an Insect and Drive Gigantism? PLoS ONE 6(7): e22610. doi:10.1371/
journal.pone.0022610
Editor: Alexander W. Shingleton, Michigan State University, United States of America
Received December 7, 2010; Accepted July 1, 2011; Published July 27, 2011
Copyright: ß 2011 Verberk, Bilton. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This study was funded by the Netherlands Organisation for Scientific Research (NWO-RUBICON fellowship no. 825.09.009). The funders had no role in
study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: wilco@aquaticecology.nl
damage to proteins, membranes and cells, and ultimately death
[2,3,6,7].
Although argued to hold for ectotherms in general [3], most of
the evidence for oxygen limitation at thermal extremes to date
comes from a variety of marine taxa, including fish, crustaceans,
bivalves, and annelids [3, 5 and references therein]. Recent studies
of terrestrial isopods and beetles [7,8] question the generality of
this mechanism, suggesting that upper and lower limits are
decoupled in terrestrial arthropods, and showing no increase in
critical thermal maximum (CTmax) with hyperoxia. Additionally,
whilst hypoxia decreased CTmax in isopods, an effect was only seen
at extreme hypoxia in beetles, casting doubt on the degree to
which oxygen limitation sets upper thermal limits in tracheates.
From a functional perspective, both the fact that the tracheal
system constitutes a single-step gas exchange system and their high
efficiency of oxygen delivery, would indeed seem to preclude
oxygen limitation as a major mechanism setting upper thermal
limits in terrestrial insects [8]. In addition, terrestrial insects can
maintain relatively constant internal oxygen levels, across steep
clines in external supply, by adjusting ventilation rates or through
compensatory developmental changes in tracheal length, diameter
or branching [9,10].
Introduction
To predict species responses to global warming trends, it is
paramount to understand the causal mechanisms underlying
thermal limits. The idea of oxygen limitation as a mechanism
setting upper thermal limits in animals was first expounded by
Winterstein [1] and has since been greatly expanded by Pörtner
and colleagues [2–5]. Recent work has extended this principle to
lower thermal limits, and sees both upper and lower critical
temperatures (CTmax and CTmin) being coupled by the common
mechanism of temperature-dependent oxygen limitation. As
ectotherms warm the demand for oxygen in their tissues increases
faster than the rate at which oxygen can be provided by cardiac
and ventilatory processes, leading to a drop in whole-animal
aerobic scope and a shift from aerobic to anaerobic metabolism.
Similarly on cooling, metabolism slows, leading to insufficient ATP
production in ventilatory muscles, reducing oxygen supply to
tissues. Deleterious thermal effects are hypothesized to set in first
at the whole-animal level, rather than lower hierarchical levels [3].
Thus, temperature-dependent oxygen limitation first lowers
whole-animal aerobic scope (and hence performance), followed
by the onset of anaerobic metabolism, finally resulting in heat
PLoS ONE | www.plosone.org
1
July 2011 | Volume 6 | Issue 7 | e22610
Oxygen, Thermal Limits and Insect Gigantism
Not all insect species and life-stages may be equally sensitive
to temperature-dependent oxygen limitation however and there
remains a need for additional studies across the range of
morphologies and lifestyles seen in extant insects [11]. In
particular, many insects have aquatic larval stages, where the
lower oxygen content and diffusion rates in water compared to
air dramatically reduce available oxygen [12,13], making
oxygen limitation more likely. In addition, larger species or
individuals may be more prone to temperature-dependent
oxygen limitation because of longer diffusion pathways [4].
Similarly, if body size ultimately constrains an animal’s capacity
to supply its tissues with oxygen, this would explain why animals
attain larger sizes in the cold, where metabolic rates are reduced
[13,14]. Such size based oxygen limitation is also central to the
leading hypothesis regarding insect gigantism in the late
Palaeozoic [15].
Here we provide the first formal test of oxygen limitation in a
freshwater tracheate, the stonefly Dinocras cephalotes (Curtis, 1827).
Their large, aquatic larvae rely predominantly on tegument
respiration, making this an ideal species to study the principle of
oxygen limitation. We examine the effects of hypoxia and
hyperoxia on CTmax and test whether variation in CTmax amongst
individual larvae is related to their oxygen consumption. Given
the central role of oxygen in hypotheses on limits to both thermal
tolerance and body size [6,13,15–17], we also discuss how the
results of our study may improve our understanding of the role of
oxygen in setting body size limits in arthropods. A second reason
to discuss insect gigantism, stems from the fact that many insects
exhibiting gigantism [18] apparently had aquatic larvae, although
establishing larval lifestyle from fossil data can be difficult [19,20].
