Protective role of trehalose in the Namib desert ant, Ocymyrmex robustior Free

Ants have colonized most habitats in terrestrial ecosystems, except for those in polar regions. They play key ecological roles as predators, scavengers and herbivores, and participate in mutualisms with many diverse organisms (e.g. insects, plants, fungi) (Wilson, 1971; Hölldobler and Wilson, 1990). Their remarkable ecological success is largely attributed to their societal structure, which is characterized by a division of labor between reproductive queens and non-reproductive workers, a regime that ensures colony maintenance (Wilson, 1971).

Some ant species are found in the hottest and driest deserts on Earth. In these environments, most organisms avoid scorching midday temperatures by being nocturnal or exploiting thermal shelters (e.g. escaping into burrows or under rocks). In contrast, these ants forage during the hottest periods of the day, when air and ground temperatures may reach up to 50°C and 70°C, respectively (Marsh, 1985; Andersen, 2007; Wehner and Wehner, 2011; Boulay et al., 2017). The resulting reduction in competition and predation allows workers to exploit food resources more effectively (Cerdá et al., 1998; Wehner, 2020). Desert ants have evolved several behavioral, morphological and physiological adaptations to cope with the extreme conditions of their habitat (reviewed in Perez and Aron, 2020). They utilize thermal refuges (e.g. shadows or elevated spots), where the air temperature is cooler, to avoid overheating (Wehner, 1987). Their long legs maximize the distance between their body and the burning ground, allowing higher running speeds, reduced foraging times and enhanced forced convective cooling (Sommer and Wehner, 2012; Centorame et al., 2019). At the molecular level, the expression of co-regulated gene clusters promotes proteome stability, the elimination of toxic residues and the maintenance of cell integrity, which likely help workers tolerate acute heat stress close to their lethal threshold (Gehring and Wehner, 1995; Willot et al., 2018; Perez et al., 2021).

Surprisingly, research has yet to examine the role of low-molecular-weight osmolytes in the thermal tolerance displayed by desert ants. Such metabolites are known to improve tolerance to abiotic stress in various taxa (Bandyopadhyay et al., 2019; Jogawat, 2019; Noer et al., 2020). For example, sugar osmolytes enhance thermal and desiccation resistance to cold stress (yeasts: Schade et al., 2004; plants: Nägele and Heyer, 2013; flies: Colinet et al., 2013) and heat stress (bacteria: Potts, 1994; plants: Rosa et al., 2009; bumble bees: Prieto, 2016). Of particular note is trehalose, which has an unmatched hydration ability because it very effectively reduces the mobility of hydrogen-bonded water molecules in saccharides (Kawai et al., 1992). Trehalose is a disaccharide osmolyte made of UDP-glucose and glucose-6-P; it is naturally synthesized by various organisms (e.g. bacteria, fungi, plants, invertebrates; Thompson, 2003; Feofilova et al., 2014; Lunn et al., 2014). It enhances protein stability and increases lipid bilayer equilibrium by placing water molecules near to the lipids' polar parts, thereby maintaining membrane fluidity (Donnamaria et al., 1994; Singer and Lindquist, 1998; Crowe, 2007; Jain and Roy, 2008). Work on bacteria, fungi and plants has shown that trehalose synthesis improves tolerance to abiotic stressors such as heat and desiccation. In insects, trehalose is the main sugar found in the hemolymph and fat body, where it has at least two functions: (1) serving as a source of energy during flight (Becker et al., 1996) and (2) acting as a cryoprotectant by reducing the supercooling point, thereby improving cold tolerance (Storey and Storey, 2012; Tang et al., 2018; Cline, 2018; Khani et al., 2007; Wen et al., 2016; Olsson et al., 2016). To our knowledge, only a single study has investigated trehalose regulation under conditions of heat stress in insects. Jin et al. (2018) showed that high temperatures can upregulate expression of the gene trehalose-6-phosphate synthase in the fat bodies of grass moths (Ostrinia furnacalis). The functional role of this compound is otherwise unexplored.

