Role of unfolded protein response and ER-associated degradation under freezing, anoxia, and dehydration stresses in the freeze-tolerant wood frogs

Abstract

The North American amphibian, wood frogs, Rana sylvatica are the most studied anuran to comprehend vertebrate freeze tolerance. Multiple adaptations support their survival in frigid temperatures during winters, particularly their ability to produce glucose as natural cryoprotectant. Freezing and its component consequences (anoxia and dehydration) induce multiple stresses on cells. Among these is endoplasmic reticulum (ER) stress, a condition spawned by buildup of unfolded or misfolded proteins in the ER. The ER stress causes the unfolded protein response (UPR) and the ER-associated degradation (ERAD) pathway that potentially could lead to apoptosis. Immunoblotting was used to assess the responses of major proteins of the UPR and ERAD under freezing, anoxia, and dehydration stresses in the liver and skeletal muscle of the wood frogs. Targets analyzed included activating transcription factors (ATF3, ATF4, ATF6), the growth arrest and DNA damage proteins (GADD34, GADD153), and EDEM (ERAD enhancing α-mannosidase-like proteins) and XBP1 (X-box binding protein 1) proteins. UPR signaling was triggered under all three stresses (freezing, anoxia, dehydration) in liver and skeletal muscle of wood frogs with most tissue/stress responses consistent with an upregulation of the primary targets of all three UPR pathways (ATF4, ATF6, and XBP-1) to enhance the protein folding/refolding capacity under these stress conditions. Only frozen muscle showed preference for proteasomal degradation of misfolded proteins via upregulation of EDEM (ERAD). The ERAD response of liver was downregulated across three stresses suggesting preference for more refolding of misfolded/unfolded proteins. Overall, we conclude that wood frog organs activate the UPR as a means of stabilizing and repairing cellular proteins to best survive freezing exposures.

Introduction

All living organisms on Earth must deal with changing environmental conditions that can influence their physiology and lifespan (Storey and Storey 2005). This includes a multitude of environmental stresses such as fluctuation in oxygen levels, the availability of food and water, and variation in temperature. Organisms must be flexible to endure the inherent variability of their environments on both daily and seasonal time frames (Hawkins and Storey 2020). Extreme changes outside of the normal range can cause structural or metabolic damage to cells/tissues or, ultimately, even death.

The North American wood frog, Rana sylvatica, is one of only a few extreme survivalists among vertebrates that can tolerate whole body freezing during the winter and many studies have focused on the molecular, physiological and biochemical adaptations that contribute to survival (Storey and Storey 2017; Layne and Lee 1995). The frogs begin to freeze below the equilibrium freezing point of their body fluids (about − 0.5 °C) and survive the conversion of up to 65–70% of their total body water into extracellular ice (Storey and Storey 2017). Freezing is challenging as it brings a multitude of consequences and stresses that must be dealt with during the winter months in order that frogs can thaw unharmed in the spring and resume normal life. While frozen, physiological changes faced by wood frogs include cessation of heartbeat, blood circulation, breathing, and detectable brain and muscle activity (Storey and Storey 2017). Ice formation in extracellular compartments also places two major stresses on cells and organs: (a) ischemia imposed due to the interruption of breathing and blood flow, and (b) cell dehydration and osmotic stress owing to water exit into extracellular ice crystals (Costanzo et al. 2013; Storey and Storey 2017).

Freezing protection is accomplished by multiple adaptations. Some are triggered immediately when freezing begins. For example, ice nucleation on the skin sends immediate adrenergic signals that trigger liver glycogenolysis to produce and export huge amounts of glucose that acts as a cryoprotectant and is rapidly distributed throughout the body before freezing can shut down blood circulation (Storey and Storey 1985; Singh and Storey 2020). Other adaptations occur more slowly in response to rising ischemia and osmotic stresses and address specific pro-survival needs of different organs as well as implementing overall metabolic rate depression (MRD) and enhancing antioxidant defenses to deal with ischemia/reperfusion stress and associated reactive oxygen species (ROS) generation (Storey 1998; Rider et al. 2006; Storey and Storey 2019; Wu et al. 2018).

All of these stresses have potentially negative consequences for intracellular metabolism and the impact of anoxia/ischemia and osmotic/dehydration stresses on cellular processes that must remain active (but at low levels) and/or provide necessary protective measures. One of these processes is protein synthesis that involves not just production of peptide chains but also folding of proteins into the correct functional conformations and addition of necessary posttranslational modifications. The endoplasmic reticulum (ER) is the site of protein folding and modification and also acts as a quality control checkpoint before proteins can take up their intracellular jobs or be secreted for extracellular functions (Vitale et al. 1993; Almanza et al. 2019). Protein folding is the function of ER resident chaperone proteins and our previous studies have shown increased expression of central regulators of ER stress, the glucose regulated proteins (Grp78 and Grp94), in liver and muscle in response to freezing in wood frogs (Storey and Storey 2019). Comparable studies of environmental stress in other species have shown similar responses by the ER machinery to fold or refold proteins, including responses to oxygen limitation by anoxia tolerant turtles (Trachemys scripta elegans) and hibernation-induced MRD in ground squirrels (Spermophilus tridecemlineatus) (Krivoruchko and Storey 2013; Mamady and Storey 2008). Combined action of the unfolded protein response (UPR) and the ERAD (ER-associated protein degradation) pathways act to restore ER balance. In the present study, we hypothesized that the UPR and ERAD pathways would be activated in wood frogs, not only by freezing but also by two of the component stresses affecting cells during freezing: anoxia/ischemia and dehydration.

The UPR is activated under stress conditions when Grp78 is confronted with a surge of unfolded proteins accumulating in the ER. This triggers Grp78 to dissociate from interactions with three receptor proteins on the ER membrane: inositol-requiring 1 kinase (IRE1), PKR-like endoplasmic reticulum kinase (PERK), and activating transcription factor 6 (ATF6) (Fig. 1) (Lee 2005). These proteins then trigger three major signaling pathways, PERK/ATF4, IRE1/XBP1 (X-box binding protein 1), and ATF6, that each play a role in cell survival (Fig. 1). These pathways act in concert to quickly activate responses that lead to refolding of misfolded proteins or direct proteins into proteasomal degradation via the ERAD pathway (Kim et al. 2006). Both IRE1 and PERK are transmembrane protein kinases that are activated via autophosphorylation under ER stress (Fig. 1) (Adams et al. 2019). Activated PERK (p-PERK) phosphorylates eIF2α (eukaryotic initiation factor 2α, phospho eIF2α) during ER stress that leads to global attenuation of mRNA translation which ward off further ingress of ER client proteins (Harding et al. 1999). Also, phosphoeIF2α (p-eIF2α) can preferentially induce activating transcription factor 4 (ATF4), a member of the ATF/CREB family of transcription factors (Mamady and Storey 2008). ATF4 is an important mediator of the UPR as it activates several pro-survival as well as pro-apoptotic genes. For example, ATF4 regulates several UPR target genes such as GADD34 (growth arrest and DNA damage inducible protein 34), a pro-survival protein that can reinitiate translation by interacting with protein phosphatase 1 (PP1) to dephosphorylate eIF2α (Mamady and Storey 2008). Among pro-apoptotic genes, ATF4 increases the expression of GADD153/C/EBP-homologous protein (CHOP) (Harding et al. 2000). Besides responding to ER stress caused by misfolding of proteins, GADD153 also responds to ROS, hypoxia, and nutrient deprivation (Krivoruchko and Storey 2013). Another gene downstream of ATF4 is the Activating Transcription Factor 3 (ATF3) that is involved in upregulation of genes involved in maintaining cell homeostasis (Harding et al. 2003).

