Naked mole-rats reduce the expression of ATP-dependent but not ATP-independent heat shock proteins in acute hypoxia

Hypoxic environments are common in nature, and for terrestrial mammals, these environments include life at high altitude and in underground burrows. In hypoxia, cellular aerobic respiration can become severely compromised, which can uncouple ATP supply from demand and disrupt cellular energy balance (Buck and Pamenter, 2006; Hochachka, 1986; Land et al., 1993). In hypoxia-intolerant mammals, prolonged hypoxic exposure can lead to organ failure and, ultimately, death. Conversely, some animals have evolved systemic and cellular adaptations to hypoxia, which enables these hypoxia-tolerant species to survive in hypoxic environments (Bickler and Buck, 2007; McClelland and Scott, 2019).

In addition to disrupting energy balance, hypoxia can also contribute to a loss of protein integrity via disruptions of redox balance, which can alter protein structure, and lead to protein damage, aggregation or misfolding, especially of mitochondrial proteins (Kaufman and Crowder, 2015; Kaufman et al., 2017). Impaired proteostasis during hypoxia may result in loss of cellular function and tissue damage in hypoxia-intolerant species. To avoid this, a key to surviving hypoxia is to maintain good proteostasis via the unfolded protein response (UPR) (Hetz et al., 2015). The two primary components of the UPR are (1) the ubiquitin–proteasome system, which limits the accumulation of damaged proteins by degrading denatured non-functional proteins and preventing their accumulations from affecting healthy proteins and the cell, and (2) molecular chaperones, also known as heat shock proteins (HSPs). HSPs are classified based on their function as either holdases, foldases or disaggregases (Díaz-Villanueva et al., 2015). Holdases are ATP independent and bind to and passively stabilize denatured proteins and prevent aggregation. Foldases are ATP dependent and actively refold denatured proteins. Disaggregases are also ATP dependent and bind to and disaggregate protein clumps. HSPs can also be classified according to their molecular weight: HSP10, 27, 40, 60, 70, 90, 110, etc.

HSPs are known to be upregulated in hypoxia-intolerant species during hypoxia, including in mice, rats, rabbits, piglets, flies, nematodes and estuarine fishes (Baird et al., 2006; David et al., 2006; Liu et al., 2013; Mestril et al., 1994; Shen et al., 2005; Tiedke et al., 2014; Tokyol et al., 2005). However, few studies have examined the HSP response to acute hypoxia in hypoxia-tolerant organisms. This is a significant gap in our knowledge as it is important to understand the role of HSPs in cytoprotective adaptations to hypoxia in animals for which hypoxia is not a stress per se. One study of the anoxia-tolerant western painted turtle found that HSP70 became significantly elevated in the brain (1.9-fold), heart (3.5-fold), liver (1.7-fold) and muscle (5.6-fold) after a 30 h forced dive (Ramaglia and Buck, 2004). In addition, HSP90 increased significantly in the brain (5.6-fold), liver (2.1-fold) and muscle (2.4-fold) after a 24 h forced dive (Ramaglia and Buck, 2004). Similarly, in the anoxia-tolerant crucian carp, HSP70 expression increased drastically in the brain and heart after 7 days of anoxia, while HSP90 was significantly elevated in the heart only after 1 day of anoxia (Stensløkken et al., 2010). Recently, Wu et al. (2018) observed that HSP60 was upregulated 1.73-fold and 1.59-fold after 4 and 24 h of anoxia, respectively, in wood frog brain. Conversely, no studies have addressed HSP responses to hypoxia in any hypoxia-tolerant mammalian model.

One of the most hypoxia-tolerant mammals is the naked mole-rat (Heterocephalus glaber). Studies in this species have revealed several physiological adaptations that are protective against hypoxia including profound depression in metabolism during acute hypoxia and metabolic plasticity following prolonged hypoxia (Chung et al., 2016; Pamenter et al., 2015). A key finding was that naked mole-rats can suppress their metabolism by up to 85% in acute hypoxia (Pamenter et al., 2018). Marked metabolic rate suppression is very important in hypoxia, as it reduces ATP demand to match limited ATP supply (Buck and Pamenter, 2006). Hypoxic metabolic suppression in this species is achieved in part by decreasing body temperature to near-ambient levels and reducing physical activity (Houlahan et al., 2018; Ilacqua et al., 2017; Kirby et al., 2018). However, the underlying cellular mechanisms of metabolic suppression are still poorly understood. Naked mole-rats can also use fructose as an alternative fuel source during O₂ deprivation (Park et al., 2017). In addition, naked mole-rats have an enhanced blood O₂ carrying capacity in hypoxia owing to the expression of a high-affinity hemoglobin (Johansen et al., 1976).