The Namib desert ant, Ocymyrmex robustior, is one of the world's most thermotolerant insects. Workers forage during the hottest part of the day, when sand surface temperatures reach up to 67°C and their body temperatures exceed 51°C (Marsh, 1985). Their diet is almost exclusively composed of heat-stricken arthropods (97.5%) and, on occasion, plant matter (Marsh, 1985). Recently, transcriptomic analyses have revealed that workers exposed to thermal stress have upregulated levels of isoforms for two genes involved in trehalose metabolism: facilitated trehalose transporter 1 and bifunctional trehalose-6-phosphate synthase/phosphatase (see Fig. S1; S. N. Araujo, R.P., Q. Willot, M. Defrance and S.A., unpublished). Here, we explored how this sugar metabolite affects heat tolerance in O. robustior. First, we tested whether trehalose-supplemented workers displayed increased survival following exposure to lethal temperatures. Second, we assessed whether worker survival was associated with metabolic levels of trehalose. Third, because heat shock can cause proteins to misfold and denature, we quantified heat-stress-induced protein damage by comparing protein aggregation levels in trehalose-supplemented workers and control workers. And fourth, we compared the relative expression of three genes involved in trehalose metabolism for these two groups of workers: the two genes mentioned above (facilitated trehalose transporter 1 and bifunctional trehalose-6-phosphate synthase/phosphatase) and the gene trehalase, which hydrolyzes trehalose.

Five colonies of Ocymyrmex robustior Stitz 1923 were collected near the Gobabeb Research Station in the Namib Desert (23°33'45''S, 15°02'31''E). A collection permit was issued by the Namibian National Commission on Research, Science and Technology (permit number RPIV01042022). Ants were reared under laboratory conditions for 2 weeks before the experiments began. The colonies were placed in plastic boxes (30×40×10 mm) with Fluon®-coated sides to prevent the ants from escaping. Each box had a thin layer of clean sand across its bottom. Colonies were supplied with glass test tubes (16×150 mm) for nesting. Water was present at the bottom of the tubes, which the ants could access via a moist cotton plug. The colonies were reared under constant environmental conditions: temperature of 25°C (±1°C), 12 h:12 h light:dark cycle and relative humidity of 30–40%. Each colony was split into two equal-sized groups that were given the same basic diet: pieces of cockroaches 2 or 3 times a week. However, one group also received a trehalose supplement (1 mol l⁻¹ solution), while the other group received a control supplement (water). Because O. robustior is a scavenger that feeds almost exclusively on dead arthropods under natural conditions (Marsh, 1985), we assumed that the control workers would not experience any carbohydrate deficiencies.

To determine the effect of trehalose on heat tolerance, we performed survival assays using the control and trehalose-supplemented workers. Thirty workers were randomly sampled from each group. They were placed individually in 100 ml cotton-plugged glass assay tubes that were then submerged in a 45°C water bath (W22, Julabo), where they remained for 8 h. No source of humidity was added. The temperature inside the tubes was monitored using 0.075-mm diameter thermocouples [Type K Thermocouple (Chromel/Alumel), RS Components, UK] connected to a digital thermometer (RS Pro RS52 Digital Thermometer, RS Components). The ants were checked every 10 min. A survival curve was established using the workers' time until death. Ants were considered dead once they had lost their locomotor abilities (i.e. muscular paralysis; Perez et al., 2021; Roeder et al., 2021).

Concentration levels of trehalose were measured for the control and trehalose-supplemented workers. For each group, 5 sets of 10 workers were weighed to the nearest 0.001 mg (Ohaus electronic microbalance) before being exposed to either a sublethal temperature of 43°C for 4 h (heat-stress treatment, HS) or a control temperature of 25°C for 4 h (non-heat stress treatment, NHS). These assays were performed using the same experimental setup described above. After treatment, ants were immediately snap frozen in liquid nitrogen.

To purify the low-molecular-weight carbohydrates, we used a modification of the method described by Soudi and Moharramipour (2012) (see also Sajwan et al., 2015). The ants were homogenized in 1 ml of 80% ethanol. After 8 min of centrifugation at 10,000 g, the supernatant was transferred to a new tube and dried at 37°C for 24 h. The pellet was then resuspended in pure H₂O. Trehalose levels were quantified using a Trehalose Assay Kit (Megazyme) in accordance with the manufacturer's instructions. First, reducing sugars were removed from the samples using an alkaline borohydride solution (10 mg ml⁻¹ of sodium borohydride in 50 mmol l⁻¹ sodium hydroxide). Then, we added NADP⁺/ATP and hexokinase/glucose-6-P dehydrogenase to the samples to convert d-glucose and glucose-6-P into gluconate-6-phosphate and NADPH. Finally, a trehalase solution was also added to convert any trehalose into d-glucose. The difference in NADPH absorbance (340 nm) between the second and third steps was measured. This difference served as a proxy for the quantity of trehalose in each sample. Trehalose concentrations standardized for ant body mass were compared between groups (trehalose-supplemented versus control workers) and treatments (HS versus NHS).

