Biogenesis and Regulation of the Freeze–Thaw Responsive microRNA Fingerprint in Hepatic Tissue of Rana sylvatica

Abstract

Background: Freeze-tolerant animals undergo significant physiological and biochemical changes to overcome challenges associated with prolonged whole-body freezing. In wood frog Rana sylvatica (now Lithobates sylvaticus), up to 65% of total body water freezes in extracellular ice masses and, during this state of suspended animation, it is completely immobile and displays no detectable brain, heart, or respirometry activity. To survive such extensive freezing, frogs integrate various regulatory mechanisms to ensure quick and smooth transitions into or out of this hypometabolic state. One such rapid and reversible regulatory molecule capable of coordinating many aspects of biological functions is microRNA. Herein, we present a large-scale analysis of the biogenesis and regulation of microRNAs in wood frog liver over the course of a freeze–thaw cycle (control, 24 h frozen, and 8 h thawed). Methods/Results: Immunoblotting of key microRNA biogenesis factors showed an upregulation and enhancement of microRNA processing capacity during freezing and thawing. This was followed with RT-qPCR analysis of 109 microRNA species, of which 20 were significantly differentially expressed during freezing and thawing, with the majority being upregulated. Downstream bioinformatics analysis of miRNA/mRNA targeting coupled with in silico protein–protein interactions and functional clustering of biological processes suggested that these microRNAs were suppressing pro-growth functions, including DNA replication, mRNA processing and splicing, protein translation and turnover, and carbohydrate metabolism. Conclusions: Our findings suggest that this enhanced miRNA maturation capacity might be one key factor in the vital hepatic miRNA-mediated suppression of energy-expensive processes needed for long-term survival in a frozen state.

1. Introduction

A key theme employed in various winter survival strategies, including hibernation, freeze-avoidance, and freeze-tolerance, is the use of hypometabolism [1]. Depressing the metabolic rate to less than 30% of active levels allows animals to survive prolonged periods of dormancy using only internal fuel reserves while preventing the buildup of deleterious waste byproducts. An excellent model of natural vertebrate freeze-tolerance is the North American wood frog Rana sylvatica (now Lithobates sylvaticus). This animal ranges from the Southern Appalachian mountains across the northern boreal forests of Canada and into Alaska, and due to subzero temperatures experienced in these climes during winter, can retreat into a frozen state of suspended animation for months at a time and endure the freezing of up to 65% of total body water as extracellular ice masses [2]. When frozen, wood frogs display no detectable brain activity, no breathing, no movement, and a flat-lined heart, and yet, when temperatures warm, they are able to thaw and return unscathed to normal life. This requires extensive molecular reorganization to reprioritize fuel/energy use to processes necessary for vital cellular functions and pro-survival mechanisms [3]. Adaptations for successful freeze-tolerance include the following: (1) minimization of cell volume reduction and excessive extracellular ice formation by synthesizing and distributing massive amounts of glucose that act as an intracellular cryoprotectant [4]; (2) global metabolic rate depression (MRD) and reprioritization of ATP usage; and (3) upregulation and activation of selected pro-survival and protective mechanisms [3]. Indeed, despite being in a hypometabolic state, wood frogs must expend energy on protective functions, such as antioxidant defenses [5], to ensure that their cells can endure the myriad of freeze-associated challenges that include mechanical damage from ice, dehydration, anoxia/ischemia, hyperglycemia, etc. [3].

Numerous molecular mechanisms are in place to orchestrate smooth transitions between active and frozen dormant states. These include regulatory mechanisms of MRD that are conserved in other hypometabolic systems, including epigenetic DNA and histone modifications, activity of signal transduction pathways, transcription factor regulation, post-translational protein modifications, and post-transcriptional regulation of gene transcripts by microRNA (miRNA) [6]. MiRNAs are a class of multifunctional molecules that are now known to be central modulators of diverse biological functions and environmental stress responses due to their ability to modulate mRNA translation [7]. These highly conserved small non-coding RNA transcripts (~22 nt) can selectively fine-tune gene expression by binding to protein-coding gene transcripts (in humans, over 60% of protein-coding genes are miRNA-regulated) to either promote their degradation or induce translational suppression [8,9,10]. MiRNA-mediated gene silencing is a prime regulatory candidate for reversible metabolic reorganization to facilitate both global MRD and selective gene activation. The inherent properties that make miRNAs excellent regulators include their capacity to be (1) easily inducible, (2) rapidly activated, (3) readily reversible, (4) energetically inexpensive, and (5) responsive to environmental stimuli [6]. This regulatory capacity is further enhanced in complexity and flexibility by the ability of a single miRNA type to target the mRNA transcripts of hundreds of different genes, and, conversely, that transcripts of any given gene can be targeted by hundreds of miRNAs [11,12].