We do not suggest that insect gigantism is restricted to insects with
early aquatic life stages. The now extinct order of Palaeodictyoptera contained several species that displayed marked gigantism
and available fossil evidence suggests a terrestrial larval stage
[21,22]. However, gigantic insects do seem to have been
predominantly aquatic; the aquatic larval stage is held to be a
common ancestral ground plan [20] for the giant griffenflies
(Protodonata, sometimes also referred to as dragonflies) and giant
mayflies (Ephemeroptera) of the late Palaeozoic, as well as the
stoneflies (Plecoptera) arising in the Permian. Additionally the
gigantic Carboniferous myriapods, the arthopleurids, are usually
considered amphibiotic, where the early life stages are aquatic
[20].
To date, attempts to understand insect gigantism in the late
Palaeozoic have mainly been approached from the perspective of
(fossilized) terrestrial adults. Yet, if oxygen limitation at thermal
extremes operates differently for aquatic larvae and terrestrial
adults, approaching the problem of historical gigantism from a
larval perspective instead may similarly shed new light on the
possible role of oxygen in setting insect body size limits. A larval
view seems furthermore promising as insects with aquatic larval
stages can circumvent the problem of structural support during
their growing phase [15], while their flying adults overcome leg
space limitations governing tracheal space limitations in walking
insects [23].
Here we demonstrate that upper thermal limits in the aquatic
life stages of an insect do indeed respond to external oxygen supply
and that these are related to the oxygen consumption of individual
larvae. These findings extend the generality of the hypothesis of
oxygen limitation of thermal tolerance, suggest that oxygen
constraints on body size may be stronger in aquatic environments,
and that oxygen toxicity may have actively selected for gigantism
in the aquatic stages of Carboniferous arthropods.
PLoS ONE | www.plosone.org
Results
Critical thermal maximum (CTmax) differed significantly across
oxygen treatments (Table 1) being raised by 1?53uC in hyperoxia
and lowered by 2?92uC in hypoxia (Fig. 1A). Individual larvae
differed in thermal sensitivity for oxygen consumption rates
(expressed by Q10 values), which significantly affected their CTmax
(Table 1). Thermal maxima were approximately 1uC lower for
individuals with a high thermal sensitivity (Q10 value of 3)
compared to individuals with a low thermal sensitivity (Q10 value
of 1) (Fig. 1B). While we did not find a direct relationship between
body mass and CTmax (Table 1; Fig. 1C), larger instars had higher
Q10 values in comparison to smaller instars (t-test, t1,41 = 2?19,
P = 0?038). This suggests that potential effects of body mass are
mediated primarily through increased thermal sensitivity, something which seems most apparent at hypoxia (Fig. 1C).
Discussion
Here we find strong evidence for the idea that a mismatch
between external oxygen supply and internal oxygen demand can
set thermal limits in aquatic insects [13]: hypoxia lowered thermal
maxima, whilst hyperoxia increased them (Fig. 1A). At the same
time, individuals that strongly increased oxygen consumption at
elevated temperatures had lower thermal maxima (Fig. 1B). These
results contrast with previous investigations on terrestrial adult
insects, and provide a proof of principle that oxygen limitation can
set upper thermal limits in aquatic insect larvae. Drawing on
differences in ontogeny (closed trachea) and ecology (aquatic
habitat), two explanations could be made for the apparent
mismatch between oxygen supply and demand in the stonefly
nymphs, causing the onset of oxygen limitation at thermal
extremes.
First, oxygen delivery may be impeded in stonefly nymphs
because of their closed tracheae and because of the lower oxygen
content and diffusion rates in water compared to air. At higher
temperatures, more oxygen is available to an aquatic organism
because of the higher diffusivity of oxygen, yet scope for aerobic
metabolism is nevertheless reduced as increases in organismal
oxygen demand exceed increases in oxygen supply [13]. Although
a closed tracheal system still represents a one-stage oxygen delivery
system, oxygen delivery is more likely to become rate limiting at
higher temperatures because of the additional step of oxygen
diffusion across the epithelium. The absence of air sacs in larvae
may further limit oxygen delivery rates by increasing the relative
importance of diffusive rather than convective movement of
oxygen in the trachea [9], although the compression and
expansion of the trachea themselves [24] may in some cases
Table 1. Statistical analysis of critical thermal maxima in
relation to ambient oxygen levels, larval oxygen consumption
and body mass.