We compared protein aggregation levels in control versus trehalose-supplemented workers exposed to the HS versus the NHS treatment. For each group, 5 sets of 6 workers were weighed to the nearest 0.001 mg (Ohaus electronic microbalance) and then heat stressed at 43°C (HS) or maintained at 25°C (NHS) for 4 h. The ants were subsequently snap-frozen in liquid nitrogen. To perform the total protein extractions, we crushed the ant bodies in 1.5-ml microtubes using a pestle and added 1 ml of protein extraction buffer (0.1% Triton X-100, 50 mmol l⁻¹ HEPES, 1 mmol l⁻¹ EDTA, 150 mmol l⁻¹ NaCl, 0.5 mmol l⁻¹ PMSF, 0.1 mmol l⁻¹ DTT and 2.5 µg ml⁻¹ leupeptin at pH 7.3). The resulting mixture was homogenized for 3 min at maximum speed in a mixer mill using 2.8-mm zirconium oxide beads. The tubes were centrifuged for 10 min at 4°C and 1200 g. The supernatant was then filtered and transferred to new tubes using a 21-gauge needle (Sterican®, Braun, Kronberg, Germany), leaving behind large pieces of cuticles and organs. We repeated this procedure until we had obtained a homogeneous aqueous phase without any impurities. The supernatant was centrifuged for 1 h at 4°C and 100,000 g (Beckman Coulter Ultracentrifuge) to precipitate any protein aggregates; native proteins remained solubilized in the supernatant. The pellet was resuspended in 1 ml of denaturation buffer (6 mol l⁻¹ guanidine hydrochloride, 0.1 mol l⁻¹ NaH₂PO₄ and 10 mmol l⁻¹ Tris at pH 8); total protein content was determined using a BCA Assay Kit (Bio-Rad) in accordance with the manufacturer's instructions. Absorbance (592 nm) was recorded using a microplate reader. Aggregate concentration was standardized by dividing the protein concentration (µg µl⁻¹) by the initial mass (mg; i.e. before treatment) for each sample. The difference in aggregation (Δaggregation; µg µl⁻¹ mg⁻¹) between the NHS and HS treatments was calculated as follows: Δaggregation=protein aggregate concentration (HS)–protein aggregate concentration (NHS). Δaggregation was calculated for all the samples from the control and trehalose-supplemented workers, and the groups' means were compared.

We quantified the relative expression of three genes that encode enzymes involved in trehalose metabolism: bifunctional trehalose-6-phosphate synthase/phosphatase (hereafter, trehalose synthase), trehalase and facilitated trehalose transporter 1 (hereafter, trehalose transporter). Trehalose synthase catalyzes trehalose formation by binding together two glucose molecules (glucose-6-phosphate and UDP glucose) to form trehalose-6-phosphate (Tre6P); it then removes the phosphate from Tre6P to produce free trehalose. Trehalase catalyzes the opposite reaction, making the glucose available again. Trehalose transporter mediates the bidirectional transfer of trehalose between the cell and the hemolymph.

Five sets of 10 workers from the trehalose-supplemented group and the control group were heat stressed at 43°C (HS) or maintained at 25°C (NHS) for 4 h using the same experimental setup described above. The workers were immediately snap-frozen in liquid nitrogen. Total RNA was extracted using TRIzol (Invitrogen, Carlsbad, CA, USA) in accordance with the manufacturer's instructions. Samples were homogenized for 3 min at maximum speed in a mixer mill (MM 301, Retsch) using 2.8-mm zirconium oxide beads (Bertin Technologies, Montigny-le-Bretonneux, France). RNA levels were then quantified utilizing a NanoDrop 1000 spectrophotometer (Thermo Fisher Scientific, Ghent, Belgium). The absorbance ratios (260 nm/280 nm) ranged from 1.8 to 2.0 for all the samples, indicating that RNA purity was adequate. After treatment with DNase (DNase I, Thermo Fisher Scientific), 1 µg of total RNA was retrotranscribed using an RT system (SuperScript^® III Reverse Transcriptase, Thermo Fisher Scientific).