Although systemic physiological and behavioral adaptions to acute hypoxia have begun to be investigated in this fascinating mammalian model, few studies have explored the cellular mechanisms of hypoxia tolerance in naked mole-rats. Specifically, little is known regarding how naked mole-rats maintain protein integrity in hypoxia. Importantly, naked mole-rats do express functional HSPs, and the basal and heat-shock-induced expressions of most HSPs are significantly higher in fibroblasts of naked mole-rat compared with mouse, suggesting that naked mole-rat HSPs are endogenously primed for heat stress (Pride et al., 2015), and presumably also for hypoxia. Given this knowledge, we explored changes in HSPs in a tissue-specific manner during acute hypoxia exposure. Because HSPs are putatively cytoprotective and naked mole-rats are hypoxia tolerant, we hypothesized that HSP gene and protein expression would be upregulated in naked mole-rat tissues to maintain cellular integrity and proteostasis during hypoxia.

Naked mole-rats (Heterocephalus glaber Rüppell 1842) were group-housed in interconnected multi-cage systems at 30°C and 21% O₂ in 50% humidity with a 12 h:12 h light:dark cycle. Animals were fed fresh tubers, vegetables, fruit and Pronutro cereal supplement ad libitum. Animals were not fasted prior to experimental trials. All experimental procedures were approved by the University of Ottawa Animal Care Committee in accordance with the Animals for Research Act and by the Canadian Council on Animal Care. Non-breeding (subordinate) naked mole-rats do not undergo sexual development or express sexual hormones, and thus we did not take sex into consideration when evaluating our results (Holmes et al., 2009).

Seventeen naked mole-rats were randomly divided into four experimental treatment groups: a control normoxia group (21% O₂; n=5), and three hypoxia treatment groups (7% O₂) of 4, 12 and 24 h duration (n=4 each). Naked mole-rats were treated in their colony groups and home cages, and were supplied with fresh fruits and vegetables during treatment. After treatment, naked mole-rats were removed from the chamber and immediately killed by cervical dislocation followed by rapid decapitation within 15 s. Brain, heart, liver and temporalis muscle were rapidly extracted over ice, flash-frozen in liquid N₂ and then stored at −80°C until analysis.

Frozen tissues were ground into a fine powder under liquid N₂ using a mortar and pestle. For western blot analysis, 50–100 mg of frozen powdered tissue was homogenized on ice in 1 ml of homogenization buffer [3 mmol l⁻¹ sucrose, 3 mmol l⁻¹ dithiothreitol (DTT), 1 mmol l⁻¹ EDTA and protease inhibitor cocktail; Sigma-Aldrich, Oakville, ON, Canada] for 20–30 s and then sonicated three times for 10 s each. Cell lysates were then centrifuged at 4000 g for 10 min at 4°C. The resulting pellet was discarded and supernatants were centrifuged again at 13,000 g for 45 min. The resulting pellet was re-suspended in homogenization buffer and stored at −20°C. Protein concentrations were determined using Bradford assays (Bradford, 1976) (Sigma-Aldrich).

For gel loading, loading buffer [65.8 mmol l⁻¹ Tris-HCl with pH 6.8, 26.3% (w/v) glycerol, 2.1% SDS, 0.05% 2-mercaptoethanol, 0.01% Bromophenol Blue; Bio-Rad Laboratories, Mississauga, ON, Canada] was added to the protein sample in 1:1 v/v ratio and boiled at 95°C for 5 min. Protein samples and an unstained molecular marker (Bio-Rad) were loaded onto 10% SDS-polyacrylamide stain-free mini-PROTEAN^® gels (Bio-Rad) and subjected to electrophoresis in running buffer (Bio-Rad) at 200 V for 45 min (using Bio-Rad PowerPac™). Precision Plus Protein™ Unstained Protein Standards (Bio-Rad) were used to determine the migration and size of proteins on the gel. The proteins were then electrophoretically transferred onto an Immun-Blot^® PVDF membrane (Bio-Rad) in transfer buffer (Bio-Rad) at 20 V for 90 min (using Bio-Rad Trans-Blot^® SD Cell). The membranes were blocked with TBS-T (20 mmol l⁻¹ Tris, 150 mmol l⁻¹ NaCl, pH 7.5, 0.1% Tween20; Thermo Fisher Scientific, Nepean, ON, Canada) with 5% skim milk (Bio-Rad) for 1 h at room temperature. Membranes were then washed three times each for 5 min in TBS-T and then incubated in primary HSP antibody overnight at 4°C. Membranes were then washed again three times each for 5 min in TBS-T and then incubated in secondary horseradish peroxidase (HRP)-linked antibody for 1 h at room temperature. Then, membranes were again washed three times each for 5 min in TBS-T. Immunoreactivity was visualized using a chemiluminescent technique by applying the detecting agent Clarity™ ECL substrates (Bio-Rad) for 5 min and imaged (using Bio-Rad Universal Hood II and ChemiDoc™ XRS+). Finally, band intensity was analyzed using Image Lab Software (Bio-Rad) for relative HSP levels using total protein as loading control.

