Chronic changes in developmental oxygen have little effect on mitochondria and tracheal density in the endothermic moth Manduca sexta Free

The level of ambient oxygen during development influences insect respiratory and mitochondrial physiology (Harrison et al., 2006; Harrison and Haddad, 2011). In Drosophila, for example, hypoxic rearing conditions are associated with increased adult tracheolar density (Harrison et al., 2018) and adult mitochondrial density (Perkins et al., 2012; VandenBrooks et al., 2018). Hyperoxic rearing sometimes has the opposite effect on tracheolar and mitochondrial densities (VandenBrooks et al., 2018). This plasticity of cellular respiratory morphology is thought to be caused by compensatory mechanisms to maintain a narrow range of oxygen partial pressures within the tracheal system to support aerobic metabolic processes. This compensation is accomplished through signaling that increases or decreases terminal branching in the tracheal system during development (Jarecki et al., 1999). Changes in the tracheal system are thought to increase oxygen transport in low oxygen environments or limit detrimental reactive oxygen species (ROS) at high oxygen concentrations. This is especially important in endothermic flying insects that rely on aerobic respiration to fund their energetically expensive mode of locomotion. Many of the studies on chronic oxygen exposure have focused on ectothermic flying insects, mainly Drosophila (Harrison et al., 2018; VandenBrooks et al., 2018). There is a paucity of studies examining developmental plasticity in response to altered oxygen in large, endothermic insects where energy demands are significantly higher. This increase is due to their relatively large size and is a consequence of increase wing beat frequencies and loading (Heinrich, 1974). Most studies of endothermic insects focus on responses to acute oxygen changes (Dzialowski et al., 2014; Henry and Harrison, 2004; Joos et al., 1997) rather than chronic exposure or only examine the ectothermic larval stages (Callier and Nijhout, 2011). Facultative endothermy evolved in some larger insects, such as bees (Heinrich and Esch, 1994), moths (Heinrich, 1971) and dragonflies, to allow increased muscle temperatures that support the wingbeat frequencies of larger, agile flyers across a wider range of environmental temperatures (Heinrich, 1987; Heinrich and Vogt, 1993). These insects increase flight muscle temperature and mass-specific power output in cooler temperatures for various ecological reasons (Glass and Harrison, 2022). Flight is an extremely costly activity that is funded by aerobic metabolism (Komai, 1998; Socha et al., 2010). High ATP consumption and production lead to a high demand for oxygen and may be impacted by chronic hypoxia or hyperoxia in different ways from those previously observed in fully ectothermic species such as Drosophila.

Aerobic mitochondrial ATP production is required for the endothermic response in insects as flight and thermoregulation are both accomplished almost exclusively through aerobic respiration of the flight muscles. Mitochondrial ATP production is an excellent measure of metabolic physiology. It can be used to look at phenotypic changes of both coupled and uncoupled respiration, in endothermic flying insects, on a cellular level (Hood et al., 2018). Measurement of mitochondrial respiration in muscle has focused on vertebrates or on genetic model organisms. Recently, there has been an increased focus on endothermic insects, bees and wasps, looking at mitochondrial function in permeabilized flight muscle fibers (Hedges et al., 2019; Teulier et al., 2016) and isolated mitochondria (Syromyatnikov et al., 2019, 2013). Insect flight muscle has evolved the use of different substrates for ATP production. For example, some bees can utilize pathways with proline and glycerol 3-phosphate for ATP production (Hedges et al., 2019; Masson et al., 2017; Teulier et al., 2016) supporting mitochondrial oxidative phosphorylation (OXPHOS) of flight muscles. We have shown that Manduca sexta adults do not utilize either the glycerol 3-phosphate or proline pathway as mitochondrial substrates (Wilmsen and Dzialowski, 2023).

As oxidative phosphorylation is oxygen dependent, changes in oxygen availability can impact ATP synthesis. The long-term effects of developmental oxygen exposure during larval and pupal stages on metabolic activity have been studied in Drosophila. Drosophila reared under developmental hypoxia exhibit decreased isolated mitochondrial oxygen consumption and decreased reactive oxygen species production (Ali et al., 2012). Perkins et al. (2012) found increased flight muscle mitochondrial densities in hypoxia-reared Drosophila, implying the potential for increased metabolic capacity. These two studies are not inherently contradictory as the quantity of mitochondria does not indicate the per unit efficiency. How endothermic insects' mitochondria respond to developmental hypoxia and hyperoxia is largely unknown.