Real-time quantitative PCR (RT-qPCR) was performed using a Rotor-Gene Q apparatus (Qiagen). We used 20-μl reaction volumes, which contained 5 ng of template cDNA, 8 nmol l⁻¹ of total primer and 10 μl of SYBR^® (Premix Ex Taq™ II, Takara Bio USA). The following temperature program was employed: an initial incubation step at 95°C for 30 s, then 40 cycles at 95°C for 5 s, and finally an annealing step followed by an extension step, each at 60°C for 30 s. Melt curve analysis confirmed the presence of a single amplicon. For standardization purposes, RPL-32 expression was used. Fold changes (FCs) were calculated using the ΔΔC_t (relative fold gene expression) method described by Livak and Schmittgen (2001). For each replicate, ΔΔC_t was calculated by subtracting mean ΔC_t under NHS conditions from ΔC_t of each replicate. Mean ΔΔC_t values for the three genes were compared between the trehalose-supplemented group and the control group. For each gene, primers were designed using common isoform sequences, which we obtained from a transcriptomic analysis performed in our laboratory (unpublished results). These sequences were purchased after aligning the isoforms using CodonCode Aligner (CodonCode Corp., USA). Transcriptome sequencing, assemblage and annotation were performed as described in Perez et al. (2021). Primer sequences, amplicon sizes and melting temperatures are given in Table S1. Primer specificity was verified via PCR coupled with electrophoresis (1.2% agarose gel). All the primers produced single bands corresponding to expected amplicon sizes.

Survival curves were compared using a Mantel–Cox test (n=30). The effect of body mass, diet and temperature treatment on trehalose concentration were assessed through a mixed model [‘aov’ command in R: aov(trehalose_concentration∼body_mass*diet*temperature+(1|colony), data=dosage)] using trehalose concentration (diet) as a fixed effect and nest of origin as a random effect (n=5). We also performed a simple linear model [‘lm’ command in R: lm(trehalose_concentration∼body_mass, data=dosage)] in order to quantify the correlation (R²) between body mass and trehalose concentration. The RT-qPCR and protein aggregation data were analyzed utilizing Wilcoxon–Mann–Whitney tests (n=5). All statistical analyses were performed using RStudio (version 4.0; https://posit.co/).

Under conditions of heat stress, trehalose-supplemented workers survived significantly longer than did control workers (Mantel–Cox test, P<0.05; Fig. 1). Whereas survival decreased substantially from the first hours for the control workers, such decrease in survival was delayed after approximately 300 min for trehalose-supplemented ants. After 8 h, several workers from the trehalose-supplemented group were still alive, whereas all workers from the control group were dead.

We found an effect of body mass on trehalose concentration (mixed model: P<0.05); however, this effect was relatively weak (R²=0.22 considering a simple linear model). There was no combined effect of body mass with other variables (interactions mass:diet and mass:temperature: P=0.49 and P=0.75, respectively). As expected, trehalose-supplemented workers had higher concentration levels of trehalose (relative to body mass) than did control (not trehalose-supplemented) workers (mean±s.d., control NHS: 0.79±0.26 µg µl⁻¹, trehalose NHS: 3.97±1.5 µg µl⁻¹; mixed model for diet: P<0.05; Fig. 2). This result confirms that the ants assimilated the trehalose added to their diets. After the heat stress treatment, levels of trehalose were low and relatively stable in the control workers (0.68±0.14) but declined dramatically (±54%), if not significantly (temperature: P=0.08, interaction diet:temperature: P=0.43), in the trehalose-supplemented workers (1.83±0.56). This finding suggests that trehalose is converted to glucose, which is then funneled into other metabolic circuits, such as glycolysis or the pentose phosphate pathway (PPP).

Following the heat stress treatment, the increase in protein aggregates was 3 times lower in the trehalose-supplemented workers than in the control workers (mean±s.d.: 0.88±0.22 versus 2.81±0.61 ng µl⁻¹ mg⁻¹; Wilcoxon–Mann–Whitney, P=0.008) (Fig. 3).