The following antibodies were used: Hsp27 Ab (Santa Cruz Biotechnology, Santa Cruz, CA, USA), catalog no. sc-13132, lot L1015–1:1000 dilution; Hsp40 Ab (Cell Signaling Technology, Danvers, MA, USA), catalog no. 4871S, lot 2–1:1000 dilution; Hsp70 Ab (Santa Cruz Biotechnology), catalog no. sc-373867, lot D1114–1:350 dilution; Hsp90 Ab (Cell Signaling Technology), catalog no. 4877S, lot 3–1:1000 dilution; anti-rabbit IgG HRP-linked goat Ab (Cell Signaling Technology), catalog no. 7074S, lot 26–1:1600 dilution; and anti-mouse IgG_K HRP-linked Ab (Santa Cruz Biotechnology), catalog no. sc-516102, lots L2017 and B0217–1:1000 dilution.

Total RNA was extracted from 50–100 mg of frozen powdered tissue using TRIzol^® LS reagent (Thermo Fisher Scientific) according to the manufacturer's instructions. The RNA concentrations and purities were determined (NanoDrop^® ND-1000 Spectrophotometer) and stored at −80°C. To ensure the RNA samples were not contaminated with genomic DNA, samples were incubated with DNaseI (Thermo Fisher Scientific; 2 µg RNA in 5 µl nuclease-free H₂O, 1 µl 10× DNaseI reaction buffer, 1 µl DNaseI, 3 µl nuclease-free H₂O) at room temperature for 12 min. This was followed by 10 min incubation with 1 µl EDTA at 65°C to inactivate the DNaseI. Next, the RNA template was combined with 375 ng random hexamers/125 ng oligo dTs (Integrated DNA Technology, Kanata, ON, Canada) and dNTPs (Thermo Fisher Scientific; 0.5 mmol l⁻¹ final concentration) and incubated at 65°C for 5 min. First-strand reaction buffer, RNaseOut™ and DTT (Thermo Fisher Scientific; 4 µl FS buffer, 1 µl DTT, 0.5 µl RNaseOut™, 0.5 µl nuclease-free H₂O) were added and the mixture was incubated at 42°C for 2 min to inhibit ribonuclease activity and preserve the RNA for reverse transcription. Lastly, 1 µl SuperScript™ II reverse transcriptase (Thermo Fisher Scientific) was added for a total volume of 20 µl and incubated at 42°C for 50 min followed by inactivation at 75°C for 15 min. The cDNA samples were then stored at −20°C.

qPCR was performed on these cDNA samples using Maxima SYBR Green qPCR master mix (Thermo Fisher Scientific) [6.25 µl SYBR Green master mix, 200 nmol l⁻¹ each gene-specific sense and antisense primers (Table 1), 1.0 µl cDNA sample, 4.75 µl nuclease-free H₂O]. qPCR was performed on 40 cycles of: melting at 95°C for 15 s, followed by annealing, elongation and acquiring steps at 60°C for 60 s (using Qiagen, Rotor-Gene Q). Standard curves were constructed by pooling 10 µl of all the samples together; 1×, 4×, 16× and 64× serial dilutions were used. All samples were run in duplicate. Non-template controls and melting curves were used to ensure that the primers were specific and there was no contamination. Two housekeeping genes, actin and EEF2 (encoding the eukaryotic elongation factor 2 protein), were used as internal controls. Gene expression analysis was performed using the NORMA-Gene normalization method (Heckmann et al., 2011).