Fig. 1.

Schematic diagram of UPR signaling and ERAD pathway proteins under ER stress. In an inactive state, IRE-1 and PERK are associated with Grp78. However, under ER stress, when the levels of misfolded/unfolded proteins rise in the ER lumen, Grp78 is competitively titrated by the excess of unfolded proteins, and this leads to activation of ATF6, PERK-1, and IRE-1 via oligomerization and autophosphorylation. Inactive ATF6 (90 kDa) translocates to the Golgi apparatus and becomes active (50 kDa), then translocates to the nucleus to guide adaptive transcriptional responses of downstream targets (Grp78, Grp94). Activated PERK stimulates eIF2α, and phosphorylates and induces ATF4, which leads to transcription of downstream gene targets (GADD153, GADD34). IRE-1 oligomerization catalyzes the splicing of XBP-1 mRNA to synthesize 50 kDa active XBP-1 protein that can induce expression of Grp chaperones (Grp78, Grp94). The action of Grps enhances the protein folding capacity of cells along with ERAD protein (EDEM) that induces proteasomal degradation of misfolded/unfolded proteins. Image created with www.BioRender.com

Upon phosphorylation, IRE-1 excises an unconventional 26-nucleotide intron from XBP-1 allowing translation of a longer sequence (376 amino acids) that produces the 40-kDa XBP-1 protein. XBP-1 is known to increase the transcription of ER chaperones (Grp78 and Grp94). Also, among different regulators of UPR signaling, the IRE1/XBP1 is mostly conserved and also regulates ERAD proteins such as EDEM (ER-degradation-enhancing-α-mannosidase-like protein) that increase cellular resistance to ER stress (Carrara et al. 2013; Mori 2009).

A third regulator of ER stress is another ER transmembrane transcription factor, ATF6 (Hiderou Yoshida et al. 1998). ATF6 shows crosstalk with components of both PERK/eIF2α/ATF4 and IRE-1/XBP-1 pathways (Senft and Ronai 2015) and can also upregulate the transcription of genes (such as EDEM) involved in ER protein folding (Cullinan and Diehl 2006). Signals are received in the cytosol that cause inactive 90-kDa ATF6 to undergo proteolytic cleavage in the Golgi apparatus to produce the active 50-kDa form (Fig. 1) (Shen and Prywes 2004). Active ATF6 then translocates to the nucleus and forms complexes with co-activators to trigger the transcription of ER chaperones (Yoshida et al. 2000).

The present study explores the UPR and ERAD regulated proteins in two tissues (liver and skeletal muscle) of wood frogs, under three stress conditions: freezing, anoxia, and dehydration. The proteins evaluated include ATF3, ATF4, ATF6, GADD34, GADD153, XBP1, and EDEM. The data suggest that both liver and muscle cells increase protein folding capacity in the ER to combat a rise in the unfolded protein load under environmental stress.

Methods

Animal preparation

Male wood frogs were collected from breeding ponds near Bishop’s Mills, Ontario, during early spring. In the laboratory, animals were washed in a tetracycline bath and then placed in plastic boxes with damp sphagnum moss at 5 °C for ~ 2 weeks before experimentation. Control frogs were sampled from these conditions. For freezing exposure, wood frogs were placed in closed plastic boxes lined with a moistened paper towel and were transferred to an incubator set at − 4 °C. The moistened paper towel quickly froze, allowing this ice to seed freezing of frog skin that, in turn, triggered ice propagation through the frog’s body. After 45 min, incubator temperature was raised to − 2.5 °C and frogs were held there for 24 h. The 24-h frozen group was then sampled. Remaining frogs in the frozen group were transferred to an incubator set at 5 °C for 8 h before sampling as the thawed group (Storey and Storey 1984).

For anoxia exposure, dechlorinated water was bubbled with 100% nitrogen gas for ~ 30 min. This water was then used to moisten paper towels that lined the bottom of the plastic chambers used for the experiment. The lids of the chambers had two ports: one to flush in nitrogen gas and the other to vent gas. These chambers were kept in a bucket containing crushed ice and were flushed with 100% nitrogen gas for 15–20 min. Then, 5 °C acclimated frogs were placed into these chambers (5–6 frogs/chamber) and lids were tightened and sealed with parafilm. Nitrogen gas was again flushed through the chambers for a further ~ 30 min and thereafter the ports were closed, and the closed chambers were replaced in 5 °C incubators. After 24 h of anoxia exposure, half of the chambers were returned one at a time to an ice bath. Flushing with nitrogen gas was restarted and frogs were quickly sampled with minimal air exposure as the 24 h anoxia-exposed group. Remaining anoxic frogs were transferred to new chambers with normal air and allowed to recover for 4 h at 5 °C before sampling. This was the 4 h aerobic recovery group.

For dehydration exposure, frogs were individually weighed and then groups of 5–6 frogs were put in tall plastic buckets (empty and without lids) and were replaced in a 5 °C incubator where they gradually lost body water by evaporation. Frogs were quickly reweighed at intervals and the percentage of body water lost was calculated using the following formula: (M_i − M_d)/(M_i × % H₂O) where M_i and M_d denote the initial mass and subsequent mass at each weighing, respectively, and the % H₂O is the percentage of body mass that is water in control frogs; this % H₂O value was 80.0 ± 1.2%. Under these experimental conditions, frogs lose ~ 0.5–1% of their body water/hour. The experiment continued until frogs lost ~ 40% of total body water, at which time half of the animals were sampled. The remaining frogs were transferred into buckets with ~ 0.5 cm water in the bottom, replaced at 5 °C and were allowed to rehydrate until they regained their initial mass (typically overnight). The fully rehydrated group was sampled from this condition.

All frogs were euthanized by pithing, followed by rapid dissection and flash freezing of liver and hind leg skeletal muscle in liquid nitrogen and storage at − 80 °C until use. All conditions for animal care, experiments, and euthanasia were pre-approved by the Carleton University Animal Care Committee (protocol #106,935) following guidelines set by the Canadian Council on Animal Care.

Preparation of tissue extracts for total protein measurements

Total soluble protein was extracted from frozen tissue samples of liver and hind leg skeletal muscle for each experimental condition, as previously described (Al-Attar and Storey 2018). Briefly, frozen tissue samples were quickly weighed (~ 50 mg/sample; n = 4 independent replicates from different animals) and homogenized (1:2 w/v) in a solution containing 20 mM HEPES buffer, pH 7.4, 100 mM NaCl, 0.1 mM EDTA, 10 mM NaF, 1 mM Na₃VO₄, 10 mM β‐glycerophosphate, a few crystals of phenylmethylsulfonyl fluoride (PMSF), and 1 μL of Sigma protease inhibitor cocktail (Burlington, ON, Canada, cat. no. P1C001.1) per milliliter of homogenization buffer using a Polytron homogenizer for 15–20 s. Homogenates were centrifuged at 12,000 × g for 15 min at 4 °C and supernatants containing total soluble proteins were collected. The Coomassie blue dye-binding method was used to measure soluble protein concentrations with the prepared reagent from Bio‐Rad (Mississauga, ON, Canada), and all concentrations were then standardized to 10 μg/μL by adding calculated small volumes of homogenizing buffer. Aliquots of total soluble protein extracts were then mixed 1:1 v/v with 2 × loading buffer containing 100 mM Tris–HCl, 4% w/v SDS, 20% v/v glycerol, 0.2% w/v bromophenol blue, and 10% v/v 2‐mercaptoethanol to give final concentrations of 5 μg/μL for liver and muscle samples. Samples were boiled for 5 min, cooled on ice for 5 min, and stored at − 80 °C until needed.