In the trehalose-supplemented ants, heat stress induced an increase in mean relative expression for two of the genes (Fig. 4). Analysis of gene expression in HS and NHS workers indeed showed a three-fold increase in the expression of the gene trehalose synthase (3.3±0.27) and a nine-fold increase in the expression of the gene facilitated trehalose transporter (9.14±9.66). Expression of the gene trehalase remained relatively unchanged between heat-shocked and non-heat-shocked workers (1.54±1.57).

Heat stress elicited dramatically different gene expression patterns in control ants. Expression did not change for the gene trehalose synthase (1.25±0.46), whereas expression of both trehalase and facilitated trehalose transporter was downregulated four-fold (4.46±3.75 and 4.48±2.24, respectively). Expression of all three genes was significantly different between the trehalose-supplemented workers and the control workers (Wilcoxon–Mann–Whitney test, P<0.01 for all comparisons).

Owing to their high surface-to-volume ratio and ectothermic physiology, insects are vulnerable to temperature fluctuations and desiccation stress under normal conditions (Willmer et al., 2009). The stakes are raised in hot deserts such as the Namib, where heat and dryness can reach record levels (Lancaster et al., 1984). The synthesis of protective, low-molecular-weight metabolites, such as sugars or amino acids, is a useful response in such environments as these compounds can be rapidly accumulated under stressful conditions (Thompson, 2003). Our results highlight the protective role that trehalose plays in the Namib desert ant, O. robustior, one of the most thermophilic terrestrial animals on Earth (Marsh, 1985; Turner et al., 2000). We observed that high concentrations of trehalose considerably improved worker heat tolerance, likely because the molecule helps ensure greater protein stability. Furthermore, our gene expression analysis indicated that trehalose supplementation influenced trehalose regulation. Past research has shown that trehalose supplementation can enhance tolerance of several abiotic stressors (e.g. heat, desiccation, salt, hypoxia) in bacteria, fungi and plants (Chen and Haddad, 2004; Iordachescu and Imai, 2008). To our knowledge, our study is the first to demonstrate that trehalose supplementation can increase heat tolerance in insects.

More specifically, we discovered that trehalose-supplemented workers had higher levels of trehalose in their hemolymph and tissues, which was associated with a lower degree of protein aggregation following heat stress. Several models have been proposed to explain how trehalose improves protein homeostasis (Crowe et al., 1987; Green and Angell, 1989; Carpenter and Crowe, 1989). The most support exists for the preferential hydration model, which hypothesizes that trehalose promotes the formation of a thick water layer around proteins, slowing them down (Olssen et al., 2020). Thermodynamically, this process increases the energy barrier, thus impeding the transition between the native and misfolded states of proteins (Xie and Timasheff, 1997). Indeed, past work has shown that the presence of trehalose in biologically active cells can limit protein aggregation (Tapia and Koshland, 2014; Kim et al., 2018). However, it is also possible that trehalose-supplemented workers displayed greater heat tolerance because they benefited from enhanced membrane homeostasis owing to an increased number of trehalose-lipid hydrogen bonds (Pereira et al., 2004). For example, nematodes (Caenorhabditis elegans) fed trehalose suffered less membrane damage after experiencing desiccation stress (Erkut et al., 2011).

Our gene expression analysis showed that after exposure to heat stress, trehalose-supplemented workers displayed upregulated trehalose synthesis but stable trehalose hydrolysis. Such dynamics should have resulted in increased levels of trehalose. Contrary to this expectation, trehalose levels decreased dramatically although not significantly (Fig. 2). This close to significant decrease in trehalose concentration would arise because trehalose concentration of one of our samples in trehalose-supplemented NHS conditions was particularly high. This suggests that other mechanisms are acting to decrease trehalose levels. One possibility is that trehalase is activated via its phosphorylation by the protein kinase A or Ca²⁺/calmodulin kinase (Souza et al., 2002). Support for this idea comes from transcriptomic data showing that the gene Ca²⁺/calmodulin kinase is strongly upregulated in O. robustior (S. N. Araujo, R.P., Q. Willot, M. Defrance and S.A., unpublished). The situation was more ambiguous in control workers: following heat stress, trehalose synthesis remained stable but trehalose hydrolysis was downregulated. Consequently, trehalose levels should have increased. Yet, no such result was observed after 4 h of heat stress. One may not exclude, however, that induction of protective molecules would be triggered after a recovery time following a heat stress (Zhang et al., 2012; Semmouri et al., 2019). For instance, carbohydrate level increases after recovery from prolonged thermal stress in Drosophila melanogaster (Gruntenko et al., 2021). Thus, we cannot conclude that trehalose production in O. robustior is a response mechanism to heat stress, but rather that trehalose is maintained at stable levels instead of being accumulated or entirely consumed. Workers storing more sugar resources may therefore be more heat tolerant under natural conditions.