MicroRNA biogenesis is an evolutionarily conserved stepwise process that has recently been shown to be under tight spatial and temporal control [9]. Both animals and plants utilize canonical and non-canonical versions of miRNA biogenesis, and the conservation of miRNAs allows cross-kingdom uptake of these small RNA species despite evolutionary differences between their respective biogenesis pathways [13,14]. The canonical biogenesis pathway in animals begins with the transcription of long primary-miRNA (pri-miRNA) by RNA polymerase II. Pri-miRNA is then cleaved into a ~70 nt long RNA hairpin structure known as a precursor-miRNA (pre-miRNA) by the RNase III endonuclease DROSHA and its partner DGCR8. Current processing models suggest that DGCR8 directs DROSHA to a specific cleavage site by recognizing the pri-miRNA ssRNA–dsRNA junction [15]. Pre-miRNA is then exported from the nucleus to the cytoplasm by the nuclear transporter EXPORTIN-5 (XPO5) and RAN-GTP shuttle that also protects the newly formed pre-miRNA from nuclear degradation [16]. Then, DICER, another RNase III endonuclease enzyme, cleaves the pre-miRNA transcript into a miRNA duplex composed of both a passenger strand (which is generally degraded) and a guide strand that is cleaved into the mature miRNA [17]. This is facilitated by the transactivator response RNA binding protein (TRBP) that complexes with DICER to monitor the length of mature miRNAs in addition to modulating the processing efficiency of select pre-miRNAs [18,19]. A third protein also present in this complex is the protein kinase RNA activator (PACT); its role in miRNA maturation is to promote DICER-mediated cleavage of the pre-miRNA [19,20]. The mature miRNA strand then associates with the multi-component RNA-induced silencing complex (RISC) and a member of the Argonaute family of endonucleases (AGO 1–4) that then binds to specific regions in the 3′ untranslated region (3′-UTR) of target mRNA transcripts. Imperfect miRNA/mRNA binding leads to translational suppression via sequestration into stress granules or processing bodies (P-bodies), whereas perfect binding results in mRNA target degradation [12].

Since this study examines miRNAs in a freeze-tolerant model, the influence of temperature on miRNA functionality must also be investigated. While the binding of miRNA to mRNA targets is mainly dictated by the degree of complementarity at the seed region, it should be noted that the 3′ end of miRNAs have been shown to bind in a manner that stabilizes the miRNA/mRNA duplex [21]. A thermodynamic threshold of −18 kcal/mol is generally used to predict miRNA/mRNA target binding, and this suggests that large fluctuations in body temperature will significantly affect miRNA targeting [21,22]. Indeed, at low temperatures, more miRNA/mRNA interactions will be stabilized, which in turn increases the dynamic regulatory potential of these molecules. This has been demonstrated in a pilot study on freeze-tolerant turtles that reported significant changes and enhanced miRNA functionality and targeting in frozen states [23], and the concept was further expanded by a large-scale analysis of cold-specific miRNA targeting in frozen wood frog brains [24]. An additional temperature-dependent effect concerns the thermodynamic stability of processed miRNA duplexes, in which even small shifts in temperature have been shown to modify strand-specific expression of miRNAs [25,26]. Taken together, the large number of miRNA species, their complementary mRNA targets, and the temperature-dependent effects on binding all act to generate an expansive regulatory RNA network that enables broad and flexible control over mRNA expression during stress.

As demonstrated by proton MRI, one of the last organs to succumb to ice accumulation and the first to thaw in wood frogs is the liver [27]. The liver is the metabolic center responsible for coordinating whole-body metabolism, encompassing functions including the following: ketogenesis, gluconeogenesis, regulation of blood sugar levels, carbohydrate storage, detoxification, protein synthesis and export, and the processing and storage of nutrients. In addition to these vital processes, the liver takes on additional roles during freezing, such as the synthesis and distribution of cryoprotectant glucose and freeze-responsive protein synthesis. This includes the enhanced synthesis of the blood clotting protein fibrinogen and novel freeze-responsive proteins FR-10 and FR-47 that function to ensure whole organism cryoprotection and freeze survival [28,29,30]. Within minutes of the initial ice nucleation event, the liver begins to convert its massive summer-accumulated glycogen stores into glucose through the activation of glycogen phosphorylase, among other enzymes [3]. Glucose is then exported from the liver and distributed to all other body tissues where high sugar concentrations act to protect against excessive cellular dehydration and minimize intracellular and extracellular ice formation [3]. While liver metabolic reorganization has been shown to be regulated by hormones, transcription factor networks, and enzymatic controls [3,31], recent work is now focusing on elucidating the role of the post-transcriptional regulation of gene expression in frozen wood frog livers. Initial studies of miRNA expression in wood frog liver identified five miRNA species (miR-26a, miR-126, miR-217, miR-21, and miR-16) that were freeze-upregulated; these were linked to anti-apoptotic functions and cell-cycle suppression [32,33]. Another study assessed miRNA expression in wood frog skeletal and cardiac muscles and reported tissue-specific differential expression of 53 miRNAs over the freeze–thaw cycle; target prediction suggested their involvement in maintaining muscle contraction and reversible protein phosphorylation [34]. A small RNA-sequencing study of related freeze-tolerant frog species, Dryophytes versicolor (formerly Hyla versicolor), identified 11 miRNAs, which were differentially regulated in the liver during freezing [35]. A study of over 100 miRNAs and their biogenesis in wood frog brains identified a global trend of reduced miRNA biogenesis capacity coupled with freeze-downregulated and thaw-downregulated miRNAs that were responsible for neuroprotective functions during hypometabolic exposures [24].

In this study, we will implement a large-scale analysis of miRNA biogenesis and regulation in wood frog livers over freeze–thaw exposure. Immunoblotting of the key components of miRNA biogenesis machinery will characterize the behavior of this pathway during freeze–thaw cycles, and RT-qPCR of 109 pre-selected miRNAs will reveal differential regulation (if any) of these small RNAs during freeze–thaw and their downstream functional roles. The pre-selected miRNAs were determined from the following sources: 8 miRNAs from a family of cold-associated “cryomiRs” [36], 23 miRNAs from a study in cardiac/skeletal muscle of R. sylvatica over the freeze–thaw cycle that share hepatic-functional roles [34], and 78 miRNAs from a study in the brain tissue of R. sylvatica over the freeze–thaw cycle [24]. Of these miRNAs, 53 in total were differentially regulated in other cold-tolerant species and/or tissues of R. sylvatica, including 6 in cardiac tissue, 6 in skeletal muscle, 3 in both cardiac and skeletal muscle, and 40 in the brain [24,34]. Further, in silico miRNA target prediction and functional enrichment at −2.5 °C and 5 °C will be performed to reveal potential alterations in miRNA/mRNA binding at the physiologically relevant temperatures wood frogs experience during freezing. Taken together, our findings will shed light on the upstream and downstream components of liver miRNAs during the freeze–thaw cycle of R. sylvatica and how they may underpin the survival of this animal during the winter months.