Preparation of nuclear protein extracts

Nuclear protein extracts were prepared from liver and muscle samples of animals for each of the three experimental conditions. Frozen tissue samples (~ 50 mg) were homogenized (1:5 w/v) in 1 × buffer A (10 mM HEPES, 10 mM KCl, 10 mM EDTA, 20 mM β-glycerol phosphate, pH 7.9) with 10 µL/mL of 100 mM dithiothreitol (DTT) and 10 µL/mL of protease inhibitor cocktail added immediately before homogenizing with a Dounce homogenizer using 3–4 gentle strokes. Samples were held on ice for 25 min and then centrifuged at 12,000 × g at 4 °C. Following centrifugation, supernatants were removed and saved as the cytoplasmic fraction. The pellet containing the nuclear fraction was re-extracted in 5 × lysis buffer B: 100 mM HEPES, 2 M NaCl, 5 mM EDTA, 50% v/v glycerol, 100 mM β-glycerol phosphate, pH 7.9 with 100 mM DTT and protease inhibitor (1:1000 v/v) added to the cocktail. An aliquot of 250 µL of 5 × buffer B was added to each pellet and the pellet was sonicated (Kontes micro-ultrasonic cell disruptor) for 5–10 s. Samples were incubated on ice for 10 min and then centrifuged for 10 min at 14,000 × g at 4 °C and the supernatant was saved. Protein concentrations were determined with the Bio-Rad protein assay and concentrations were adjusted to 2 μg/μL for both tissues. Samples were stored at − 80 °C until use. To evaluate the separation of cytoplasmic and nuclear fractions, immunoblotting was carried out as described below and tested with antibodies to α-tubulin (SC- 31779; Santa Cruz Biotechnology, Dallas, TX) and acetyl-histone H3 (CS-9649; Cell Signaling Technology) to assess any cross-contamination. The cytoplasmic fractions from liver and muscle showed a higher expression of α-tubulin protein as compared to the nuclear fractions. Conversely, histone protein levels were higher in the nuclear fractions from liver and muscle as compared with cytoplasmic fractions for both tissues (Tessier and Storey 2010).

Immunoblotting

To evaluate relative protein expression levels of selected targets in liver and muscle, proteins were separated on SDS-PAGE gels. Equal amounts of soluble protein were loaded into each well and gels were electrophoresed using a Bio-Rad Mini Protean III apparatus at 180 V in 1 × running buffer (3.2 g Tris base, 18.4 g glycine, 1 g SDS per liter) until the desired separation was achieved. Proteins were then electroblotted onto polyvinylidene difluoride (PVDF cat no. IPVH00010) membranes by wet transfer in prechilled 1 × transfer buffer (25 mM Tris pH 8.5, 192 mM glycine, 20% v/v methanol) at 4 °C for 70–90 min at 160 mA. After transfer, membranes were blocked with non-fat milk 2.5% in 1 × TBST (20 mM Tris base pH 7.6, 150 mM NaCl, 0.05% v/v Tween 20) for 30 min. Membranes were then washed before probing with specific primary antibodies (1:1000 v/v dilution in 0.5% TBST) overnight at 4 °C on a rocking platform.

Membranes were then washed for 3 × 15 min using 0.5% TBST, followed by incubation with secondary antibody for 2 h. Depending on the primary antibody, secondary antibodies were horseradish peroxidase (HRP)–linked anti-rabbit IgG secondary antibody, anti-goat IgG secondary antibody, or anti-mouse IgG secondary antibody (1:2000 v/v dilution). Following incubation, membranes were washed as described above before visualization using enhanced chemiluminescence (H₂O₂ and luminol). Blots were then stained using Coomassie blue (0.25% w/v Coomassie brilliant blue, 7.5% v/v acetic acid, 50% v/v methanol). The following antibodies from Santa Cruz Biotechnology (Dallas, TX) were used: ATF-3 (sc-188), ATF-4 (CREB-2) (sc-200), ATF-6 (sc-14253), GADD 34 (sc-794), GADD 153 (sc-7351), EDEM (sc-27391), and XBP-1 (inactive) (sc-32138). The antibody for XBP-1 (active) was a gift from Dr. L. Glimcher (Harvard School of Public Health, Boston, USA).

Statistical analysis

Bands on Western blots were visualized using a ChemiGenius Bio-Imager (Syngene, Frederick, MD) and band intensities were analyzed using the associated Gene Tools software. Band densities in each lane were standardized against the collective intensity of a group of Coomassie-stained protein bands in the same lane. The Coomassie-stained group was prominent, consistent across all lanes and well separated from the band of interest. This method is more accurate as compared to standardizing band intensities against a housekeeping protein such as GAPDH (Eaton et al. 2013). Band densities of experimental samples were then standardized relative to the corresponding control band densities. Immunoblot data are presented as mean ± SEM, n = 4 independent biological replicates. Statistical testing used the SigmaPlot 12.0 program to conduct a one-way ANOVA followed by a Student–Newman–Keuls test with p < 0.05 accepted as a significant difference. Statistical testing of cytoplasmic versus nuclear protein levels (mean ± SEM, n = 4 independent biological replicates) used the Student’s t-test, p < 0.05.

Results

ATF4 and downstream targets in liver and muscle during freezing, anoxia, and dehydration stress

Figure 2a and b show total protein levels of ATF4 and its downstream proteins (ATF3, GADD 34, and GADD 153) in liver and skeletal muscle comparing control conditions (5 °C acclimated) with responses to 24-h freezing at − 2.5 °C and subsequent 8-h thaw at 5 °C. ATF4 protein levels increased by 3.6-fold in liver of 24-h frozen frogs but after 8 h thawed recovery had returned to near control levels (Fig. 2a). Similarly, muscle ATF4 levels also increased significantly by 1.3-fold after 24-h freezing (Fig. 2b) but, unlike liver, were reduced to 50% compared to the control value after 8 h of thawed recovery. Downstream targets of ATF4 also showed strong responses in both tissues in response to freezing. ATF3 increased ~ 5-fold in liver and GADD34 levels rose 2-fold (Fig. 2a); both reverted close to control levels during the 8-h thawing recovery. By contrast, muscle ATF3 levels increased significantly during freezing (1.4-fold of control values) and remained high (1.5-fold over control values) during 8-h thawing and GADD 34 levels did not change across either freezing or thawing (Fig. 2b). By contrast, GADD153 levels in both liver and muscle were strongly suppressed to just 15 and 5% of control values, respectively, during freezing and remained low after 8-h thawing (25% in liver and 5% in muscle, as compared with controls) (Fig. 2a, b).