This study examined the effect of chronic hypoxia and hyperoxia exposure during larval and pupal development on adult M. sexta flight muscle mitochondrial respiration and underlying muscle morphometrics, such as tracheal morphology and mitochondrial density. Manduca sexta are a physiological model organism because they are highly endothermic with the adult moth thorax temperatures reaching up to 45°C in flight (Heinrich, 1971). The flight muscles, relying on aerobic mitochondrial metabolism, need to receive sufficient oxygen to meet the aerobic respiration demand through the internal respiratory tracheal system. While ectothermic insect species show plasticity in the development of their trachea in response to developmental hypoxia or hyperoxia, it is unclear whether larger endothermic insects exhibit the same developmental plasticity or are constrained by the requirements afforded by the high oxygen demand of flight muscles. We hypothesized that rearing M. sexta larvae and pupae in hypoxia would lead to greater tracheal endowment and mitochondrial respiration rates of flight muscles in adults while hyperoxia-reared adults would have lower tracheal endowment and mitochondrial respiration rates.

Manduca sexta (Linnaeus 1763) eggs were purchased from Carolina Biological Supply. Upon arrival, eggs were carefully separated and placed in containers with mixed hornworm media (Carolina Biological Supply) to hatch into larvae (see below). Once larvae were big enough to handle without harming them (about 11 days after arrival), they were weighed, an ID was assigned to each larva, and they were moved to individually labeled vials with the same hornworm media. Larvae were allowed to grow, and tubes were cleaned every other day to reduce mold growth. When larvae in their 5th instar had reached the wandering phase immediately prior to pupation, they were weighed again and placed in a dark vial containing wood shavings for a week to pupate. After 5 days in the dark, pupating larvae were checked daily to see whether they had completed pupation. Once they had pupated, they were removed from the dark vial to prevent them from entering diapause, weighed and placed into large plastic bins (57.2×40.6×32.4 cm). The pupae were checked daily to determine when they eclosed. Adult moths were marked with Testors™ acrylic paint, to keep track of the date they hatched, and maintained in the plastic bins. Adults were provided access to sugar water as described in Wilmsen and Dzialowski (2024) ad libitum until they were used for experiments. The moths were weighed a final time 3–4 days after they had eclosed. Moths were maintained on an 18 h:6 h light:dark cycle, at 25°C, and a humidity of ∼38%.

To quantify developmental effects of chronic hypoxia or hyperoxia exposure on the metabolic phenotype of adult M. sexta, once eggs were received, they were immediately placed in one of three ­O₂ levels: 10%, 21% or 30% O₂. The animals were maintained chronically at these O₂ levels throughout their entire development. To maintain constant O_2­, the vials containing the developing animals were placed in tightly closed plastic bins and oxygen was regulated with a ROXY-4 gas regulator (Sable Systems International) and oxygen sensors (Maxtec, LLC). To maintain hypoxia, the ROXY-4 controlled the influx of excess nitrogen gas into the chamber and for hyperoxia, it controlled the influx of excess oxygen into the chamber. Small fans inside the bins pulled in some outside air to ensure humidity levels did not drop too low and circulated the air to ensure that the O₂ level was consistent throughout the container.

Mitochondrial respiration was measured in permeabilized flight muscle from adult M. sexta using an O2k Oxygraph (Oroboros Instruments). Thorax flight muscles were isolated, prepared and respirometry was run using methods adapted from Hedges et al. (2019) and Teulier et al. (2016). Moths were chilled at 4°C for 7 min, the thorax was removed, and the flight muscles were dissected out and placed in ice-cold BIOPS (in mmol l⁻¹: 2.77 CaK₂EGTA, 7.23 K₂EGTA, 5.77 Na₂ATP, 6.56 MgCl₂·6H₂O, 20 taurine, 15 Na₂-phosphocreatine, 20 imidazole, 0.5 dithiothreitol and 50 MES; pH 7). Muscle fibers were gently teased apart and then rocked at 4°C for 5 min in MIR05 (in mmol l⁻¹: 0.5 EGTA, 3 MgCl₂, 60 lactobionic acid, 20 taurine, 10 KH₂PO₄, 20 Hepes and 110 sucrose, with 1 g l⁻¹ BSA; pH 7.1). Between 0.5 and 2 mg of tissue were placed into the respiratory chambers of the O2k Oxygraph. We assessed differences in mitochondrial oxygen consumption rate between the three developmental oxygen treatments at two different temperatures, 25 and 40°C. We examined these two temperatures because they include the ambient temperature that a resting flight muscle would experience in our experiments and the elevated flight muscle temperature during flight. Initially, the substrates pyruvate (5 mmol) and malate (2 mmol) were added to assess LEAK respiration through complex I (Fig. S1). This was followed by the addition of saturating ADP (5 mmol) to stimulate OXPHOS respiration through complex I. To test substrate use by complex II, succinate (10 mmol) was added. Cytochrome c was then added to test mitochondrial membrane integrity. N,N,N′,N′-Tetra-methyl-p-phenylenediamine (TMPD, 0.5 mmol) and ascorbate (2 mmol) were added last to stimulate oxygen flux of complex IV, cytochrome oxidase (COX). Using two O2k Oxygraphs, permeabilized flight muscle mitochondria respiration was measured at both 25 and 40°C for each moth simultaneously.