Source
SS (Type III)
d.f.
F ratio
P-value
Oxygen treatment
131.897
2
45.338
,0.0001
Q10 oxygen consumption rates
12.132
1
8.341
0.0062
Body mass (mg dry weight)
.356
1
.245
0.6234
ANCOVA statistics on critical thermal maxima. Significant results are indicated in
bold. Thermal maxima were highest for hyperoxia (36 kPa) and lowest for
hypoxia (14 kPa). In addition thermal maxima were lowest for larvae which
consumed more oxygen at higher temperatures. (SS = Sum of squares;
d.f. = degrees of freedom).
doi:10.1371/journal.pone.0022610.t001
2
July 2011 | Volume 6 | Issue 7 | e22610
Oxygen, Thermal Limits and Insect Gigantism
Figure 1. Differences in critical thermal maxima (CTmax) in the stonefly Dinocras cephalotes at three different levels of oxygen (a), the
relationship between CTmax of the stonefly nymphs and their thermal sensitivity in oxygen consumption (b) and their body mass
(c). Differences in CTmax were consistent with the mechanism of oxygen limitation: hypoxia lowered CTmax, while hyperoxia increased CTmax (a) and
thermal maxima were lower for individuals which strongly increased their oxygen consumption rates at higher temperatures (high Q10 values). Each
bar represents the average (6 s.e.) of 15 nymphs. Letters indicate significant differences (P,0.05; Tukey HSD post hoc test following an ANOVA
including only oxygen treatment: F2,41 = 44?06, P,0?001).
doi:10.1371/journal.pone.0022610.g001
although the thermal limits associated with these performance
measures will be lower than those for the short-term survival
reported here [2,4,6].
In each of the above explanations the conditions that make
oxygen limitation more likely arise from both the aquatic nature
(lower oxygen availability and higher thermal buffering) and the
closed tracheal system (limiting oxygen delivery and regulatory
capacity). Consequently, oxygen limitation may be especially likely
for insects that have life stages with closed tracheal systems and live
in an aquatic or essentially aquatic environment (e.g. endoparasites, endophytic species, some rotten wood borers, rotten fruit
specialists, etc.). Thus, many insects may be affected by oxygen
limitation at some stage during their life cycle; indeed different life
stages vary in their susceptibility to hypoxia [10,34,35] and
thermal tolerance [36].
Similarly, both of the above explanations for the onset of oxygen
limitation at thermal extremes could underlie insect gigantism.
While they do not preclude additional evolutionary routes toward
gigantism (which seems most probable for entirely terrestrial taxa
like the Palaeodictyoptera), they could explain why gigantism was
apparently frequent amongst arthropods with juvenile aquatic life
stages. Importantly, each explanation makes very different
predictions. The first, of impeded oxygen delivery, follows the
existing explanation that increased atmospheric levels of oxygen in
the late Palaeozoic permitted the evolution of larger body sizes.
The basic difference is that oxygen limitation first sets in at the
larval stage, either owing to the lower availability of oxygen in
water compared to air [12,13] or the additional barrier of diffusion
across the epithelium.
Oxygen limitation in larvae fits with the observation that oxygen
delivery does not seem to become much more challenging for
larger bodied adults of terrestrial insects [10,15]. Although larger
animals are predicted to be more prone to oxygen limitation, such
size dependency may only be evident under certain conditions
[15,17], given that larger individuals may have modified
respiratory structures, and change their respiratory behaviour to
compensate for reductions in oxygen supply capacity associated
with larger size [10,15]. Costs associated with such compensatory
changes may include tracheal hypertrophy [9,23], or increased
thermal sensitivity (see results). These costs may be reflected in the
generate substantial convective movement [25]. An increased
difficulty of oxygen delivery in an aquatic environment fits with the
fact that CTmax at normoxia is generally reported to be higher for
terrestrial than aquatic arthropod life stages [7,8,26–29]. As CTmax
is reached at lower temperatures in aquatic taxa, this is less likely
to be a result of thermal damage at the cellular level such as the
disruption of membrane structure and problems associated with
protein folding [30,31], which would make oxygen limitation more
decisive in setting thermal limits in aquatic life stages, rather than
one of several factors as suggested for terrestrial insects [7,8]. One
assumption here is that aquatic taxa have not considerably altered
their membrane fluidity and protein activity in response to the
more stable, often lower thermal regime in aquatic habitats (see
below). If oxygen is more important at less extreme thermal
maxima relative to other factors (membrane fluidity and protein
stability), it would explain why CTmax increased with hyperoxia in
the stonefly nymphs studied here (Fig. 1A) and in aquatic mayfly
nymphs studied by Whitney [26], but not in the terrestrial
tracheates with open tracheal systems studied so far [7,8].