Fig. 2.

Effect of freezing, anoxia, and dehydration stresses on the relative expression levels of ATF4 pathway proteins (ATF4, ATF3, GADD34, and GADD153) in liver (a) and skeletal muscle (b) of wood frogs. Mean values for controls (at 5 °C) were set to 1.0 and experimental data are expressed relative to controls for freeze/thaw exposure (− 2.5 °C for 24 h, then 8-h thaw at 5 °C), anoxia/recovery (24-h anoxia, then 4-h aerobic recovery at 5 °C), and dehydration (40% dehydration, then full rehydration at 5 °C). Data are mean ± SEM, n = 4 independent biological replicates. Statistical testing used one-way ANOVA and Student–Newman–Keuls post hoc test with p < 0.05 accepted as significant. *Significantly different from controls

Unlike the response to freezing by liver, ATF4 protein levels in liver of anoxic frogs decreased significantly, falling to 48% of control values and remained at this level during 4-h aerobic recovery (Fig. 2a). However, muscle ATF4 levels showed a similar pattern as was seen during 24-h freezing with a significant increase (1.5-fold as compared to controls) when frogs were exposed to 24-h anoxia, but levels dropped to ~ 20% of control values after 4-h aerobic recovery (Fig. 2b). Liver ATF3 protein levels decreased significantly to 54% of control values after 24-h anoxia and remained at this level after aerobic recovery (Fig. 2a), whereas in muscle ATF3 levels rose by 3.5-fold over control values during 24-h anoxia and remained elevated (3.7-fold) and significantly higher than controls during aerobic recovery (Fig. 2b). GADD 34 levels increased in anoxic liver (by 1.4-fold) but returned to control values after aerobic recovery (Fig. 2a). Muscle levels of GADD 34 protein levels were also upregulated during 24-h anoxia exposure (by 2.2-fold) and decreased during recovery but still remained significantly elevated (1.5-fold) over controls (Fig. 2b). By contrast, GADD153 protein decreased significantly under anoxia to 43% of aerobic values and remained low after 4-h aerobic recovery (Fig. 2a), whereas in muscle GADD153 increased significantly during anoxia (1.3-fold) but fell to ~ 50% of control values during aerobic recovery (Fig. 2b).

Dehydration to 40% of total body water lost produced a different pattern of ATF4 response in liver as compared with freezing or anoxia. Liver ATF4 protein levels increased significantly by 2.4-fold above control values in response to 40% loss of body water and remained high (2.1-fold over controls) after rehydration (Fig. 2a), whereas in muscle levels of ATF4 were reduced to 31% of control values and remained low when animals were allowed to rehydrate fully (Fig. 2b). Interestingly, despite an increase in ATF4, the levels of ATF3, GADD34, and GADD153 remained unchanged during dehydration/rehydration stress in liver (Fig. 2a). However, unlike in liver, muscle ATF 3 levels were suppressed to about 30% of control values and remained low during recovery (~ 40% of controls) (Fig. 2b). Although dehydration stress did not affect GADD34 levels in muscle, during rehydration GADD34 levels increased by 1.4-fold over controls (Fig. 2b). Interestingly, GADD 153 content was reduced to 45% of control values during 40% dehydration but returned to control values during rehydration (Fig. 2b).

XBP1 and EDEM expression in liver and muscle under freezing, anoxia, or dehydration stress

The X-box binding protein (XBP1) exists in two forms. To detect both forms, two different antibodies were used that detected inactive (Inac XBP1, 30 kDa) and active (Act XBP1, 50 kDa) forms of XBP1 protein. During 24-h freezing, levels of Inac XBP1 in wood frog liver increased significantly by 1.5-fold over controls but returned to control values after thawing (Fig. 3a). Levels of the Act XBP1 also increased markedly during freezing (by 3.5-fold) and remained high after 8-h thawing (4.2-fold over controls) (Fig. 3a). EDEM protein levels in liver decreased significantly during freezing to 62% of control values and remained low after 8-h thawing (55% of controls) (Fig. 3a). In muscle, the levels of Inac XBP1 remain unchanged during freezing and subsequent 8-h thawing (Fig. 3b). By contrast, levels of Act XBP1 decreased significantly (to 40% of controls) during the 24-h freeze but returned to near control values after thawing (Fig. 3b). Levels of EDEM in muscle showed an opposite response to that of liver. Muscle EDEM increased by 3.2-fold during freezing and rose further to almost 6-fold higher than control values during 8-h thawing (Fig. 3b).

Fig. 3.

Effect of freezing, anoxia, and dehydration stresses on the relative expression of XBP1 pathway proteins (Inac XBP1, Act XBP1, EDEM) in a liver and b skeletal muscle of wood frogs. Data are mean ± SEM, n = 4 independent biological replicates. Statistical testing used one-way ANOVA and Student–Newman–Keuls post hoc test with p < 0.05 accepted as significant. *Significantly different from controls

Anoxia exposure led to a strong significant decrease in Inact XBP1 protein levels in liver (to ~ 45% of controls) during 24 h of anoxia exposure and remained low during aerobic recovery (Fig. 3a). However, Act XBP1 in liver responded similarly to that seen during freeze/thaw with significant increases in levels after both 24-h anoxia (1.65-fold) and 4-h recovery (2.8-fold) as compared with controls (Fig. 3a). Liver EDEM levels decreased significantly under anoxic conditions by 50% and remained low (43% of controls) during 4-h aerobic recovery (Fig. 3a). Muscle levels of the inactive form of XBP1 decreased significantly to ~ 32% of the control value after 24 h of anoxia but rose to ~ 66% of controls after 4-h aerobic recovery (Fig. 3b). By contrast, the amount Act XBP1 did not change under anoxia but declined to ~ 45% of controls during recovery (Fig. 3b). Muscle EDEM levels showed a similar pattern as in liver. EDEM levels declined during 24-h anoxia (to 44% of controls) and remained low at 37% of controls during recovery (Fig. 3b).

During dehydration stress, protein levels of inactive XBP1 rose significantly in response to 40% dehydration (by 1.6-fold) but returned to near control values after full rehydration (Fig. 3a). Act XBP1 showed a similar response increasing significantly by 2.6-fold during 40% dehydration but decreasing to 1.7-fold over controls after full rehydration (Fig. 3a). EDEM showed a very strong decrease in the liver during dehydration, levels being reduced to only 16% of controls but rebounding partially during rehydration to 43% of control values (Fig. 3a). Muscle showed opposite trends in response to dehydration as compared to liver. In muscle, Inac XBP1 levels were significantly reduced in both dehydration and rehydration (to 56% and 50% of controls, respectively) (Fig. 3b). Also, Act XBP1 levels decreased to 58 and 57% of controls, under dehydration and rehydration stress (Fig. 3b). However, EDEM levels remained unchanged during dehydration and rehydration in wood frog muscle (Fig. 3b).