Moth flight muscle mitochondria were isolated from individual moths using methods adapted from Syromyatnikov et al. (2013). Moths were sedated in the freezer for 7–10 min and then the flight muscle was removed. All isolation steps were performed on ice. A single moth's flight muscles were cut into several smaller pieces using fine scissors in isolation buffer (in mmol l⁻¹: 100 sucrose, 220 mannitol, 1 EGTA and 20 Hepes, with 0.2% BSA, pH 7.1) and then transferred to 10 ml of isolation buffer containing 1 mg ml⁻¹ protease (Nagarase, Sigma). The muscle was digested for 10 min at 4°C with rocking, then rinsed 3 times in isolation buffer and placed in ∼10 ml isolation buffer. Flight muscle was then homogenized using an overhead homogenizer (Scilogex SCI20-S) at 60 rpm for 4–5 passes. The solution was strained through two layers of cheesecloth and placed in a centrifuge tube for centrifugation at 600g for 10 min and the resulting pellet discarded. Then, the supernatant was centrifuged at 8500g for 10 min, the supernatant was discarded, and the pellet resuspended in 1 ml of isolation buffer. The solution was centrifuged a final time at 8500g for 10 min and suspended in 250 µl respiration media (in mmol l⁻¹: 100 sucrose, 220 mannitol, 1 EGTA and 20 Hepes, pH 7.1). The suspension was allowed to sit for at least 15 min before being added to the O2k Oxygraph respirometer. Protein concentration of isolated mitochondria was obtained using a standard Bradford Assay (Pierce™ Coomassie Protein Assay). Isolated mitochondrial respiration was measured using an O2k Oxygraph. The concentration of protein was 30 µg ml⁻¹ for each chamber. Mitochondrial respiration rates were determined using the substrate protocol described above for the permeabilized fibers (Fig. S1) and all measurements were performed only at 40°C.

Flight muscle tissue was prepared following a modified protocol developed by Henry Lujan (Baylor Medical Center). Moths were chilled at 4°C for 7 min until they were inactive. Flight muscles were dissected out and fixed overnight in 2.5% glutaraldehyde solution (100 µl 25% EM glutaraldehyde in 900 µl cacodylate buffer). The tissue was washed 3 times in cacodylate buffer for 10 min each. Muscle tissue was then incubated with osmium stain [3 µl 1 mmol l⁻¹ calcium chloride buffer, 400 µl 2% potassium ferrocyanide, 250 µl 4% osmium tetroxide (OsO₄), 247 µl 0.2 mol l⁻¹ cacodylate buffer] for 2.5 h. After incubation, the tissue was washed in cacodylate buffer twice for 10 min and placed in 1% uranyl acetate for 2.5 h. The tissue was again washed twice in buffer for 10 min each before undergoing acetone dehydration. To dehydrate the tissue, it was placed in 50% and then 70% acetone twice for 10 min each and 90% and 100% acetone twice for 15 min each. Tissue was then infiltrated with epoxy resin by subsequent incubation of 3:1 acetone:epoxy for 1 h, 1:1 acetone:epoxy overnight, 1:3 acetone:epoxy for 3 h and finally 100% epoxy for 4 h. Tissue was placed in fresh epoxy in the correct orientation, either for longitudinal sectioning or for cross-sectioning, and the epoxy was cured in a 60°C oven for 3 days. After the epoxy blocks were cured, the tissue samples were cut into ultra-thin sections ∼50 nm thick using an Ultra Microtome (Leica EM UC7) and sections were placed on 3 mm formvar-coated copper grids for viewing on a transmission electron microscope (TEM). The grids were then stained for 5 min with 2% lead citrate followed by 15 min with 2% uranyl acetate.