Second, changes in external oxygen levels may have stronger
impacts on internal oxygen levels in aquatic invertebrates, and
hence thermal tolerance, if their oxygen regulatory capacity is
more limited than that of terrestrial invertebrates. Insects are
forced to regulate internal oxygen levels within a fairly narrow
range, balancing the risk of asphyxiation with that of oxygen
toxicity [10,32,33]. Whereas terrestrial insects can simply open or
close spiracles to regulate oxygen uptake, such regulation is
unavailable to aquatic stages with closed tracheal systems such as
Dinocras. Equally, fluctuations in ectotherm oxygen consumption
are reduced in aquatic habitats owing to their thermally buffered
nature. In short, the ability to regulate internal oxygen levels is
inherently limited in taxa with closed tracheal systems, while the
need to do so is lower in aquatic habitats. With poor regulation of
oxygen intake in Dinocras, it is perhaps not too surprising to find a
relationship between CTmax of individual stonefly nymphs and
their thermal sensitivity in oxygen consumption (Fig. 1B). As these
measurements of oxygen consumption were performed at
ecologically realistic temperatures, aerobic scope for feeding,
growth and reproduction may be likewise affected by the interplay
between external oxygen supply and organismal oxygen demand,
PLoS ONE | www.plosone.org
3
July 2011 | Volume 6 | Issue 7 | e22610
Oxygen, Thermal Limits and Insect Gigantism
long term, underlying observed variation in body size across
environmental gradients of temperature or oxygen availability
[13,14]. Here we found support for size related performance in the
hypoxic treatment only (Fig. 1C). A possible explanation for this is
that the tracheal network of the larger individuals, which
developed under normoxia in the field, was of insufficient capacity
when larvae approached their thermal limits under hypoxia. At
higher levels of oxygen, other limits may have set in that were less
dependent on size, involving the failure of oxygen delivery systems
operating at the cellular level [14].
A fundamental problem with a passive oxygen ceiling
constraining maximum body size is that it is inferred from extant
taxa having physiologies optimized to normoxic conditions and
therefore may not reflect the evolutionary limit of tracheal
breathing [37]. The second explanation of limited oxygen
regulatory capacity recasts oxygen as an active driver of gigantism
by focussing on the risks of having too much oxygen, rather than
too little. In aquatic larvae with closed tracheal systems and limited
ability to regulate internal oxygen levels, internal oxygen levels
would be expected to closely track environmental oxygen levels.
Whilst species are likely to have experienced and evolved
responses to cope with periods of hypoxia [38], the same may
not apply for hyperoxia, putting aquatic larvae at greater risk of
oxygen poisoning than terrestrial adults [39]. If large body sizes
are more sensitive to hypoxia and asphyxiation, they may equally
confer protection from oxygen toxicity [40], constituting an
antioxidant response [41].
In support of oxygen as an active driver of increasing body size,
Loudon [9] found that beetle larvae increased in body mass when
they were transferred from hypoxia to normoxia during their
development. The logic is that larvae which started their
development in hypoxia increased their tracheal size, but could
not decrease them again upon returning to normoxia, as the new
trachea are built around the old trachea of the previous larval
instar. Although increasing body mass entails costs, body mass
results from the net effect of many different factors [14]. During
development, internal hypoxia acts as a signal and may stimulate
tissue differentiation instead of growth, thus affecting the size
reached. Similarly hyperoxia may trigger an increase in body size
as a readily available way to effectively escape oxygen toxicity,
possibly enabled through lower costs in ventilation or tracheal
investments [10,40,42]. Hence hyperoxia may actively drive
evolutionary increases in body mass, even in small insects [40].
Direct evidence for oxygen toxicity in a range of freshwater
invertebrate species is provided by Fox & Taylor [32] who found
that smaller juvenile stages are more sensitive to hyperoxic
conditions than their larger aquatic adults.
An active selection for larger body sizes under hyperoxia would
fit with the reappearance of giant mayflies in the putative high
oxygen atmosphere at the end-Cretaceous [43] and the persistence
of giant insects during putative lower levels of atmospheric oxygen.