ATF6 expression in liver and muscle during freezing, anoxia, and dehydration stress

Similar to XBP1, ATF6 exists in two forms: inactive (Inac ATF6, 90 kDa) and active (Act ATF6, 50 kDa). The relative protein levels of Inac ATF6 increased in the liver of wood frogs after 24-h freezing exposure and remained high after 8-h thawing (4.7- and 4.5-fold higher, respectively), as compared to controls (Fig. 4a). However, this did not correlate with a change in the active form of ATF6 protein; relative protein levels of Act ATF6 remained unchanged under all three conditions (control, 24-h freezing, and 8-h thawing) (Fig. 4a). In muscle, the levels of Inact ATF6 did not change during 24-h freezing or 8-h thawing (Fig. 4b). However, unlike liver, Act ATF6 levels rose significantly (1.6-fold) and remained elevated during thawing (1.45-fold over controls) (Fig. 4b).

Fig. 4.

Effect of freezing, anoxia, and dehydration stresses on the relative expression of ATF6 pathway proteins (Inac ATF6, Act ATF6) in a liver and b muscle of wood frogs. Data are mean ± SEM, n = 4 independent biological replicates. Statistical testing used one-way ANOVA and Student–Newman–Keuls post hoc test with p < 0.05 accepted as significant. *Significantly different from controls

Anoxia exposure also led to a significant increase in protein levels of inactive ATF6 in the liver (by 1.7-fold), as compared with controls, and remained at this level after 4-h aerobic recovery. Interestingly, Act ATF6 also increased under anoxia, rising significantly by 2.7-fold but returned to control values after 4-h aerobic recovery (Fig. 4a). In muscle, the protein levels of Inac ATF6 did not change during anoxia but rose significantly (1.7-fold as compared to controls) during 4-h recovery (Fig. 4b). Despite the rise in inactive levels of this protein during recovery, the levels of the active form did not change across the two conditions (Fig. 4b).

The levels of liver ATF6 during dehydration stress showed a similar response as in freezing. Dehydration stress led to a 1.5-fold increase in Inac ATF6 that was regained after rehydration (Fig. 4a). Despite the rise in Inac ATF6, the levels of Act ATF6 did not change across the two experimental conditions (Fig. 4a). By contrast, in dehydrated muscle the levels of Inac ATF6 did not change during 40% dehydration stress but rose (2.2-fold) during rehydration, as compared to controls (Fig. 4b). Similarly, the active form of this transcription factor was also unchanged during dehydration but increased by 1.3-fold, compared to controls, during rehydration (Fig. 4b).

Nuclear distribution of selected transcription factors in the UPR and ERAD responses to freezing, anoxia, and dehydration in liver and muscle of wood frogs

Since transcription factors (TFs) have dynamic behavior and their movement inside the nucleus, mostly under various stress/stimuli, can activate downstream signaling, we measured the nuclear levels of major TFs involved in ER stress signaling. Hence, it is important to gain information on the relative amounts of transcription factors in the nucleus under the various stress conditions. The data in Fig. 5a and b show the effects of freezing, anoxia, or dehydration on the nuclear content of ATF4, ATF3, ATF6 (active), and XBP1 (active) in wood frog liver and skeletal muscle, respectively. For muscle, data for a short time of freezing exposure (6 h) was also included since many transcription factors mediate changes in gene expression that would be expected to occur rapidly to prepare tissues long-term freezing survival. This would be especially true of leg skeletal muscle that freezes much earlier than do core organs like liver.

Fig. 5.

Relative nuclear protein expression of ATF4, ATF3, ATF6, and XBP1 in a liver and b skeletal muscle of wood frogs under control, 24-h freezing, 24-h anoxia, and 40% dehydration conditions. Data are mean ± SEM, n = 4 independent biological replicates. Statistical testing used one-way ANOVA and Student–Newman–Keuls post hoc test with p < 0.05 accepted as significant. *Significantly different from controls

ATF4

The distribution of ATF4 in nuclear fractions from liver and muscle is shown in Fig. 5a and b. Liver samples from both 24-h freezing and 40% dehydrated wood frogs showed upregulation of nuclear protein levels of ATF4 by 2-fold and 1.7-fold, respectively, as compared to controls, whereas 24-h anoxia exposure did not affect nuclear ATF4 content (Fig. 5a). In muscle, nuclear levels of ATF 4 did not change across 24-h freezing, 24-h anoxia, and 40% dehydration but only changed during early freezing (6 h) rising by 1.5-fold over control values (Fig. 5b).

ATF 3

In liver, nuclear ATF3 levels increased significantly during both freezing and anoxia exposures, by 1.7- and 2-fold, respectively (Fig. 5a). However, the nuclear content of ATF3 was unaltered under dehydration stress. In muscle, nuclear levels of ATF3 were unchanged after 6-h freezing but showed significant but small increases after 24-h freezing and 40% dehydration (1.2- and 1.4-fold, respectively) (Fig. 5b). However, 24-h anoxia triggered a very large 4.2-fold increase in nuclear ATF3 protein levels in muscle (Fig. 5b).

ATF 6

Liver ATF6 levels in the nuclear fraction increased under all three stress conditions. After 24-h freezing, nuclear ATF6 protein levels were elevated by 1.9-fold over control whereas increases were 1.5-fold under anoxia and 2.3-fold under 40% dehydration (Fig. 5a). In muscle, the nuclear ATF6 content increased significantly under all experimental conditions (Fig. 5b). Under 6-h and 24-h freezing, anoxia, and dehydration stresses, ATF6 nuclear levels increased significantly by 3.4-fold, 2.7-fold, 3-fold, and 4-fold respectively (Fig. 5b).

XBP1

There was no significant change in nuclear protein levels of XBP1 during 24-h freezing in liver (Fig. 5a). However, levels increased significantly by 1.7-fold and 2-fold in nuclear fractions of liver from anoxic and dehydrated frogs, respectively (Fig. 5a). In muscle, the nuclear levels of XBP1 significantly increased in samples from both 6-h and 24-h frozen frogs (by 1.6-fold and 1.8-fold, respectively) (Fig. 5b). However, neither anoxia nor dehydration affected muscle nuclear XBP1 levels.

Phosphorylation of eIF2α during freezing, anoxia, and dehydration stress in liver and muscle

The effects of environmental stress on the phosphorylation state of the eukaryotic initiation factor 2 alpha subunit (eIF2α) were also assessed in wood frog liver and muscle. The alpha subunit is considered to be the main regulatory subunit controlling translation initiation, and phosphorylation by a variety of protein kinases strongly inhibits eIF2α to halt global translation, sparing the preferential translation of ATF4. Figure 6a and b show the relative expression phospho-eIF2α in liver and muscle samples from frogs after 24-h freezing, 24-h anoxia, or 40% dehydration, compared with control values. In liver, freezing had no significant effect on p-eIF2α content, but both anoxia and dehydration exposures led to significant increases in p-eIF2α by 2.6- and 3.6-fold, respectively (Fig. 6a). By contrast, the response of p-eIF2α in muscle was opposite (Fig. 6b) with levels significantly increasing to 1.7-fold over controls during 24-h freezing, remaining unchanged during 24-h anoxia, and decreasing to close to ~ 50% of control values in response to 40% dehydration stress (Fig. 6b).

Fig. 6.