For detection of differences in HSP protein or gene expression, two-way ANOVA (with organ and hypoxia treatment as independent variables and protein level or gene expression as the dependent variable) was performed using GraphPad Prism software. Dunnett's post hoc analysis was conducted for specific statistically significant differences with respect to the control group within an organ, with a multiple comparison corrected α-level of 0.05. All data were then presented as means±s.e.m.

The gene profile of Hsp27 was consistent globally. Hsp27 mRNA levels stayed constant for the first 12 h of acute hypoxia in all organs (Fig. 1A). Interestingly, at the 24 h mark, we observed significant global reductions in Hsp27 mRNA across all organs tested: 74% in the brain, 74% in the heart, 87% in the liver and 83% in the muscle (Fig. 1A). By contrast, HSP27 protein levels remained unchanged throughout the 24 h hypoxic treatment in the brain, heart and muscle (Fig. 1B, blue, red and purple bars, respectively). However, in the liver, the HSP27 protein level significantly increased by 90% after 4 h of acute hypoxia (Fig. 1B, green bars). Conversely, after 12 h, liver HSP27 was significantly reduced by 66% compared with the normoxic control.

The gene profile of Hsp40 was similar to that observed for Hsp27. Specifically, the Hsp40 mRNA levels remained unchanged for the first 12 h of acute hypoxia in most organs, except in the liver (Fig. 2A). Then, at the 24 h mark, Hsp40 was significantly reduced by 79% in the brain, 79% in the heart and 81% in the muscle (Fig. 2A, blue, red and purple bars, respectively). Unlike in other organs, liver Hsp40 mRNA was significantly upregulated by 317% at 12 h and 293% at 24 h of hypoxic treatment compared with the control group (Fig. 2A, green bars). By contrast, HSP40 protein levels remained at control levels throughout the 24 h hypoxic treatment in the brain, heart and muscle (Fig. 2B blue, red and purple bars, respectively). Consistent with the gene data, in the liver, HSP40 protein levels were significantly increased by 72% after both 12 and 24 h of acute hypoxia treatment with respect to the control group (Fig. 2B, green bars).

Hypoxia-induced changes in Hsp70 gene expression varied in direction, magnitude and time across different tissues. Hsp70 mRNA shared similar patterns in the brain and the heart, wherein both were unchanged at 4 h, significantly upregulated to 199% of that of the control at 12 h, but then significantly downregulated by 55%, with respect to normoxic controls, at 24 h of hypoxia (Fig. 3A, blue and red bars, respectively). In the liver, Hsp70 was significantly reduced by 80% at 12 h and by 93% at 24 h (Fig. 3A, green bars). Finally, in muscle, Hsp70 remained unchanged after 4 and 12 h and was then significantly reduced by 70% at the 24-h mark compared with controls (Fig. 3A, purple bars). Similarly, hypoxia-induced HSP70 responses were also different in different tissues. In the brain, HSP70 protein expression remained unchanged after 4 and 12 h of hypoxia and was then significantly reduced by 57% at 24 h compared with controls (Fig. 3B, blue bars). In the heart, after just 4 h, HSP70 protein expression increased significantly to 217% of that of the control (Fig. 3B, red bars). However, at 12 h, HSP70 was significantly reduced by 74% of controls, but, by 24 h, this reduction became non-significant at 46% with respect to normoxic controls (Fig. 3B, red bars). In the liver, HSP70 remained unchanged after 4 and 12 h of hypoxia and was then significantly reduced by 54% at the 24-h mark with respect to normoxic controls (Fig. 3B, green bars). Finally, in the muscle, HSP70 levels were significantly reduced by 86% and 83% after 12 and 24 h, respectively (Fig. 3B, purple bars).

Unlike Hsp70, Hsp90 gene expression was globally decreased during hypoxia across all organs tested. In both the brain and the heart, Hsp90 mRNA decreased by 24% at 4 h, 58% at 12 h and 87% at 24 h of hypoxic treatment (Fig. 4A, blue and red bars, respectively). In the liver, Hsp90 mRNA decreased significantly by 68% at 4 h, 64% at 12 h and 99% after 24 h of hypoxia (Fig. 4A, green bars). Finally, in the muscle, Hsp90 remained unchanged after 4 h but was then significantly reduced by 70% at 12 h and by 84% at 24 h of hypoxia (Fig. 4A, purple bars). The HSP90 protein response had a similar pattern to that of the Hsp90 gene profile. In the brain, the HSP90 protein level was significantly reduced by 20% at 4 h, 35% at 12 h and 23% at 24 h of hypoxia (Fig. 4B, blue bars). Similarly, in the heart, HSP90 decreased significantly by 24% at 4 h, 41% at 12 h and 62% at 24 h of hypoxia (Fig. 4B, red bars). In the liver, HSP90 remained unchanged after 4 h, but was significantly reduced by 45% at 12 h and 61% at 24 h (Fig. 4B, green bars). In the muscle, HSP90 also remained unchanged after 4 h, and was also significantly reduced by 83% at 12 h and 78% at 24 h in hypoxia (Fig. 4B, purple bars).