2. Materials and Methods

2.1. Animal Experiments

Male wood frogs were collected from spring meltwater ponds near Oxford Mills, Ontario, Canada. Frogs were washed in a tetracycline bath and then acclimated for two weeks at 5 °C in plastic containers with a layer of damp sphagnum moss (frogs were not fed). Active (non-stressed) control frogs (n = 4–5) were sampled from this condition. For the 24 h frozen condition, frogs (n = 8–9) were moved into plastic containers lined with a damp paper towel and moved to an incubator set at −4 °C for 45 min to cool the frogs and trigger ice nucleation. After this time, the temperature was raised to –2.5 °C, and a 24 h freezing exposure was timed from that point. Frozen frogs (n = 4–5) were randomly sampled from this condition. Remaining frogs were assigned to the recovery group and transferred to 5 °C, where they thawed for 8 h. All control, 24 h frozen, and 8 h thawed frogs were euthanized by double-pithing. The liver was rapidly excised, flash-frozen in liquid nitrogen, and stored at −80 °C. Animal care and experimentation protocols had the prior approval of the Carleton University Animal Care Committee (protocol #106935; valid 11 June 2017 to 11 June 2020) and followed guidelines set by the Canadian Council on Animal Care (protocol #13683). Frogs were collected under permit #1085726 (valid 13 March 2017 to 31 December 2017) issued by the Ontario Ministry of Natural Resources.

2.2. Total Soluble Protein Extraction

Total soluble protein extracts were prepared from the liver of control, 24 h frozen, 8 h thawed wood frogs. Samples (~500 mg) of frozen tissue were homogenized 1:2 w/v using a Polytron PT10 homogenizer and chilled homogenization buffer (20 mM HEPES, 200 mM NaCl, 0.1 mM EDTA, 10 mM NaF, 1 mM Na₃VO₄, 10 mM β-glycerophosphate, pH 7.4) with a few crystals of phenylmethylsulfonyl fluoride and 1 μL/mL of protease inhibitor (Bioshop; Cat# PIC002, Burlington, ON, Canada) added just prior to homogenization. Homogenates were centrifuged for 15 min at 10,000× g (4 °C), and supernatants containing soluble protein were collected. Protein concentration of each sample was quantified using the BioRad protein assay, as per the manufacturer’s instructions (Cat# 5000002; Hercules, CA, USA). Protein concentrations were then standardized to 10 µg/µL by addition of small aliquots of homogenization buffer. Standardized samples were mixed 1:1 v/v with SDS buffer (100 mM Tris-HCl, 4% w/v SDS, 20% v/v glycerol, 0.2 w/v bromophenol, 10% v/v β-mercaptoethanol, pH 6.8) to a final concentration of 5 µg/µL. Finally, samples were boiled for 10 min to denature and linearize all proteins and then stored at −40 °C until use.

2.3. Immunoblotting

Equal amounts of total protein homogenates (25–40 µg depending on target being probed) of control, 24 h frozen, and 8 h thawed samples were loaded on 6–15% discontinuous SDS-PAGE. Gels were run on a BioRad Mini Protean III system at 180 V for 1–3 h at 4 °C in running buffer (25 mM Tris-base, 190 mM glycine, 0.1% w/v SDS, pH 7.6). Resolved protein gels were then transferred to 0.45 µm pore polyvinylidene difluoride membranes at 160 mA for 1.5–16 h in pre-chilled transfer buffer (25 mM Tris-base, 192 mM glycine 10% v/v methanol, pH 8.5) at 4 °C. Transferred membranes were air-dried for 15 min, reactivated in methanol for 5 min and incubated in 2–10% v/v skim milk in TBST (20 mM Tris-base, 140 mM NaCl, 0.05% v/v Tween-20) for 10–45 min with rocking at room temperature (RT). Blocked membranes were then washed 3 × 5 min in TBST and subsequently incubated overnight with the primary antibody of interest (1:1000 v:v dilution in TBST) with rocking at 4 °C. Primary antibodies were purchased for DROSHA (NeoBiolab; Cat# A8336), DGCR8 (GeneTex; Cat# GTX130061), DICER (SantaCruz; Cat# SC-30226), AGO1 (GeneTex; Cat# GTX47799), AGO2 (ECM BioSciences; Cat# AP5281), p-AGO2^Tyr393 (ECM BioSciences; Cat# AP5311), p-AGO2^Ser387 (ECM BioSciences; Cat# Ap5291), RAN (GeneTex; Cat# GTX114139), EXPORTIN-5 (GeneTex; Cat# 130727), TRBP (GeneTex; Cat# GTX485546), and PACT (GeneTex; Cat# GTX114215).

Next, membranes were washed with TBST (3 × 5 min) and probed for 30 min at RT with horseradish peroxidase-conjugated goat anti-rabbit secondary antibody (1:8000 v/v dilution in TBST; Cat# APA002P BioShop; Burlington, ON, Canada). Finally, membranes were again washed 3 × 5 min in TBST, and protein bands were visualized using enhanced chemiluminescence (H₂O₂ and Luminol) and imaged using the ChemiGenius BioImaging System (Syngene; Frederick, MD, USA). Total protein levels were visualized by staining membranes with Coomassie blue (0.25% w/v Coomassie Brilliant Blue, 7.5% v/v acetic acid, 50% methanol) and subsequently destained with destain solution (50 mL ddH₂O, 50 mL acetic acid, 150 mL methanol).