Relative protein expression phospho-eIF2α in a liver and b muscle in of wood frogs, comparing control, 24-h freezing, 24-h anoxia, and 40% dehydration conditions. Data are mean ± SEM, n = 4 independent biological replicates; statistical analysis used the Student’s t-test, *p < 0.05

Discussion

The endoplasmic reticulum responds to stresses that cause an increase in unfolded and misfolded proteins; such stress may be short term, such as environmental insults, or long term, such as metabolic diseases including type 2 diabetes and Alzheimer’s disease (Ozcan and Tabas 2012). The UPR and ERAD pathways are stress-responsive pathways that are activated in the ER upon accumulation of unfolded/misfolded proteins and are required to maintain ER homeostasis (Almanza et al. 2019). Freeze-tolerant vertebrates like wood frogs (R. sylvatica) are equipped with a natural tolerance to whole body freezing as well as the component stresses of freezing (dehydration and anoxia). All three of these stresses can individually cause ER stress for they can impose a reduction in metabolic function and energy restriction, large changes in cell volume and osmolality, and both anoxia and oxidative stress, the latter typically arising when O₂ levels rapidly return to normal after thawing (Storey and Storey 2013). Studies have shown that overall rates of new protein synthesis and protein modification/packaging are reduced as a part of freeze tolerance but selected proteins are upregulated that aid survival (Storey and Storey 2017). The presence of misfolded/unfolded proteins in the ER has been linked to the induction of the glucose-regulated ER chaperones, Grp78 and Grp94 (Zhu and Lee 2015; Kozutsumi et al. 1988), both of which are regulated in a tissue-specific manner in wood frogs (Storey and Storey 2019). Therefore, we were interested in determining how wood frogs deal with the potential disruption of ER function during freezing and/or its associated conditions (dehydration and anoxia) in order to establish the role of the UPR and ERAD pathways in establishing ER stability while frozen. In the current study, three main pathways of the UPR were studied in wood frog liver and skeletal muscle: (1) PERK/eIF2α/ATF4, (2) ATF6, and (3) IRE1/XBP1 (Fig. 1). The overlap between selected UPR and ERAD downstream targets was also explored in response to the three different stresses (freezing, anoxia, and dehydration).

UPR signaling and ERAD pathway in liver and muscle

PERK/eIF2α/ATF4 pathway in liver and muscle of wood frogs under stress

The overall response of the transcription factor, ATF4, was upregulation during 24-h freezing in the liver of wood frogs. This was evident from the rise in total protein levels during freezing (Fig. 2a) in conjunction with the rise in nuclear ATF4 content (Fig. 5a), suggesting that accumulation of this transcription factor is an early ER stress response to freezing. However, levels of the upstream activator of ATF4, p-eIF2α, did not change significantly during freezing, as compared to controls (Fig. 6a). Despite this, upon accumulation in the nucleus, ATF4 upregulation (Fig. 5a) was associated with increased levels of its downstream targets such as ATF3 whose total protein levels (Fig. 2a) and nuclear levels (Fig. 5a) were upregulated during 24-h freezing. ATF3 is known to repress the expression of GADD 153 (Fig. 2a) that is a known player in ER stress-mediated apoptosis (Wolfgang et al. 1997). The low levels of GADD 153 during freeze/thaw (Fig. 2a) could also be a result of elevated GADD 34 during freeze/thaw in liver (Fig. 2a) since GADD 34 has been shown to reduce GADD 153 content in liver (Inaba et al. 2015). This could further help to keep apoptotic regulators at bay while frogs are frozen (Novoa et al. 2001). Interestingly, both ATF3 and GADD34 also play a crucial role in regulating cell’s metabolism (Ku and Cheng 2020; Nishio and Isobe 2015). ATF3 plays a role in suppressing gluconeogenesis via downregulating a key enzyme PEPCK (phosphoenolpyruvate carboxykinase) during stress (Ku and Cheng 2020). The downregulation of PEPCK in the liver of freezing wood frog in another study (Kiss et al. 2011) aligns with the observed downregulation of ATF3 (Figs. 2a and 5a). Also, GADD34 has been shown to suppress mTORC (mammalian target of rapamycin) during stress that can promote global protein translation and is one of the key players in regulating cellular homeostasis during stress (Holczer et al. 2016). Also, mTORC has been shown to be downregulated in various naturally stress-tolerant animals (hibernators such as thirteen-lined ground squirrels and reptiles such as red-eared slider turtles) (Logan et al. 2019; Szereszewski and Storey 2018), hinting at the possible role of GADD34 in maintaining the low levels of mTORC to conserve the available energy and halt energy-expensive pathways such as protein synthesis. Therefore, both ATF3 and GADD34 could enhance the capacity of the liver to remain metabolically active over freeze/thaw to fulfill major survival needs (e.g., cryoprotectant production and distribution), utilizing energy sources judiciously and minimizing oxidative stress during thawing (reperfusion injury) by abating the apoptotic response (Storey and Storey 2017).

Interestingly, the overall response of ATF4 and downstream genes under its control in skeletal muscle responding to freezing (Fig. 2b) was similar to that seen in liver with upregulation of major players involved in abating an apoptotic response. Total protein levels of ATF4 were significantly higher than controls during freezing (Fig. 2b) in conjunction with a rise in p-eIF2α levels in skeletal muscle (Fig. 6b), suggesting preferential selection of p-eIF2α for ATF4 activation during 24-h freezing in wood frog muscle. Interestingly, nuclear levels of ATF4 only rose during the first 6 h of freezing (by ~ 1.5-fold) (Fig. 5b) but had returned to control values after 24-h freezing. This observation suggests a crucial role for active ATF4 during early freezing in peripheral tissues such as skeletal muscle that, together with skin, are the first tissues to freeze (Rubinsky et al. 1994). This early freezing response could underlie an overexpression of downstream ATF3 total protein and nuclear levels (Figs. 2b and 5b, respectively) during 24-h freezing. ATF3 total protein levels remained high even during thawing (Fig. 2b) although GADD 34 levels did not change but strong upregulation of ATF3 might be enough to mitigate expression of the apoptotic regulator GADD 153 (Wolfgang et al. 1997) during both 24-h freezing and 8-h thawing (Fig. 2b). This response was similar to that seen in liver during freezing and may be one of the wood frog’s defense strategies to maintain the metabolic integrity of muscle tissue during freeze/thaw. Hence, overall, both liver and muscle showed upregulation of key proteins involved in minimizing apoptotic damage during freeze/thaw, and this also supports previous studies that showed heightened anti-apoptotic or antioxidant machinery during cycles of freeze/thaw in wood frogs (Zhang et al. 2021; Zhang and Storey 2013).

Under anoxic conditions, the total protein levels of ATF4, GADD 153, and ATF3 all decreased, and these remained low during the 4-h aerobic recovery suggesting a reduced response by the ATF4 pathway or lower ER stress occurring under anoxia/recovery conditions in the liver. Interestingly, levels of p-eIF2α increased significantly during 24-h anoxia that seems to disagree with decrease in total protein and nuclear levels of ATF4 (Figs. 2a and 6a) and the increase in GADD34 levels (Fig. 2a). Despite this, the nuclear levels of ATF3 were upregulated during 24-h anoxia (Fig. 5a) and might keep apoptotic damage due to GADD 153 action under check by lowering the expression of this protein in anoxia. Alternate regulation of ATF3 under anoxic conditions has been shown in another study that can regulate the cell cycle via suppressing cyclin D1 gene expression (Ameri et al. 2007). Also, a separate study of wood frog liver exposed to 24-h anoxia showed suppressed levels of cyclin D1 (Zhang and Storey 2012). This also suggests other possible roles and multiple levels of regulation of this major transcription factor in order to maintain homeostasis under stress conditions. Overall, regulation of downstream targets was similar to frozen liver, but marked differences in ATF3 and ATF4 could be attributed to anoxic liver not undergoing cryoprotectant synthesis or disruptive blood flow compared to freezing liver (Al-Attar and Storey 2018).