The stained grids were imaged using a JEOL 1400+ transmission electron microscope at the UT Dallas Molecular and Protein Analysis Core (MoPAC). Images were taken at ×5000 magnification and analyzed using ImageJ. Image analysis methods were adapted from Lam et al. (2021). Briefly, two images from each individual in both muscle orientations were analyzed for mitochondria and myofibril morphology. The images were first split into quadrates using the quadrat picker plugin on ImageJ and two randomly selected quadrates were analyzed. Both mitochondria and myofibrils were assessed for their total cross-sectional area of the muscle. Each structure was outlined using the freeform tool, added to the RIO manager, and measured. To obtain mitochondria density, the area of each structure type was summed within a quadrant and then divided by the total area of the quadrant to get a measure of the structure area (µm²) as a function of total area examined (µm²).

Methods for prepping and imaging flight muscle trachea were modified from VandenBrooks et al. (2018). Adult moths were placed in the freezer for 10 min to chill them and then the head and abdomen were removed from the thorax. A cut was made along the ventral median line of the thorax to allow fixative to adequately penetrate the tissue. The thorax was then fixed in 4% paraformaldehyde (PFA) overnight (Electron Microscopy Sciences). Once the thorax tissues were fixed, they were taken out of PFA and cut. First, the thorax was cut in half along the ventral midline and then a section ∼1 mm thick was cut on either side of the midline. This resulted in two sections for each thorax. After sections were cut, they were placed in phosphate-buffered saline (PBS) wash for 10 min; this step was repeated 3 times. The sections were then moved to a 3:1 PBS:glycerol solution for 10 min followed by a 1:1 PBS:glycerol solution for an additional 10 min. The sections were mounted on 1.75 mm deep concave slides (Carolina Biological Supply) with 100% glycerol solution.

The slides were imaged using a Zeiss LSM710 confocal microscope. A laser wavelength of 405 nm was used to excite the tracheae with an emission band of 422–528 nm. Because insect trachea is primarily composed of chitin, they auto-fluoresce under this wavelength of light. Magnification was set to ×40 water immersion lens and an image Z-stack with depth of approximately 0.40 µm with a 1 µm distance between sections was taken. Pixel size was 0.104 µm pixel⁻¹. The smallest tracheole tips are 20–40 nm (VandenBrooks et al., 2018), so our resolution may not have been fine enough to resolve the smallest tracheoles. Images were then processed and analyzed using ImageJ software.

ImageJ software was used to determine both tracheal density and tracheole diameter. Analysis of tracheal morphology was made on 32 slices of a 2D Z-stack for each section; the resulting image was then background subtracted, and the threshold adjusted so only the trachea were visible. After thresholding, the Vessel Analysis Plugin (Teplyi and Grebchenko, 2019) was used to measure tracheal density of the smaller trachea in the compressed Z-stack of 32 slices as well as tracheal diameter. As another measure of tracheal density, images were analyzed by adding two lines, 50 µm long, perpendicular to the muscle fiber and the resulting plot profile was used to determine the number of tracheae that intersected the line.

Oxygen flux for permeabilized fibers was analyzed using independent mixed effect linear models for each substrate, with rearing oxygen level and temperature as independent variables (Table S1). A two-way ANOVA was used to analyze the isolated mitochondria respiration, with developmental oxygen treatment and substrate as the independent factors. Substrate contribution ratio and coupling efficiency for permeabilized fibers were analyzed by mixed effect linear models, with temperature and rearing oxygen levels as factors. Confocal images were analyzed using one-way ANOVA to look at the effect of developmental oxygen concentration on morphology (Table S2). Analysis of data from TEM images was conducted with a linear mixed model with oxygen treatment as the fixed factor and animal number for the replicate measurement as a random factor. Statistics were run using GraphPad Prism version 9 and R running on JAMOVI version 2. Data are reported as means±s.e.m.