2.4. RNA Isolation

RNA isolation from liver samples of control, 24 h frozen, and 8 h thawed wood frogs (n = 4 individuals) was conducted as previously described [37]. Briefly, samples (~50 mg) of frozen tissue were homogenized and extracted in Trizol-chloroform. RNA was precipitated with isopropanol and washed with 70% ethanol prior to being air-dried and resuspended in RNase-free water. RNA concentration and purity were assessed spectrophotometrically with the 260/280 nm ratio using a Take3 micro-volume quantification plate (BioTek; Winooski, VT, USA) and a PowerWave HT spectrophotometer (BioTek). Only samples with 260/280 ratios between 1.8 and 2.2 were used for miRNA analyses. Total RNA integrity was assessed by running isolated RNA on a 1% agarose gel stained with SYBR Green and by verifying the presence of sharp bands for 28S and 18S ribosomal RNA. RNA isolates were then standardized to equal concentrations by the addition of small volumes of RNase-free water and stored at −80 °C.

2.5. Polyadenylation and Stem–Loop Reverse Transcription

RNA samples were prepared for miRNA analysis as described previously [38]. Polyadenylation was performed using the Epi-Bio PolyA tailing kit, where each 10 μL reaction contained 3 μg total RNA, 1 mM ATP, and 0.5 μL (2 U) of E. coli poly (A) polymerase in buffered solution (0.1 M Tris-HCl, pH 8.0, 0.25 M NaCl, and 10 mM MgCl₂). Reactions were incubated at 37 °C for 30 min to adenylate, 95 °C for 5 min to arrest the reaction, and then immediately chilled on ice. Polyadenylated products (10 μL samples) were combined with 5 μL of 250 pM stem–loop RT adapter primer, heated to 95 °C for 5 min to denature RNA, cooled to 60 °C for 5 min to allow adapter annealing and placed on ice for 1 min (Supplementary Table S1). The SuperScript™ III Reverse Transcriptase kit was used on the polyadenylated and stem–loop primer-annealed RNA samples. Briefly, each sample was combined with 1 µL mouse Maloney leukemia virus (M-MLV) reverse transcriptase (2 U), 1 µL deoxynucleotide triphosphate (dNTP) mixture containing 25 mM of each nucleotide (Cat# R1121; ThermoFisher Scientific; Waltham, MA, USA), 2 µL 0.1 M dithiothreitol, and 4 µL 5× first-strand buffer. Samples were incubated at 16 °C for 30 min, 42 °C for 30 min, and 85 °C for 5 min. cDNA was serially diluted and frozen at −20 °C.

2.6. Relative microRNA Quantification

MicroRNA-specific forward primers were designed using all annotated frog (Xenopus tropicalis and Xenopus laevis) miRNAs deposited on miRBase v18 and Xenbase [39,40]. All miRNA, universal reverse primer, and reference gene primers were designed as described previously [38] and are listed in Supplemental Table S1. These candidates were experimentally tested via RT-qPCR, and primers that (1) generated a single peak melt curve and (2) were highly conserved via cross-referencing on miRBase were included in the analysis. Primers were synthesized by Integrated DNA Technologies. All RT-qPCR assays were performed as previously described [41] using a CFX Connect™ Real-Time PCR Detection System (BioRad; Cat# 1855201), following MIQE guidelines [42]. Each 20 µL RT-qPCR reaction consisted of 2 µL of diluted RT product, 10 µL of ddH₂O, 4 µL of 1 M trehalose (BioShop; Cat# TRE222), 2 µL RT-qPCR buffer (100 mM Tris-HCl, pH 8.5, 500 mM KCl, 1.5% Triton X-100, 20 mM MgCl₂, 2 mM dNTPs, and 100 nM fluorescein), 0.5 µL formamide (BioShop; Cat# FOR001), 0.5 µL of 25 mM miRNA-specific forward primer, 0.5 µL of 25 mM universal reverse primer, 0.1 µL of 100x SYBR green mix diluted in dimethyl sulfoxide (Cat# S7585; Invitrogen; Waltham, MA, USA), 0.16 µL of 25 mM dNTPs, and 0.125 µL of 5 U/µL Taq polymerase (BioShop; Cat# TAQ001.1). The following PCR program was used: 95 °C for 3 min, followed by 40 cycles of 95 °C for 15 s, and 60 °C for 1 min. To ensure primer specificity and the amplification of a single PCR product, all RT-qPCR assays were subject to post-run melt–curve analysis; reactions that generated non-specific products were rejected.

2.7. Bioinformatics microRNA Target Identification and Pathway Enrichment

To identify potential miRNA–mRNA interactions, mature miRNA sequences were searched against the 3′UTR sequences from the Xenopus tropicalis reference genome available on the UCSC table browser (JGI 7.0/xenTro7) using the temperature-sensitive miRNA target prediction program FINDTAR3 (v.3.11.12) [43]. Target predictions were performed at −2 °C and 5 °C for miRNAs that significantly changed during freezing and thawing, respectively. The following default parameters were used with FINDTAR3: AT and GC weight of 5, GT weight of 2, a gap opening penalty of −8, a gap extension penalty of −2, target duplex with maximum threshold-free energy −20 kcal/mol, and demand strict 5′ seed pairing.

The list of predicted miRNA targets generated by FINDTAR3 was then functionally enriched to examine the connectivity of miRNA-targeted mRNA. This was done by mapping protein–protein interactions using STRING X. tropicalis medium confidence interactions [44]. Two groups were queried separately: (1) the 10 miRNAs upregulated in frozen livers and (2) the 13 miRNAs upregulated in thawed livers. Protein interaction networks were then modeled using CYTOSCAPE software v3.10.1 [45] and clustered based on STRING combined scores using Markov Clustering (MCL) with the following default parameters: MCL granularity of 2 and edge weight cut-off of 0.5. Enrichment for biological processes was performed using the Gene Ontology (GO) annotations available for X. tropicalis using the PANTHER classification system (v.11.1). The enriched biological processes discussed and highlighted are statistically significant clusters, as determined by Bonferroni tests with p < 0.05.