Examples of the latter include large griffenflies (Protodonata) in
the late Permian [44] and abnormally large dragonflies (Odonata)
during the Triassic/Jurassic [45]. Similarly, oxygen as an active
driver of gigantism would predict a shift in size spectra such that
average size increases, rather than a unilateral broadening of size
spectra where only the body size of the largest species increases, as
predicted by a passive oxygen ceiling. While establishing changes
in modal sizes from the fossil record is fraught with difficulties, size
spectra in extant amphipod assemblages would support oxygen as
an active driver, where not only maximum body size within an
assemblage, but also average and even minimum body size is
greater at higher ratios of oxygen supply to demand [13,46].
PLoS ONE | www.plosone.org
Thus, a larval view of Paleozoic hyperoxia-enabled gigantism in
insects may be very informative. We suggest the aquatic larval
stage as a route to gigantism, explaining the predominance of
aquatic life stages amongst extinct gigantic insects. In aquatic
juvenile stages, impeded oxygen delivery may have resulted in
stronger constraints on body size, whilst larger body sizes may
have been actively selected for to avert oxygen toxicity. The
widespread gigantism in marine species further supports this
hypothesis, with examples of gigantism [47,48] found predominantly at times when oxygen levels were perhaps not higher than
current ones, but rising [49], effectively putting the animals at risk
of oxygen poisoning. Each of the two explanations for a larger role
of oxygen in aquatic stages yields testable predictions concerning
the relative rates of evolution of larger and smaller body sizes, and
the shifting or broadening size spectra.
Materials and Methods
Animals were collected from the River Dart, Devon, UK and
maintained in the lab at 1061uC in a 12 L:12 D regime. They
were kept in a flow-through aquarium (10 L?min21), fed with
artificial pond water [50], buffered and diluted to reflect the pH
and conductivity of the field site (pH 6.4–6.6, 70–150 mS?cm21)
and fed chironomid larvae. Oxygen consumption was measured
for each larva at 10uC and 15uC using closed glass respiration
chambers of 67.5–68.5 ml. The chambers were immersed in a
temperature controlled bath (60.1uC) and stirred using underwater magnetic stirrers to ensure mixing of water. Respiration
chambers were fitted with a fine nylon mesh forming a false
bottom to prevent contact between the larvae and the magnetic
stirrer bar. Individuals were allowed to acclimate for 10 min
before the chambers were closed and left for 60 min. Oxygen
content was measured before and after using an O2 electrode
(1302 Oxygen Electrode, Strathkelvin Instruments) that was
connected to a calibrated meter (Oxygen Meter 929, Strathkelvin
Instruments). On average, larvae depleted oxygen levels only by
5% (with a maximum of 18%) and oxygen consumption was
expressed as mg O2 (mg wet mass)21 h21. Oxygen consumption of
larvae at 10uC was significantly correlated with their consumption
at 15uC (Partial correlation, corrected for differences in body mass:
r = 0.511, P,0.001), justifying the calculation of Q10 values for
individuals. One individual was removed from further analysis as
abnormally low oxygen consumption measured at 10uC resulted in
an unrealistic Q10 value of 99.The Q10 values calculated for each
specimen were correlated with the CTmax values of the exact same
individuals.
To assess CTmax at normoxia, animals were placed in small
flow-through chambers (70670630 mm; flowrate 0.016 L?s21)
fed with water from a 25 L header tank passing through a tubular
counter current heat exchanger. Water in the header tank was of
the same composition as that used to maintain animals, and was
bubbled with a mixture of 20% oxygen and 80% nitrogen,
obtained using a gas-mixing pump (Wösthoff, Bochum, Germany),
and covered with 18 mm thick expanded polystyrene sheeting to
prevent equilibration with the atmosphere. Before the start of the
experiment, animals were left to acclimate for 1 h at the starting
temperature of 10uC, after which temperature in the experimental
chambers was increased at 0.25uC min21, using a Grant R5 water
bath with a GP200 pump unit (Grant Instrument (Cambridge)
Ltd, UK), connected to the heat exchanger. Temperatures were
logged using a HH806AU digital thermometer (Omega Engineering Inc., USA). CTmax was recorded as the point at which animals
no longer showed any body movement or muscular spasms.