Compared to liver, the response of ATF4 pathway proteins in muscle was the opposite of that seen in anoxic liver. The total protein levels of ATF4 rose significantly under 24-h anoxia (Fig. 2b) despite a lack of change in p-eIF2α levels (Fig. 6b). The nuclear levels of ATF4 stayed the same during anoxia exposure (Fig. 4a). Despite this, ATF3 levels were elevated, for both total protein (Fig. 2b) and nuclear protein (Fig. 5b). This argues for a separate regulatory mechanism independent of ATF4. One possibility could be the same as in anoxic liver where inhibition of cell cycle activity via suppressing cyclin D levels has been demonstrated in anoxic muscle of wood frogs (Roufayel et al. 2011). Interestingly, levels of GADD153 were also high during 24-h anoxia (Fig. 2b), despite higher nuclear and total levels of ATF3 and total levels of GADD34 (Fig. 2b). The increased levels of GADD 153 hints at an apoptotic route via overexpression of the pro-apoptotic protein, Bax (BCL2 Associated X, Apoptosis Regulator) (Oyadomari and Mori 2004), that has also been shown to upregulate in a separate study in anoxic wood frog muscle (Gerber et al. 2016). The same study (Gerber et al. 2016) also showed downregulation of another pro-apoptotic factor such as phosphorylated P-53 (p-P53 at ser 46) during 24-h anoxia and levels of anti-apoptotic factors like Bcl-2 increased during 4-h aerobic recovery during which wood frogs face reperfusion-induced ROS damage and need a heightened anti-apoptotic and protein refolding response as shown in some of the wood frog studies (Gerber et al. 2016; Storey and Storey 2019). Furthermore, elevated levels of ATF3 and GADD34 (Fig. 2b) during 4-h aerobic recovery could act in conjunction to reduce GADD153 levels (Fig. 2b) below control levels to keep a check on apoptotic cascade route.

Under dehydration stress, ATF4 levels rose significantly in liver, a response that was similar to that seen during freezing (Fig. 2a). This occurred in conjunction with an increase in liver p-eIF2α content (Fig. 6a) that could activate ATF4 expression and suggests a more active response by ATF4 under dehydration, as compared to anoxia conditions. This was further supported by the active nuclear levels of ATF4 (Fig. 4a) that also rose significantly in response to dehydration. Interestingly, however, the typical downstream effects of ATF4 upregulation (i.e., enhanced levels of GADD34, GADD153, and ATF3) were not seen during dehydration or rehydration (Figs. 2a and 5a). This suggests that liver ATF4 may have other downstream targets under dehydration/rehydration conditions (Tsuru et al. 2016) or that dehydration may have lesser consequences for ER action in liver, possibly since core organs may be less affected by dehydration as compared to peripheral tissues (e.g., skin, muscle). This is different from the effect of freezing since ice growth throughout the body affects the hydration state of all organs.

The results for skeletal muscle under dehydration stress were different from freezing or anoxia. ATF4 protein levels were downregulated in 40% dehydrated frogs and remained low after rehydration (Fig. 2b). Interestingly, muscle dehydration was the only stress/tissue combination to show reduced expression of p-eIF2α (Fig. 6b) which could be a possible reason for the lower ATF4 expression observed (Fig. 2a), and for the increased expression of GADD 34, the latter being known induce protein phosphatase 1 (PP1) that can dephosphorylate eIF2α. Also, levels of ATF3 remained low during both dehydration and rehydration, but GADD 153 declined during both conditions (Fig. 2b). One possible mechanism for a lower expression of GADD153 could be the rise in ATF3 nuclear levels (Fig. 5b) (Wolfgang et al. 1997). Furthermore, the reduced levels of most of the major protein components of the PERK pathway in response to dehydration hinted at little or no ER stress in muscle when water content was reduced.

IRE1/XBP1 pathway in liver and muscle during freezing, anoxia, and dehydration stress

The levels of inactive XBP1 (30 kDa) and spliced active protein (50 kDa) were analyzed. In liver, both inactive XBP1 (Inac XBP1) and active XBP1 (Act XBP1) rose during 24-h freezing and Act XBP1 also remained high after 8-h thawing (Fig. 3a). The upregulation of active XBP1 might be explained by a known action of XBP1 in regulating Grp expression. Furthermore, ATF6 can bind to the XBP-1 promoter and induce expression of other genes such as EDEM (Chen et al. 2016). Although no change in the nuclear levels of XBP1 occurred (Fig. 5a), EDEM levels were downregulated after both 24-h freezing and 8-h thawing (Fig. 3a), suggesting that the liver may use an alternate pathway beside ERAD to dispose of excess and misfolded proteins or, alternately, that liver has a low buildup of misfolded proteins during freezing and might instead promote refolding of proteins during freezing via molecular chaperones (Storey and Storey 2019; Vabulas et al. 2010). This also could be seen in PERK/ATF4 pathway which also links to a state of low apoptotic damage to the proteins during freezing. In muscle during freezing, the inactive form XBP1 (Inac XBP1) did not change and active XBP1 (Act XBP1) levels were reduced during 24-h freezing (Fig. 3b). However, although active XBP1 total protein levels remained low during freezing, nuclear levels were elevated during early freezing (6 h) and increased further during 24-h freezing (Fig. 5b). This suggests an important role for XBP1 from early freezing (Fig. 5b) through to full 24-h freezing, thereby timely inducing levels of EDEM that could act to remove misfolded or excess proteins via proteasomal degradation during freezing and/or thawing (Fig. 3b).

Anoxia stress also triggered elevation of Act XBP1 (Fig. 3a) in liver as well as upregulation of nuclear levels of XBP1 (Fig. 5a). Despite this, the XBP1 downstream target, EDEM, was downregulated in both 24-h anoxia and 4-h aerobic recovery suggesting a possibility of ER stress in anoxic liver but not likely building up misfolded protein in excess. Furthermore, the upregulated nuclear levels of XBP1 during anoxia and dehydration could also lead to elevation of GRPs (Grp78 and Grp94) as shown in a separate wood frog study and that may support more efficient protein folding as well as production and secretion of novel freeze responsive proteins (J. M. Storey and Storey 2019). In muscle, the regulation XBP1 (active and inactive) and EDEM were either unchanged or downregulated (Fig. 3b). Overall, low expression of major proteins involved in ER stress and ERAD, along with unchanged nuclear levels of XBP1 (Fig. 5b), suggests a downplay of this pathway possibly due to lower or no ER stress during anoxia/recovery in wood frogs.