2.8. Quantification and Statistics

Relative protein densitometric quantification was performed on chemiluminescent immunoblot protein bands using GeneTools Software v4.3.17 on a ChemiGenius BioImaging System (Syngene, Frederick, MD, USA). Immunoblot band intensity in each lane was standardized against a group of Coomassie blue-stained protein bands to correct for any minor variations in sample loading [46]. Immunoblot data for each experimental condition are expressed as means (±SEM) relative to control values, with n = 4 samples from different animals. Statistical analysis was performed by one-way ANOVA and Dunnett’s post-hoc test (p < 0.05 accepted as significant) using the RBIOPLOT statistics and graphing R package [47].

MicroRNA relative expression levels were calculated using the comparative ΔΔCq method [37]. Raw Cq values were transformed to the 2^−Cq form, such that the miRNA of interest could be standardized to the endogenous U6 snRNA reference gene. U6 snRNA was experimentally determined to be a suitable reference gene based on its stable expression in R. sylvatica liver under all experimental conditions, tested as described by [48]. Data are mean relative expression levels (±SEM) relative to control values, n = 4 independent biological replicates from different animals at each sampling point. Statistically significant changes in miRNA relative expression were identified using one-way ANOVA and Dunnett’s post-hoc test (p < 0.05 accepted as significant). Statistical analyses and histogram generation were performed using the RBIOPLOT statistics and graphing R package [47].

3. Results

3.1. Protein Expression of miRNA Biogenesis and Processing Machinery

Immunoblotting was used to examine the abundance relative to controls (standardized to 1.0) of miRNA biogenesis proteins in wood frog livers over a freeze–thaw cycle. During freezing, microprocessor protein levels of DROSHA and DGCR8 were significantly upregulated by 1.89 ± 0.11-fold and 1.80 ± 0.08-fold as compared with controls, respectively (Figure 1). This significant upregulation was sustained during thawing for both DROSHA and DGCR8 with levels that were 1.894 ± 0.192-fold and 2.152 ± 0.202-fold greater than controls, respectively. Protein levels of the nuclear export machinery, XPO5 and RAN, were also significantly upregulated during freezing to 2.019 ± 0.120-fold and 2.046 ± 0.190-fold over controls, respectively (Figure 1). Again, this upregulation was sustained during thawing for both XPO5 and RAN with levels that were 1.522 ± 0.163-fold and 2.349 ± 0.095-fold over controls, respectively. However, protein abundance levels of the RISC complex components, DICER, TRBP, and PACT, all remained constant over the course of the freeze–thaw cycle (Figure 1). AGO1 and AGO2 total protein abundance also remained constant over freezing and thawing (Figure 1). However, the relative phosphorylation of AGO2 changed significantly over freeze/thaw. Levels of p-AGO2^Ser387 decreased significantly during thawing by 53.6% relative to controls, whereas p-AGO2^Tyr393 content decreased significantly by 86.3% during freezing and remained low at 72.4% below controls after thawing (Figure 1).

3.2. Differential miRNA Expression over the Freeze–Thaw Cycle

A large-scale analysis of the expression of 109 miRNA species using RT-qPCR identified 20 freeze–thaw responsive miRNAs in wood frog livers (Figure 2). Of these, 13 miRNAs (11.9%) were freeze-responsive, and 15 miRNAs (13.8%) were thaw-responsive, with 8 miRNAs (7.34%) overlapping between freezing and thawing. The majority of significantly differentially expressed miRNAs were upregulated during both freezing and thawing. Of the 13 miRNAs found to display significant differential expression during freezing, the following 10 (9.2% of total miRNAs) increased by 1.48–2.57-fold relative to controls: rsy-miR-101a-3p, rsy-miR-181a-3p, rsy-miR-192-5p, rsy-miR-199a-5p, rsy-miR-200a-3p, rsy-miR-210-3p, rsy-miR-221-3p, rsy-miR-222-3p, rsy-miR-22-3p, and rsy-miR-301-3p (Figure 2 and Supplementary Table S2). By contrast, only three miRNAs (2.8% of total miRNAs) were found to be significantly downregulated to 0.42–0.63 of control levels during freezing: rsy-miR-93a-5p, rsy-miR-9406-3p, and rsy-miR-9-5p (Figure 2 and Supplementary Table S2).

During thawing, 15 miRNAs were significantly differentially expressed and, similar to the effects of freezing, the majority of these miRNAs were upregulated (Figure 2). Of the 15 thaw-responsive miRNAs, the following 13 (11.9%) were significantly upregulated during thawing to 1.432–3.481 of control values: rsy-miR-10a-5p, rsy-miR-145-5p, rsy-miR-181a-3p, rsy-miR-192-5p, rsy-miR-199a-5p, rsy-miR-200a-3p, rsy-miR-208-3p, rsy-miR-210-3p, rsy-miR-221-3p, rsy-miR-23-3p, rsy-miR-27-3p, rsy-miR-30a-3p, and rsy-miR-429-3p (Figure 2; Supplementary Table S2). Only rsy-miR-93a-5p and rsy-miR-9406-3p were found to decrease to 0.426 ± 0.069 and 0.592 ± 0.085 of control levels, respectively, during thawing (1.8%; Figure 2; Supplementary Table S2).