In the present study, we examined the HSP response during 24 h of hypoxic exposure in the hypoxia-tolerant naked mole-rat for the first time. Our study produced two salient findings. First, HSP27 and HSP40 expression remained largely unchanged throughout the hypoxic treatment. Second, HSP70 and HSP90 expression were globally reduced in all tissues during the hypoxic treatment. These results contradict our initial hypothesis that HSP gene and protein expression would be upregulated in naked mole-rat tissues to maintain cellular integrity and proteostasis during hypoxia. Nonetheless, closer examination of the ATP dependence of these HSPs suggests that naked mole-rats prioritize energy savings during hypoxia and regulate their HSP response accordingly to provide some degree of proteostasis at a minimal energetic cost.

Despite refuting our hypothesis, the divergent changes in HSPs in our study are intriguing. The smaller HSPs, HSP27 and HSP40, remained unchanged in hypoxia, whereas the larger HSPs, HSP70 and HSP90, were globally reduced in hypoxia. These divergent patterns may be explained by the functional classification and ATP dependency of these HSPs. Specifically, the large HSPs (HSP70 and HSP90) are classified as foldases and are ATP-dependent chaperones that actively refold unfolded proteins via ATP hydrolysis (Díaz-Villanueva et al., 2015). Because these larger foldases require ATP to function, their downregulation in an ATP-deficient environment such as hypoxia would help to conserve energy for other more important cellular processes. Interestingly, the downregulation of these HSPs is delayed such that in the first 12 h of hypoxia, when cellular ATP is presumably less depleted, there is an activation of these ATP-dependent HSPs for cytoprotection of hypoxia-induced protein damage. Indeed, we even observed an upregulation of Hsp70 mRNA in the brain and the heart at 12 h. However, during longer periods of hypoxia (24 h), when ATP levels are presumably compromised and/or metabolic rate suppression is enhanced, these larger HSPs are downregulated at both the mRNA and protein level. Conversely, the small HSPs (HSP27 and HSP40) function as holdases: they are ATP-independent co-chaperones that recognize and stabilize unfolded proteins and bring them to the foldases for re-folding (Díaz-Villanueva et al., 2015). Because these smaller holdases do not require ATP, their function is presumably maintained during prolonged hypoxia despite any hypoxia-related ATP deficit. In summary, the initial trigger of hypoxia results in the activation of ATP-dependent HSPs; however, over time there appears to be a switch from relying on ATP-dependent to ATP-independent HSP activity in hypoxia. This would presumably help conserve energy during long-term hypoxia, when ATP levels may be compromised.

Although we do not see global downregulation of HSP expression during hypoxia, Hsp gene transcription exhibited a very consistent global trend such that all four Hsp genes examined were significantly downregulated after 24 h of hypoxia in all four organs tested. Because gene expression is the precursor to protein expression, this suggests that a longer hypoxic exposure would cause a global downregulation in the expression of all four HSP proteins. Indeed, although HSP27 and HSP40 do not require ATP for their normal cellular function, ATP is nonetheless required for protein synthesis to maintain the expression of these HSPs. Thus, it would become increasingly energetically expensive for the cell to maintain HSP protein levels during prolonged hypoxia, given the reduced energy production when O₂ is limited.