Animals which were at this point transferred to fully oxygenated
4
July 2011 | Volume 6 | Issue 7 | e22610
Oxygen, Thermal Limits and Insect Gigantism
water of 10uC recovered with no apparent lasting damage. Below
critical maxima, larvae initiated in repeated swimming behaviour
(around 29uC; interpreted as attempts to escape experimental
conditions) and fell upon their backs (around 30uC) and an onset
of spasms was observed (around 31uC). We used a dynamic
method (acute exposure to changing temperatures) in contrast to
most work performed on marine taxa, which used static methods
(chronic exposure to constant temperatures [3,5 but see 2]). As
faster rates of warming can result in higher critical thermal
maxima [51,52], we employed the same rate of warming as
employed in previous studies on terrestrial insects to minimize
confounding effects arising from methodological differences in a
direct comparison of our results with theirs. To assess CTmax at
hypoxia (5% O2, 95% N2) and hyperoxia (60% O2, 40% N2), the
gas mixture was adjusted 10 min after placing the animals in the
small flow-through chambers. In this way animals were gradually
exposed to hypoxic and hyperoxic conditions during acclimation.
In several test runs, oxygen levels were measured after adjusting
the gas mixture to determine when and at which value oxygen
levels stabilized. From these measurements the relative contributions of the gas mixture and normal air to the (hypoxic or
hyperoxic) test water could be determined. The oxygen levels
stabilised after 1 hour at 36 kPa (34–38) and 14 kPa (13.3–14.8)
for the hyperoxia and hypoxia treatment, respectively.
Acknowledgments
We thank Andrew Atfield, Roger Haslam, Rick Preston, Peter Russell,
Richard Ticehurst and Saar Verberk for logistic support. Two anonymous
referees made critical, much appreciated comments that improved the
manuscript.
Author Contributions
Conceived and designed the experiments: WV DB. Performed the
experiments: WV. Analyzed the data: WV. Wrote the paper: WV DB.
References
1. Winterstein H (1905) Wärmelähmung und Narkose. Z Allg Physiol 5: 323–350.
2. Frederich M, Pörtner HO (2000) Oxygen limitation of thermal tolerance defined
by cardiac and ventilatory performance in spider crab, Maja squinado.
Am J Physiol – Regul Integr Comp Physiol 279: R1531–R1538.
3. Pörtner HO (2002) Climate variations and the physiological basis of temperature
dependent biogeography: systemic to molecular hierarchy of thermal tolerance
in animals. Comp Biochem Physiol A 132: 739–761. (doi:10.1016/S10956433(02)00045-4).
4. Pörtner HO (2006) Climate-dependent evolution of Antarctic ectotherms: An
integrative analysis. Deep-Sea Res II 53: 1071–1104. (doi:10.1016/
j.dsr2.2006.02.015).
5. Pörtner HO (2010) Oxygen- and capacity-limitation of thermal tolerance: a
matrix for integrating climate-related stressor effects in marine ecosystems. J Exp
Biol 213: 881–893. (doi:10.1242/jeb.037523).
6. Pörtner HO (2001) Climate change and temperature-dependent biogeography:
oxygen limitation of thermal tolerance in animals. Naturwissenschaften 88:
137–146. (doi:10.1007/s001140100216).
7. Stevens MM, Jackson S, Bester SA, Terblanche JS, Chown SL (2010) Oxygen
limitation and thermal tolerance in two terrestrial arthropod species. J Exp Biol
213: 2209–2218. (doi:10.1242/jeb.040170).
8. Klok CJ, Sinclair BJ, Chown SL (2004) Upper thermal tolerance and oxygen
limitation in terrestrial arthropods. J Exp Biol 207: 2361–2370. (doi:10.1242/
jeb.01023).
9. Loudon C (1989) Tracheal hypertrophy in mealworms: Design and plasticity in
oxygen supply systems. J Exp Biol 147: 217–235.
10. Harrison JF, Frazier MR, Henry JR, Kaiser A, Klok CJ, et al. (2006) Responses
of terrestrial insects to hypoxia or hyperoxia. Resp Physiol Neurobiol 154: 4–17.
(doi:10.1016/j.resp.2006.02.008).
11. Lighton JRB (2007) Hot hypoxic flies: Whole-organism interactions between
hypoxic and thermal stressors in Drosophila melanogaster. J Therm Biol 32:
134–143. (doi:10.1016/j.jtherbio.2007.01.009).
12. Jones JD (1972) Comparative Physiology of Respiration. Special Topics in
Biology Series. London: Edward Arnold Ltd.
13. Verberk WCEP, Bilton DT, Calosi P, Spicer JI (2011) Oxygen supply in aquatic
ectotherms: Partial pressure and solubility together explain biodiversity and size
patterns. Ecology 92: 1565–1572. (doi:10.1890/10-2369.1).