There was a significant effect of chronic rearing oxygen exposure during the larval and pupal stages on permeabilized flight muscle fiber mitochondrial oxygen flux rate (Table S1; Fig. 1). There was no difference in the LEAK oxygen flux between moths raise at different oxygen levels. Hypoxia-raised animals had increased oxygen fluxes when compared with control and hyperoxia-reared animals during OXPHOS using complex I stimulated with ADP (Table S1; Fig. 1) as well as OXPHOS using complex I and II stimulated with succinate (Table S1; Fig. 1). COX activity was significantly different between the oxygen groups (Table S1), but this did not appear after correction by the post hoc tests. The coupling efficiency and substrate contributions were not different between the three different rearing conditions (Table S1; Fig. 1B,C). There was an increase in oxygen flux of between 30% and 40% with the addition of succinate (Fig. 1C).

The effect of developmental oxygen concentration was also examined on isolated flight muscle mitochondria at 40°C. There were no significant differences in mitochondrial respiration rates within any substrates when comparing the three different developmental oxygen conditions when isolated mitochondria were used to measure respiration (P=0.7257; Fig. 2; Table S1). Coupling efficiency was lower in the controls compared with the hypoxia- and hyperoxia-reared adults (Fig. 2B). There were no differences in the substrate contributions between the rearing oxygen conditions (Fig. 2C).

Using both flight muscle TEM and confocal images, we analyzed the morphometrics of mitochondria, myofibrils and trachea in response to rearing oxygen level (10%, 21% or 30% O₂). Two different section perspectives, cross-section and longitudinal, were used to assess the mitochondria and sarcomeres (Fig. 3). The cross-sectional preparation provides a view of the mitochondria surrounding each myofibril and is a better way to analyze both mitochondria and myofibril density in the muscle as they are more discrete shapes and less sensitive to orientation. The longitudinal preparation provides a view of the myofibrils along the repeating sarcomeres, allowing for measurement of sarcomere length as well as mitochondrial density. For the cross-sectional orientation, there were no significant differences between oxygen development treatments in mitochondrial density (Fig. 4A; Table S2), total myofibril density (Fig. 4B; Table S2) or mean individual myofibril size (Fig. 4C; Table S2). When looking at the longitudinal section, there were no differences in mitochondrial density (Fig. 4D; Table S2) or sarcomere length (Fig. 4E; Table S2).

Flight muscle trachea can be visualized using a confocal microscope because tracheal chitin auto-fluoresces (Fig. 5). Typically, larger tracheae are found near the outer edge of each muscle fiber and start to branch in towards the center. As these tracheae proceed toward the center, they branch into tracheoles that run along each myofibril, parallel to the sarcomeres. From the confocal images, there were no differences in the tracheal area in the compressed Z-stack (Fig. 6A; Table S2), tracheal size (Fig. 6B; Table S2) or mean number of tracheal intersections (Fig. 6C; Table S2).

Mitochondrial function changes have been shown to be extremely important in an organism's response to environmental stressors. These changes can be seen both on the evolutionary scale and at the phenotypic level. In deer mice adapted to high elevation, there is an increase in mitochondrial quantity as well as increased fiber respiration in the gastrocnemius muscle when compared with lower altitude mice (Scott et al., 2018). Inter-tidal invertebrates have also evolved various strategies to cope with acute hypoxia or even anoxic conditions that they are exposed to regularly (Sokolova, 2018). There are some inconsistencies in the literature when it comes to phenotypic mitochondrial responses to hypoxia in insects. For example, Drosophila reared in hypoxia showed a decrease in ROS production and oxygen consumption in isolated mitochondria compared with normoxic animals (Ali et al., 2012). When flour beetles (Tribolium castaneum) (Wang et al., 2018) and Drosophila (Bosco et al., 2015) were exposed to 3 h of hypoxia, they had reduced citrate synthase activity compared with controls (Wang et al., 2018), indicating reduced metabolic activity. These short-term exposures are contradictory to findings for Drosophila raised in hypoxia for multiple generations that have increases in mitochondrial densities (Perkins et al., 2012), indicating an increased whole-tissue oxygen respiration capacity. When a subspecies of locust (Locusta migratoria) that live at high altitude was compared with a subspecies inhabiting lowlands, the high-altitude subspecies had increased complex I activity in hypoxia compared with those living near sea level (Zhao et al., 2013). In this study, developmental hypoxia had a significant effect on permeabilized flight muscle fiber mitochondrial oxygen flux when utilizing both complex I and II substrates, inducing higher oxygen respiration than both normoxic and hyperoxic animals (Fig. 1). These results indicate that development in hypoxia increased OXPHOS capacity at the tissue level but did not affect LEAK or oxygen flux through complex V (COX).