The following six miRNAs were significantly elevated (5.5%) during both freezing and thawing: rsy-miR-181a-3p, rsy-miR-192-5p, rsy-miR-199a-5p, rsy-miR-200a-3p, rsy-miR-210-3p, and rsy-miR-221-3p. Conversely, rsy-miR-93a-5p and rsy-miR-9406-3p were significantly downregulated during both freezing and thawing (1.8%; Figure 2; Supplementary Table S2).

3.3. Bioinformatic Analyses of miRNA-Targeted Pathways

A functional cluster enrichment of downstream miRNA targets was performed using temperature-sensitive miRNA/mRNA target prediction, protein–protein interaction analysis, and the identification of enriched biological processes. The key processes targeted by the 10 freeze-upregulated miRNAs were: (1) intracellular signal transduction. (2) RNA processing and splicing, (3) protein ubiquitination, (4) carbohydrate catabolic processes and hexose metabolism, (5) DNA replication, (6) translation pre-initiation complex formation, (7) protein de-ubiquitination, (8) cell redox homeostasis, (9) ATP hydrolysis, (10) microtubule nucleation and polymerization, (11) intraciliary transport, and (12) lipid phosphorylation (Figure 3; Supplementary Table S3). The key networks targeted by the subset of 13 thaw-upregulated miRNAs included the following processes: (1) chromatin remodeling, (2) protein ubiquitination, (3) RNA processing and splicing, (4) carbohydrate catabolism and hexose metabolism, (5) tissue development, (6) microtubule nucleation and polymerization, (7) translation pre-initiation complex formation, (8) ATP hydrolysis, (9) intraciliary transport, (10) lipid phosphorylation, and (11) DNA replication (Figure 4; Supplementary Table S4). All miRNA-targeted processes are listed in the supplementary table. Due to the current limitations of the annotated X. tropicalis protein network interactions, the majority of remaining clusters were unclassified or mapped to non-liver-specific processes.

4. Discussion

In species that are cold tolerant, a subset of cold-associated miRNAs dubbed “cryomiRs” appears to be present in both vertebrates and invertebrates and has been implicated in biological processes involving muscle atrophy prevention, cell cycle regulation, and glucose/lipid metabolism [36]. The roles that miRNAs play in coordinating complex liver functions during periods of environmentally induced hypometabolism have also been reported in evolutionarily distant species ranging from hibernating South American marsupials [37] to cold-tolerant insects [49,50] to estivating African clawed frogs [51]. The wood frog liver must be under tight transcriptional and translational control to coordinate cryoprotective processes (e.g., glucose synthesis and distribution, among others) against a backdrop of global hypometabolism [3]. As such, the main question of interest for this study was—does the liver rely on miRNA-mediated silencing to regulate and suppress non-vital processes that are not required during prolonged periods in the frozen state?

4.1. Members of the miRNA Biogenesis Pathway Were Upregulated During Freezing and Remained High During Thaw

A tight regulation of proteins/enzymes involved in miRNA biogenesis plays a large part in generating the unique miRNA expression fingerprints seen in different animals, tissues, developmental stages, cell types, subcellular compartments, and stress responses. Our investigation of the protein levels of key miRNA biogenesis factors suggested an enhanced capacity for miRNA biosynthesis in wood frog liver under frozen and thawed conditions. Significantly elevated protein levels of the microprocessor complex subunit (DROSHA and DGCR8) were identified during freezing (Figure 1), and such enhancements have been shown to promote miRNA processing [52,53]. DROSHA has been implicated as the main regulator and rate-limiting step of miRNA biogenesis, and studies on Xenopus oocytes have demonstrated that increased DROSHA protein levels strongly boost miRNA biogenesis and pri-miRNA processing [54]. The nuclear export of pre-miRNA to the cytoplasm for further processing also serves as a critical regulatory step in miRNA biogenesis, and increased levels of XPO5 and its regulator, the small GTPase RAN, observed during freezing suggest enhanced miRNA nuclear export (Figure 1). Accordingly, the overexpression of XPO5 has been shown to enhance RNA interference mediated by endogenous miRNAs [55,56]. Once in the cytoplasm, the pre-miRNA is further cleaved by the endonuclease DICER and its binding partners TRBP and PACT to form a ds-miRNA duplex [19]. The unchanged levels of RISC proteins DICER, TRBP, and PACT over the freeze–thaw cycle suggest that this step proceeds unaffected by freezing stress. A possible explanation is that non-canonical miRNA biogenesis pathways, such as the DROSHA and DGCR8-independent pathway, spliceosome-dependent mechanisms, or the terminal uridylyl transferase-dependent pathway, all converge at DICER processing and, therefore, levels of this global protein must remain constant (Figure 1) [57]. The constant protein levels of DICER observed herein match previous measurements of DICER levels in the liver of frozen wood frogs and demonstrate the consistency and reproducibility of these findings [32].

The mature miRNA guide strand binds to a member of the AGO protein family, typically AGO1 or AGO2, to induce the cleavage or translational suppression of target mRNAs. Interestingly, protein levels of both AGO1, which interacts with ~30% of miRNAs, and AGO2, the sole AGO protein member with slicer activity that interacts with ~60% of miRNAs, remained constant over the freeze–thaw cycle (Figure 1) [58]. However, studies have shown that AGO2 functionality is heavily modulated by post-translational modifications (PTMs), and further examination revealed significantly reduced levels of p-AGO2^Tyr393 during freezing and thawing (Figure 1). Since phosphorylation at Tyr-393 reduces AGO2 binding to DICER and leads to inhibition of pre-miRNA processing and maturation, the reduction in p-AGO2^Tyr393 content indicates an overall increase of AGO2 miRNA processing capacity [59]. AGO2 is involved in both miRNA biogenesis and function, and its phosphorylation at Ser-387 has been shown to lead to reduced mRNA cleavage and enhanced translational repression [60]. Hence, the thaw-downregulated levels of AGO2^Ser387 suggest that mRNA cleavage, as opposed to translation inhibition or destabilization, is favored during thawing. However, this observation requires a deeper investigation of miRNA translational silencing machinery and an analysis of stress granule and P-body components. Overall, our examination of the protein and PTM levels of the main miRNA biogenesis factors suggests a potential activation of canonical miRNA biosynthesis during both freezing and thawing (Figure 1). This enhanced capacity for miRNA biogenesis is likely facilitating the selective increase of miRNAs identified to be overexpressed during freezing and thawing (Figure 2).