14. Atkinson D, Morley SA, Hughes RN (2006) From cells to colonies: at what levels
of body organization does the ‘temperature-size rule’ apply? Evol Dev 8:
202–214. (doi: 10.1111/j.1525-142X.2006.00090.x).
15. Harrison JF, Kaiser A, VandenBrooks JM (2010) Atmospheric oxygen level and
the evolution of insect body size. Proc R Soc Lond B 277: 1937–1946.
(doi:10.1098/rspb.2010.0001).
16. Chapelle G, Peck L (1999) Polar gigantism dictated by oxygen availability.
Nature 399: 114–115. (doi:10.1038/20099).
17. Woods HA, Moran AL, Arango CP, Mullen L, Shields C (2009) Oxygen
hypothesis of polar gigantism not supported by performance of Antarctic
pycnogonids in hypoxia. Proc R Soc Lond B 276: 1069–1075. (doi:doi:10.1098/
rspb.2008.1489).
18. Graham JB, Dudley R, Aguilar NM, Gans C (1995) Implications of the late
Palaeozoic oxygen pulse for physiology and evolution. Nature 375: 117–120.
(doi: 10.1038/375117a0).
19. Wootton RJ (1988) The historical ecology of aquatic insects: an overview.
Palaeogeogr Palaeoclim Palaeoecol 62: 477–492. (doi:10.1016/00310182(88)90068-5).
20. Kukalová-Peck J (1983) Origin of the insect wing and wing articulation from the
arthropodan leg. Can J Zool 61: 1618–1669. (doi:10.1139/z83-217).
PLoS ONE | www.plosone.org
21. Wootton RJ (1972) Nymphs of Palaeodictyoptera (Insecta) from the Westphalian
of England. Paleontology 15: 662–675.
22. Shear WA, Kukalová-Peck J (1990) The ecology of Paleozoic terrestrial
arthropods: the fossil evidence. Can J Zool 68: 1807–1834. (doi:10.1139/z90262).
23. Kaiser A, Klok CJ, Socha JJ, Lee W-K, Quinlan MC, et al. (2007) Increase in
tracheal investment with beetle size supports hypothesis of oxygen limitation on
insect gigantism. Proc Natl Acad Sci USA 104: 13198–13203. (doi:10.1073/
pnas.0611544104).
24. Westneat MW, Betz O, Blob RW, Fezzaa K, Cooper WJ, et al. (2003) Tracheal
respiration in insects visualized with synchrotron X-ray imaging. Science 299:
558–560. (doi:10.1126/science.1078008).
25. Socha JJ, Lee W-K, Harrison JF, Waters JS, Fezzaa K, et al. (2008) Correlated
patterns of tracheal compression and convective gas exchange in a carabid
beetle. J Exp Biol 211: 3409–3420. (doi:10.1242/jeb.019877).
26. Whitney RJ (1939) The Thermal Resistance of Mayfly Nymphs from Ponds and
Streams. J Exp Biol 16: 374–385.
27. Chown SL (2001) Physiological variation in insects: hierarchical levels and
implications. J Ins Physiol 47: 649–660. (doi:10.1016/S0022-1910(00)00163-3).
28. Calosi P, Bilton DT, Spicer JI, Atfield A (2008) Thermal tolerance and
geographical range size in the Agabus brunneus group of European diving beetles
(Coleoptera: Dytiscidae). J Biogeogr 35: 295–305. (doi:10.1111/j.13652699.2007.01787.x).
29. Wijnhoven S, van Riel MC, van der Velde G (2003) Exotic and indigenous
freshwater gammarid species: physiological tolerance to water temperature in
relation to ionic content of the water. Aquat Ecol 37: 151–158. (doi:10.1023/
A:1023982200529).
30. Feder ME, Hofmann GE (1999) Heat-shock proteins, molecular chaperones,
and the stress response:Evolutionary and Ecological Physiology. Annu Rev
Physiol 61: 243–82. (doi:10.1146/annurev.physiol.61.1.243).
31. Somero GN (2004) Adaptation of enzymes to temperature: searching for basic
‘‘strategies’’. Comp Biochem Physiol B 139: 321–333. (doi:10.1016/
j.cbpc.2004.05.003).
32. Fox HM, Taylor AER (1955) The tolerance of oxygen by aquatic invertebrates.
Proc R Soc Lond B 143: 214–225. (doi:10.1098/rspb.1955.0006).
33. Hetz SK, Bradley TJ (2005) Insects breathe discontinuously to avoid oxygen
toxicity. Nature 433: 516–519. (doi:10.1038/nature03106).