Taken together, our gene and protein expression results suggest that energy conservation is prioritized over cytoprotective protein chaperoning in naked mole-rats during hypoxia. Indeed, protein synthesis is one of the most energetically expensive cellular processes (Stouthamer, 1973), and many studies have demonstrated that translational arrest of protein synthesis is a hallmark of hypoxia tolerance. For example, in the heart of the anoxia-tolerant red-eared slider turtle, protein synthesis is reduced 3-fold after both 2 and 3 h of anoxia, and mitochondrial protein expression is particularly suppressed (Bailey and Driedzic, 1996). Similarly, the fractional rate of protein synthesis is reduced by 92% in hepatocytes of western painted turtles after 12 h of anoxic treatment (Land et al., 1993). In the liver of hypoxia-treated anoxia-tolerant goldfish, protein synthesis drops by 94% compared with normoxic controls, along with a 5.5-fold increase in AMPK activity (Jibb and Richards, 2008). AMPK is a molecular switch that inhibits anabolic processes while promoting catabolic processes to maintain energy balance in a low-energy situation, including protein synthesis (Hardie, 2007). Finally, a reduction in protein synthesis is also observed in the liver, heart and muscle of the anoxia-tolerant crucian carp in hypoxia (Smith et al., 1996).

As briefly discussed in the Introduction, an alternative means of cytoprotection and maintaining good proteostasis in hypoxia is the selective degradation of damaged proteins. Besides being non-functional, damaged proteins can accumulate and aggregate with each other and interfere with other normal cellular functions. Many studies have demonstrated that proteolytic capacity is often upregulated in hypoxia. For example, LON (a quality-control mitochondrial protease) mRNA expression is significantly induced in the heart, lung and liver of mice exposed to 10% O₂ for 3 weeks by 4.7-fold, 3.3-fold and 2.9-fold, respectively, compared with normoxic controls (Fukuda et al., 2007). Another study found that hypoxia-tolerant hard clams and oysters are able to maintain steady-state activity of ATP-dependent and -independent mitochondrial proteases under anoxia and 5% O₂ (Ivanina and Sokolova, 2016). In contrast, anoxia and hypoxia strongly suppress ATP-dependent mitochondrial protease activity in hypoxia-intolerant scallops; this suppression is also associated with the accumulation of oxidative damage to mitochondrial proteins (Ivanina and Sokolova, 2016). Similarly, naked mole-rats have a unique ubiquitin–proteasome system that gives them generalized resilience against a variety of stressors. The proteasome of naked mole-rats has significantly higher chemotrypsin- and trypsin-like catalytic activities relative to mice (Rodriguez et al., 2012). This means that naked mole-rats are more efficient at removing damaged or unfolded proteins to prevent their aggregation from affecting healthy proteins or harming the cell during stress. More interestingly, the naked mole-rat proteasome is more resistant to inhibition, both stress and drug induced, through an association with small molecular factors HSP40 and HSP70 (Rodriguez et al., 2014). These molecular chaperones both enhance the naked mole-rat proteasome activities and preserve its function during stressful conditions, although the role of these pathways in hypoxia has yet to be investigated.

In evaluating our results, it is important to consider that naked mole-rats can tolerate O₂ levels as low as 0% for 18 min, 3% for hours, and 8% for days to weeks in a laboratory setting (Chung et al., 2016; Pamenter et al., 2015; Park et al., 2017). It is therefore possible that 7% O₂ is not sufficiently stressful to trigger the HSP response in naked mole-rats. Thus, examining the HSP response at a lower O₂ concentration might provide a better understanding of its role in hypoxia tolerance. However, we consider this unlikely as we did observe changes in both gene and protein expression at this level of hypoxia, suggesting active regulation of these chaperones. In addition, HSPs are regulated via phosphorylation; however, our primary antibodies were not specific and bind to both isoforms. Thus, measuring the phosphorylated form of these HSPs, using phosphorylation-specific primary antibodies, would provide a better understanding of the regulation of HSP activity during hypoxia in this species. Nonetheless, this study is the first to address hypoxia-mediated regulation of HSPs in a hypoxia-tolerant mammal and represents a useful first step in this research area.

In summary, and contrary to results in other hypoxia- and anoxia-tolerant vertebrates, hypoxia exposure of 7% O₂ for 24 h did not result in the upregulation of four molecular chaperones (HSP27, HSP40, HSP70 and HSP90) in naked mole-rats. Interestingly, we found that ATP-independent HSP27 and HSP40 remained unchanged (except the liver), whereas ATP-dependent HSP70 and HSP90 were globally reduced by hypoxia. Based on these results, we propose that energy conservation is prioritized over cytoprotective protein chaperoning in naked mole-rats during hypoxia. We speculate that these HSP responses are a unique adaptation in this species that enables them to minimize energy expenditure to maintain a healthy proteostasis, allowing them to survive in severely but intermittently hypoxic underground burrows.

We would like to thank the University of Ottawa animal care and veterinary services team for their assistance in animal handling and husbandry.