4.2. Differential Regulation of Cold-Associated miRNAs May Link to Cryoprotection and Suppression of Energy-Expensive Processes

Of the 109 miRNAs examined in this study, 10 were significantly upregulated, while only 3 were downregulated during freezing (Figure 2). Functional in silico target enrichment of the 10 freeze-upregulated miRNAs predicted the suppression of the following energy-expensive processes via miRNA action: RNA processing and splicing, DNA replication, intracellular signal transduction, microtubule nucleation and polymerization, protein turnover, and ATP hydrolysis (Figure 3 and Supplementary Table S3). The predicted suppression of these pathways makes metabolic sense since a well-known characteristic of freezing is the global suppression of ATP-demanding processes, such as cell growth and proliferation [3]. Indeed, a previous study of the cell cycle in the liver of frozen wood frogs showed strong evidence of cell cycle suppression due to reduced protein levels of cyclin-dependent kinases (Cdks) and cyclins A and B1 [61]. The current study provides a miRNA-mediated mechanism for cell cycle suppression since freeze-upregulated miRNAs were predicted to target Cdk 45, cyclin E2, Origin recognition complex subunits 1 and 6, DNA replication complex GINS3, and the transcription factor E2F4 that controls cell cycle progression and proliferation (Supplementary Table S3).

One of the most energetically expensive cell processes is protein synthesis and turnover, which utilizes 25–30% of total cellular ATP output [62]. Therefore, it makes metabolic sense that these pathways are predicted to be miRNA-silenced during freezing (Figure 3). Specific miRNA targets predicted to be inhibited include the following: translation pre-initiation complex proteins, elongation initiation factors, poly-A binding protein, and 60S acidic ribosomal protein P0 (RPLP0) (Figure 3 and Supplementary Table S3). Interestingly, a study on RPLP0 in wood frog liver showed a 2-fold freeze-upregulation of RPLP0 transcript levels but reported that RPLP0 protein levels remained unchanged. This disconnect between mRNA transcript and protein levels could be a result of the targeting of rsy-miR-101a-3p to the RPLP0 mRNA transcript [63] (Supplementary Table S3). In addition to reducing protein translation, frozen frogs also need to maintain their existing protein pool and must, therefore, limit protein degradation. One possible miRNA-mediated mechanism for this is via the silencing of both protein ubiquitinating proteins (UBE2E3—ubiquitin-conjugating enzyme E2 and E3) and de-ubiquitinating proteins (USP1, USP30, USP46—ubiquitin-specific proteases 1, 30, and 46) observed herein (Figure 3 and Supplementary Table S3).

Wood frog freezing survival is critically dependent on their ability to mobilize vast amounts of cryoprotectant glucose in the liver via a glycogenolytic response that raises liver glucose levels from 5 mM to greater than ~300 mM within a few hours and exports huge amounts of glucose for uptake by other organs [64]. To facilitate the synthesis of cryoprotectant glucose and its distribution to the rest of the body, various routes of glucose catabolism must be inhibited. A subset of the freeze-upregulated miRNAs was predicted to do exactly that by facilitating the suppression of genes coding for various enzymes of carbohydrate and hexose catabolic pathways (Figure 3). For example, the targeting of rsy-miR-199a-5p to the aldob gene that encodes aldolase B (the liver isozyme), which is one of the central steps in glycolysis, suggests a novel miRNA-mediated mechanism for suppression of hexose phosphate catabolism—a necessity in order to direct the vast majority of carbon from glycogenolysis into glucose synthesis as the cryoprotectant (Supplementary Table S3). Indeed, liver aldolase activity was significantly suppressed by about 40% during freezing in wood frogs but rebounded after thawing [65]. Furthermore, suppression of other carbohydrate catabolism pathways may also be required during freezing. Freeze-upregulated rsy-miR-210-3p was predicted to target NADP-isocitrate dehydrogenase, an enzyme that catalyzes the oxidative decarboxylation of isocitrate into α-ketoglutarate (Supplementary Table S3). The freeze-induced downregulation of rsy-miR-9-5p, observed herein, is implicated in promoting hepatic gluconeogenesis and glucose production via a FOXO1-mediated mechanism—a link first characterized in obese mice with reduced miR-9 levels that contributed to activated gluconeogenesis (Figure 2) [66]. Taken together, these newly identified hepatic miRNA mechanisms can help to prevent glucose breakdown and promote net glucose synthesis and output from the liver for wood frog cryoprotection.

Coincidentally, rsy-miR-181a was also overexpressed in the liver of frozen wood frogs and is likely playing a role in instigating the insulin resistance required to allow glucose to rise to cryoprotectant levels. While the pancreatic response of frozen frogs to ~300 mM rising glucose levels has been shown to be intact, leading to a doubling of plasma insulin levels during freezing, the subsequent uptake of glucose into insulin-sensitive organs is not observed [67]. Despite this feature being previously attributed to the novel sequence difference in wood frog insulin that has been associated with a low potency insulin, our findings point to a possible novel underlying miRNA-mediated mechanism that could also be contributing to the freeze-responsive insulin resistance required for freezing [68,69]. It is interesting to note that hibernating thirteen-lined ground squirrels also displayed elevated levels of miR-181a during torpor [70]. The authors suggest this likely contributes to the observed reversible hepatic insulin resistance exhibited over torpor-arousal cycles. In squirrels, the stimulation of insulin resistance during the initial periods of hyperinsulinemia could act to maintain the storage of fats required to fuel hibernation, and miR-181a might be involved in coordinating this adaptive response [71,72].