34. Woods HA, Hill RI (2004) Temperature-dependent oxygen limitation in insect
eggs. J Exp Biol 207: 2267–2276. (doi:10.1242/jeb.00991).
35. Klok CJ, Kaiser A, Lighton JRB, Harrison JF (2010) Critical oxygen partial
pressures and maximal tracheal conductances for Drosophila melanogaster reared
for multiple generations in hypoxia or hyperoxia. J Ins Physiol 56: 461–469.
(doi:10.1016/j.jinsphys.2009.08.004).
36. Marais E, Terblanche JS, Chown SL (2009) Life stage-related differences in
hardening and acclimation of thermal tolerance traits in the kelp fly, Paractora
dreuxi (Diptera, Helcomyzidae). J Ins Physiol 55: 336–343. (doi:10.1016/
j.jinsphys.2008.11.016).
37. Butterfield NJ (2009) Oxygen, animals and oceanic ventilation: an alternative
view. Geobiol 7: 1–7. (doi:10.1111/j.1472-4669.2009.00188.x).
38. Hoback WW, Stanley DW (2001) Insects in hypoxia. J Ins Physiol 47: 533–542.
(doi: 10.1016/S0022-1910(00)00153-0).
39. Van Voorhies WA (2009) Metabolic function in Drosophila melanogaster in response
to hypoxia and pure oxygen. J Exp Biol 211: 3409–3420. (doi:10.1242/
jeb.031179).
40. Klok CJ, Hubb AJ, Harrison JF (2009) Single and multigenerational responses of
body mass to atmospheric oxygen concentrations in Drosophila melanogaster:
5
July 2011 | Volume 6 | Issue 7 | e22610
Oxygen, Thermal Limits and Insect Gigantism
41.
42.
43.
44.
45.
46.
evidence for roles of plasticity and evolution. J Evol Biol 22: 2496–2504.
(doi:10.1111/j.1420-9101.2009.01866.x).
Lane N (2002) Oxygen: The Molecule that made the World. Oxford: Oxford
University Press.
Owerkowicz T, Elsey RM, Hicks JW (2009) Atmospheric oxygen level affects
growth trajectory, cardiopulmonary allometry and metabolic rate in the
American alligator (Alligator mississippiensis). J Exp Biol 212: 1237–1247.
(doi:10.1242/jeb.023945).
Grimaldi D, Engel MS (2005) Evolution of the Insects. New York: Cambridge
University Press.
Nel AN, Fleck G, Garrouste R, Gand G (2008) The Odonatoptera of the Late
Permian Lodève Basin (Insecta). J Iberian Geol 34: 115–122.
Okajima R (2008) The controlling factors limiting maximum body size of insects.
Lethaia 41: 423–430. (doi:10.1111/j.1502-3931.2008.00094.x).
Chapelle G, Peck LS (2004) Amphipod crustacean size spectra: new insights in
the relationship between size and oxygen. Oikos 106: 167–175. (doi:10.1111/
j.0030-1299.2004.12934.x).
PLoS ONE | www.plosone.org
47. Rudkin DM, Young GA, Elias RJ, Dobrzanski EP (2003) The world’s biggest
trilobite – Isotelus rex new species from the Upper Ordovician of northern
Manitoba, Canada. J Paleont 77: 99–112.
48. Braddy SJ, Poschmann M, Tetlie OE (2008) Giant claw reveals the largest ever
arthropod. Biol Lett 4: 106–109. (doi: 10.1098/rsbl.2007.0491).
49. Berner RA, VandenBrooks JM, Ward PD (2007) Oxygen and Evolution.
Science 316: 557–558.
50. ASTM (1980) Standard practice for conducting acute toxicity tests with fishes,
macroinvertebrates and amphibians. In: American standard for testing and
materials, Philadelphia, Pennsylvania, USA. pp 279–280.
51. Terblanche JS, Deere JA, Clusella-Trullas S, Janion C, Chown SL (2007)
Critical thermal limits depend on methodological context. Proc R Soc Lond B
274: 2935–2942. (doi:10.1098/rspb.2007.0985).
52. Peck LS, Clark MS, Morley SA, Massey A, Rossetti H (2009) Animal
temperature limits and ecological relevance: effects of size, activity and rates
of change. Funct Ecol 23: 248–256. (doi: 10.1111/j.1365-2435.2008.01537.x).
6
July 2011 | Volume 6 | Issue 7 | e22610