This effect of oxygen treatment during development was not reflected when the mitochondria were isolated and measured (Fig. 2). It may be that there were changes in physiology in the more intact muscle fibers that influence mitochondrial oxygen flux. When these factors are removed during mitochondria isolation, this difference may be rendered negligible. In humans, change in mitochondrial capacity associated with high altitude is attributed in part to the difference in metabolic substrates utilized. Tibetan Sherpa who live at high altitudes have been shown to rely on glucose as a fuel rather than lipid oxidation (Murray et al., 2018). This could be a contributing factor in this study. Moth flight muscle has been shown to readily utilize lipid oxidation (Hansford and Johnson, 1976) to make use of fats reserves present in these adult moths (Downer and Matthews, 1976). We did not investigate the lipid oxidation of the muscle mitochondria here but it is possible that the difference seen in the permeabilized muscle fibers is at least in part due to a change in substrate usage in the hypoxic animals towards glucose and away from lipid metabolism. The inconsistencies in the literature and our findings indicate that there is still much we do not understand about how this system operates.

When looking at tracheal density, we did not see any significant changes in response to developmental oxygen. The resolution on the confocal images may have been insufficient to capture changes because the smallest trachea we were able to resolve was ∼0.4 μm in diameter. In addition, our sample size was small and may not have had the power to resolve changes. However, there was no evidence of a trend in any of our morphological data. In studies with Drosophila, changes in tracheal structure in response to developmental hypoxia typically are seen only in the smallest tracheoles (diameter ≤0.2 μm) (Harrison et al., 2018; VandenBrooks et al., 2018). But with cockroaches (Blatella germanica) exposed to developmental hypoxia or hyperoxia, there are changes, with hypoxia inducing larger tracheal volumes and hyperoxia producing smaller tracheal volumes. These changes were seen even in the larger main leg trachea when correcting for mass (VandenBrooks et al., 2012). With all of this in mind, while we can say that we did not see any changes on a larger scale, we cannot conclude whether there were changes to the trachea at the smallest level in our moths.

When it comes to mitochondrial and myofibril densities, there was no effect of developmental treatment on muscle morphology, even though we saw differences in mitochondrial oxidative capacity. Differences in mitochondrial density did approach significance in both orientations, but there was no clear, consistent pattern between them. It should be noted that the sample size was low with the linear mixed model being slightly underpowered, with power of between 0.6 and 0.67. Our results suggests that the Manduca system has limited phenotypic plasticity in response to altered developmental oxygen. This is inconsistent with the responses observed in dipterans. Drosophila raised in hypoxia over multiple generations have increased mitochondrial densities and decreased muscle fiber bundles (Perkins et al., 2012). The use of synchronous muscle necessitates more investment in the sarcoplasmic reticulum to generate enough Ca²⁺ and more ATP use for equivalent muscle contraction because about 50% of ATP generated is used to power Ca²⁺ pumps (Rome and Lindstedt, 1998). This may lead to an inflexible system that is optimized for highly aerobic flight at our current ambient oxygen concentration, at least when it comes to mitochondrial density and trachea. Or potentially the phenotypic changes do not occur during a single generation and a multigenerational study may reveal significant changes.

In summary, we found that, surprisingly, these moths had very little developmental plasticity when we measured mitochondrial respiration, tracheal morphology and muscle mitochondrial morphology. This is one of only a few studies looking at mitochondrial respiration in the moth M. sexta. The oxygen consumption of both permeabilized fibers and isolated mitochondria corresponded well to values that are known for other flying insects such as bees, wasps and Drosophila. There was a different response to developmental hypoxia at the permeabilized fiber level compared with isolated mitochondria, indicating some change that was only associated with intact tissue. However, there was also no change in tracheal, mitochondrial or myofiber morphology or density that could have accounted for the mitochondrial respiration differences. These differences coupled with the inconsistencies in the literature are puzzling and warrant more research to tease apart the causes and effects of chronic oxygen treatment in insects.

The Results and Discussion in this paper are reproduced from the PhD thesis of S.M.W. (Wilmsen, 2022). Block cutting and TEM imaging were performed using the facilities at UT Dallas Molecular and Protein Analysis Core (MoPAC).