Evidence from wood frogs, other hypometabolic animals, and oxygen-deprived organisms has shown that the enhancement of antioxidant defenses is critical for protection against rapid reoxygenation upon thawing or during arousal from hibernation or reintroduction of oxygen into anoxic systems [3]. Therefore, it was surprising to find that a subset of the freeze-upregulated miRNAs included those that were predicted to modulate cellular redox by suppressing the mRNA transcripts of glutaredoxin 2, peroxiredoxin 1, and thioredoxin reductase (Supplementary Table S3). These enzymes all act to boost the antioxidant capacity of cells and promote the reduction of thioredoxin, an essential scavenger of reactive oxygen species [73]. The incongruence observed between the prediction of miRNA-mediated suppression of this antioxidant process, which has been previously shown to be active, could be attributed to the fact that many major antioxidant enzymes examined in wood frogs are primarily regulated at the post-translational level [3,5]. Indeed, a study of superoxide dismutase (SOD) from frozen frogs found that while total protein levels of Cu-SOD and Mn-SOD were not upregulated during freezing, these enzymes were significantly post-translationally modified and displayed enhanced activity during freezing [74]. This observation that antioxidant enzymes might be primarily modulated at the post-translational level suggests that cells need not use their limited energy stores to synthesize new antioxidant enzymes, thereby providing a possible energy-saving strategy in the frozen state.

Our examination of miRNA regulation after 8 h thawing found 13 miRNAs that were upregulated and only 2 miRNAs that were downregulated. Downstream targeting analysis of the thaw-upregulated miRNAs showed sustained targeting of many ATP-demanding processes that were predicted to be suppressed during freezing (Figure 3 and Figure 4). While frogs that have been thawed for 8 h display a normal physical posture and reactions, a full metabolic and molecular recovery is likely to be a slow process. Indeed, the reconversion of glucose into glycogen stores, the catabolism of anaerobic end products, and the restoration of ATP and phosphagen pools have been shown to take several days to return to control levels [64,75]. This evolved delay in recovery is metabolically sensible since it would facilitate survival and allow a rapid response if freezing were to recur again quickly. As such, our findings suggest that miRNAs are involved in maintaining this adaptive delay by sustaining the suppression of these energy-expensive processes 8 h into thawing.

Our current miRNA findings in the liver are in stark contrast to the responses observed previously in the brains of frozen wood frogs [24]. Brains from frozen and thawed frogs exhibited a global downregulation of miRNA biogenesis proteins that was coupled with a reduced capacity for miRNA biogenesis. The effects of this reduced processing capacity were seen in the downregulation of 39 of the 41 differentially expressed miRNAs over the freeze–thaw cycle [24]. Downregulated brain miRNAs were linked with mediating neuroprotective mechanisms that could fine-tune synaptic functions and maintain cell functions required for successful whole-brain endurance of freezing and of their component anoxia and dehydration stresses. This contrasts with our current liver study, where both miRNA biogenesis and differentially changing miRNAs were generally upregulated during freeze/thaw. These opposite trends make metabolic sense as they appear to support the unique metabolic needs of each tissue. The activation of miRNA-induced neuroprotection and maintenance is required to ensure that after thawing, wood frogs are able to restore normal brain function and illustrate a mechanism that potentially allows brains to circumvent the extensive translational silencing being undertaken across the rest of the body. In the liver, by contrast, miRNA action appears to limit the use of ATP stores by suppressing various non-essential energy-expensive anabolic and pro-growth functions in the frozen state in favor of selected targeted actions such as production and export of cryoprotectant glucose. In the liver of another freeze-tolerant frog, D. versicolor, levels of DROSHA, DICER, TRBP, and XPO5 were upregulated during freezing [76]. While only two of these proteins (DROHSA and XPO5) directly mirror our results, the functional consequence of increased miRNA biogenesis pathway activity remains the same for both species. Furthermore, a small RNA-sequencing study in D. versicolor liver predicted a subset of 11 miRNAs (7 upregulated and 4 downregulated) to be differentially regulated during freezing [35]. While none of the predicted miRNAs overlapped with our results, the downstream results of miRNA-mediated inhibition of signaling pathways, protein ubiquitination, cell cycle, and chromatin/nuclear process share many commonalities with our study, as already discussed above [35]. This points to a semi-conserved response in both frog species to suppress these processes in the liver during freezing, which expands our knowledge of freeze-tolerance and miRNA regulation during this process as a whole.

5. Conclusions

Overall, our findings highlight a miRNA-mediated mechanism that can be, in part, responsible for facilitating freeze-tolerance in wood frog liver. The general freeze- and thaw-induced miRNA upregulation observed herein may contribute to the measured increase in the protein levels of key miRNA biogenesis and processing factors. The miRNA fingerprint of livers from frozen and thawed frogs was predicted to silence key ATP-demanding functions such as those involved in pro-growth processes and in maintaining cellular homeostasis, both of which are non-essential when animals are in a hypometabolic state. This pattern of miRNA-mediated silencing of energy-expensive processes during extreme stress appears to be evolutionarily conserved across distant species and survival strategies. This study emphasizes the complexity of the expansive miRNA networks required for successful freeze-tolerance and highlights select miRNA species (and their target mRNAs) that could be further investigated as hepatic “on/off” switches with potential therapeutic applications.