The dehydrated liver showed a similar protein expression pattern as in freezing (Fig. 3a), although the upregulation of Inac XBP1 and Act XBP1 (Fig. 3a) and a rise in nuclear levels of XBP1 (Fig. 5a) did not lead to upregulation of EDEM levels during dehydration/rehydration. Actually, EDEM levels declined under both conditions (Fig. 3a). This again suggests that either liver under dehydration stress uses an alternate path to ERAD to remove excess or misfolded proteins (Tsuru et al. 2016) or has a low buildup of misfolded proteins during dehydration. In muscle, the levels of both Inac XBP1 and Act XBP1 during dehydration and rehydration remained low (Fig. 3b). In particular, Act XBP1 levels fell during both 40% dehydration (42% of control values) and 24-h freezing (60% of control) (Fig. 3b) compared to control values. This hints that XBP-1 control in muscle might be linked with cell hydration. Interestingly, with the lower expression of Act XBP1, neither nuclear levels of XBP1 (Fig. 5b) nor EDEM levels (Fig. 3b) changed significantly under dehydration stress hinting that little or damage occurs to the proteome under these conditions.

ATF6 pathway in liver and muscle during freezing, anoxia, and dehydration stress

A third signaling molecule involved in the UPR is the transcription factor, ATF6, that in its inactive form (90 kDa, Inact ATF6) is present in the cytoplasm but is converted to an active form (50 kDa, Act ATF6) under ER stress causing Act ATF6 to translocate to the nucleus to induce downstream genes common to the XB1 pathway (Mamady and Storey 2008). In liver, Inac ATF6 increased significantly by over 4-fold during 24-h freezing and 8-h thawing, although Act ATF6 remained unchanged (Fig. 4a). Despite this, the nuclear levels of Act ATF6 rose rapidly (Fig. 5a). As mentioned previously, active ATF-6 can also induce XBP-1, Grp78, and Grp94 gene expression once inside the nucleus (Fig. 5a) (Yoshida et al. 2000; Shuda et al. 2003). The upregulation active ATF6 (formed as a result from proteolytic cleavage due to accumulation of misfolded protein) (Fig. 5a) does hint at possible ER stress but with less preference to send misfolded protein for degradation via ERAD pathway. This was evident from downregulation of EDEM (Fig. 4a) which suggests that proteasomal degradation of proteins is lowered and focus on protein refolding capacity might be higher. Also, a previous study of wood frogs showed an upregulation of Grp78 and Grp94 under stress conditions (Storey and Storey 2019). Unlike in liver, in skeletal muscle Inac ATF6 total protein levels remained unchanged while Act ATF6 levels changed during both 24-h freezing and 8-h thawing (Fig. 4b). Interestingly, levels of ATF6 in the nucleus rose during the first 6 h of freezing stress and remained significantly higher than controls (Fig. 5b). This suggests that upregulated ATF6 signaling could work by regulating EDEM protein levels in the ERAD pathway. The ERAD pathway is activated to move unfolded and misfolded proteins back to the cytosol where they can be degraded via proteasome degradation (Oslowski and Urano 2011). High levels of the main protein EDEM during both freezing (4-fold) and recovery (8-fold) (Fig. 3b), in conjunction with active ATF6 levels, suggests that during freezing, the increased amounts of misfolded and unfolded proteins could be targeted via this route for proteasomal degradation and reduce the burden on the cell’s apoptotic machinery. These results also align with a study from our laboratory that showed a higher amount of protein oxidative damage in muscle from 24-h frozen frogs with lesser activity of the multicatalytic proteasome (MCP) suggesting that significant protein damage occurred during freezing in skeletal muscle but recovered during 8-h thawing (increase in MCP activity) (Woods and Storey 2006) which also matches our current results (EDEM levels rose 8-fold during thawing; Fig. 3b). This suggests that the frozen ischemic state may tolerate protein damage but after thawing, protein degradation pathways go into high gear to clear misfolded or damaged proteins (Woods and Storey 2006).

The responses to both anoxia and dehydration stresses were similar in liver of the wood frogs (Fig. 4a). Total protein levels of Inac ATF6 rose under both anoxia and dehydration stress and remain elevated during recovery, whereas Act ATF6 changed only during 24-h anoxia (Fig. 4a). Despite this, nuclear levels of ATF6 were higher across both stresses: a 1.9-fold increase under anoxia and 2.3-fold under dehydration (Fig. 5a). This overall upregulation of active ATF6 in the liver nucleus under anoxia and dehydration stresses (compared to freezing in liver) indicates a downplay of proteasomal degradation via the ERAD pathway protein, EDEM, under anoxia or dehydration (Fig. 3a) with a probable focus again on supporting the protein refolding capacity via upregulation of molecular chaperones (Storey and Storey 2019). In addition to this action under UPR signaling, ATF6 has also been shown to bind to cofactors such as the CREB Regulated Transcription Coactivator 2 (CRTC2) that binds to the gluconeogenic transcription factor CREB (cAMP response element binding protein) and thereby inhibits gluconeogenesis (Rutkowski 2019). Since under freezing and dehydration conditions (but not in anoxia), a large amount of glucose (~ 200 mM) is produced via glycogenolysis in the liver and gluconeogenesis is inhibited, there is a possibility that ATF6 might also be contributing to maintaining cryoprotectant levels under these conditions or regulating the levels of molecular chaperones (e.g., Grp94) to inhibit apoptosis (Al-attar et al. 2017).

Under anoxia and dehydration stress conditions in muscle, the overall nuclear response of the ATF6 pathway (Fig. 5b), again hinted that upregulated nuclear levels of this transcription factor could aid in regulating protein folding via inducing Grp94, which has been shown to increase during dehydration/rehydration stress in wood frogs (Storey and Storey 2019).

Conclusion

UPR signaling was triggered under all three stresses (freezing, anoxia, dehydration) in liver and skeletal muscle of wood frogs with most tissue/stress responses consistent with an upregulation of the primary targets of all three UPR pathways (ATF4, ATF6, and XBP-1) to enhance the protein folding/refolding capacity under these stress conditions. Interestingly, only frozen muscle showed an acute rise in an ERAD protein (EDEM) that could also be linked with a rise in nuclear levels of activators (ATF4, ATF6, and XBP1) after 6-h freezing exposure. The rise in EDEM in freezing muscle hints at proteasomal degradation of misfolded proteins and reduce the burden on the cell’s apoptotic machinery. Apart from freezing, anoxic muscle showed little sign of apoptosis via upregulation of GADD153, but corresponding anti-apoptotic proteins were also upregulated and these could potentially hold apoptosis to a minimum. Both anoxia and dehydration led to upregulation of key proteins (ATF6, XBP-1) even during the recovery period, hinting that these transcription factors might also have roles in addressing the challenges of restoring homeostasis during recover from stress.

The liver UPR and ERAD responses were similar to muscle with a difference of downregulation or no change of EDEM (ERAD) protein across three stresses, suggesting that the overall response to ER stress is to utilize the available energy to maintain cellular homeostasis by upregulating molecular chaperone response like Grp78 and Grp98 via ATF4-ATF-6-XBP-1 regulation and not be going through acute apoptotic stress (also evident from downregulation of GADD153). This further suggests that in liver, the focus would be on more refolding of misfolded/unfolded proteins since liver is the most active organ involved in cryoprotectant synthesis and upregulation of cell protective and survival pathways.

Funding

Research was supported by a Discovery grant to KBS from the Natural Sciences and Engineering Research Council of Canada (#6793); KBS holds the Canada Research Chair in Molecular Physiology.

Declarations

Conflict of interest

The authors declare no competing interests.