In response to heat stress, the gene trehalose transporter was upregulated in the trehalose-supplemented workers but downregulated in the control workers. Given that facilitated trehalose transporter 1 mediates the bidirectional transfer of trehalose between the cell and the matrix, we speculate that this transport mechanism may somehow be downregulated when trehalose concentrations are low. In contrast, when concentrations are high, it may be beneficial to allow trehalose to circulate and provide energy to the muscles (Thompson, 2003; Liu et al., 2013). Furthermore, high levels of circulating trehalose could feed the PPP, which generates pentose sugar and NADPH. The latter is a cofactor in antioxidative processes that are part of the heat stress response (Hayes et al., 2005; Ralser et al., 2007). Consistent with this hypothesis, genes encoding PPP-related proteins (e.g. fructose-bisphosphate aldolase and glucose-1-dehydrogenase) were upregulated during heat stress in O. robustior (S. N. Araujo, R.P., Q. Willot, M. Defrance and S.A., unpublished). Taken together, these observations suggest that trehalose supplementation, acting in synergy with other cellular mechanisms, may boost heat tolerance. Indeed, trehalose supplementation has been found to enhance PPP activity and thereby improve thermal tolerance in the fungus Pleurotus ostreatus (Yan et al., 2021).

Our results raise three main questions. First, did trehalose-supplemented workers survive longer than control workers under heat stress conditions because of differences in nutrition? As noted above, O. robustior is a scavenger that feeds almost exclusively on dead arthropods (Marsh, 1985), which are high in protein. However, carbohydrates are also present and account for 7.2% of insect dry mass (Bell, 1990). Therefore, it seems unlikely that control workers were malnourished or underfed because they were given large quantities of cockroaches. Furthermore, previous work in the ant Aphaenogaster picea has shown that starvation actually delays time to knockdown under conditions of heat stress (Nguyen et al., 2017). Second, how does O. robustior obtain carbohydrates under natural conditions? Because trehalose enhances resistance to heat stress, workers would be expected to seek out carbohydrates. As mentioned above, carbohydrates were present in the dead insects scavenged by workers. Another source of carbohydrates is plant matter, especially seeds, which the workers harvest. And animals that consume a diet low in carbohydrates (i.e. carnivores) have evolved a high rate of gluconeogenesis, which generates glucose via the catabolism of non-carbohydrate carbon substrates (Walton and Cowey, 1983; Washizu et al., 1999; Sanders, 2016). Third, would we have seen the same results if we had given the workers a carbohydrate other than trehalose? It seems likely. A previous study indeed showed that carbohydrates such as sucrose or fructose enhance thermal tolerance in insects (Bujan and Kaspari, 2017). Furthermore, any source of sugar can be converted into glucose, which is used to produce trehalose – the main carbohydrate energy storage molecule used by insects. That said, we chose to focus on trehalose in this study because transcriptomic research showed that genes involved in trehalose metabolism were strongly upregulated in O. robustior under conditions of heat stress (S. N. Araujo, R.P., Q. Willot, M. Defrance and S.A., unpublished). In addition, previous proteomics research conducted in vitro has shown that trehalose provides better biomolecular protection against thermal stress than do sucrose, maltose, glucose and fructose (Crowe et al., 1996; Sola-Penna and Meyer-Fernandes, 1998; Porchia et al., 1999; Cline, 2018; Olssen et al., 2020).

In conclusion, this study has demonstrated that trehalose significantly boosts thermotolerance in the Namib desert ant, O. robustior, by providing a significant source of energy and possibly by acting as a protein stabilizer. Further research should explore whether trehalose supplementation enhances thermal tolerance of insects in general, and whether this sugar acts directly or indirectly to improve heat resistance.

We thank Quentin Willot for his help with the field sampling, Marie Overtus for her help with ultra-centrifugation procedure, and Gillian Maggs-Kölling, Eugene Marais and the Gobabeb Namib Research Institute for their logistical support. Thanks also to J. Pearce-Duvet for her language-editing services.