The effect of hibernation on protein phosphatases from ground squirrel organs

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ABB Archives of Biochemistry and Biophysics 468 (2007) 234–243 www.elsevier.com/locate/yabbi

The effect of hibernation on protein phosphatases from ground squirrel organs Justin A. MacDonald a

a,*

, Kenneth B. Storey

b

Department of Biochemistry and Molecular Biology, University of Calgary, 3330 Hospital Drive NW, Calgary, Alta., Canada T2N 4N1 b Institute of Biochemistry and Department of Biology, Carleton University, 1125 Colonel By Drive, Ottawa, Ont., Canada K1S 5B6 Received 7 August 2007, and in revised form 1 October 2007 Available online 13 October 2007

Abstract Protein phosphorylation has been identified as a reversible mechanism for the regulated suppression of metabolism and thermogenesis during mammalian hibernation. The effects of hibernation on the activity of serine/threonine and tyrosine protein phosphatases (PP1, PP2A, PP2C and PTPs) were assessed in five organs of Richardson’s ground squirrel. Each phosphatase subfamily responded differently during torpor, and each showed organ-specific patterns of activity changes. The distribution of PP1 catalytic subunit (PP1c) isoforms (a, d, c1) was assessed in five organs, and changes in the subcellular distribution of PP1 were observed during hibernation in liver and muscle. For example, in muscle, cytosolic PP1 content increased and myofibril-associated PP1 decreased during torpor. PP1c from ground squirrel liver was purified to homogeneity and characterized; temperature effects on PP1c maximal activity suggested that temperature had little or no effect on relative dephosphorylation potential at low temperatures. However, nucleotide inhibition of PP1c by ATP, ADP and AMP was much weaker at 5 C compared with 37 C assay temperatures. PP2A activity decreased in three organs (brown adipose, kidney, brain) during hibernation whereas PP2C activity was increased in liver and brain. PTPs were assessed using both a general substrate (ENDpYINASL) and a substrate (DADEpYLIPQQG) specific for PTPs containing the SH2-binding site; both revealed hibernation-associated changes in PTP activities. Changes in protein phosphatase activities suggest the relative importance of these modules in controlling metabolic function and cellular processes during mammalian hibernation.  2007 Elsevier Inc. All rights reserved. Keywords: Mammalian hibernation; Torpor; Spermophilus richardsonii; Phosphorylation; Protein phosphorylation; Protein phosphatase; PP1; PP2A; PP2C

Winter hibernation allows small mammals to minimize metabolic energy costs at a time when a scarcity of food and cold environmental temperatures endanger normal life. By hibernating, animals can reduce their energy requirements by at least 90% and survive for many months while slowly catabolizing body lipid reserves [1]. While hibernating, metabolic rate drops to low levels, frequently to as little as 1–2% of the normal resting rate of euthermic animals, and core body temperature falls to near ambient (often to 0–5 C) [2]. Research in recent years has demonstrated that metabolic rate depression during hibernation is an active, controlled process that includes the coordinated suppres*

Corresponding author. Fax: +1 613 520 4389. E-mail address: [email protected] (K.B. Storey).

0003-9861/$ - see front matter  2007 Elsevier Inc. All rights reserved. doi:10.1016/j.abb.2007.10.005

sion of metabolism [3], the reorganization of various cellular functions [4–6], and the induction of selected genes and processes to provide long term stability in the dormant state [7,8]. One of the biochemical regulatory mechanisms that figures prominently in metabolic suppression during hibernation is reversible protein phosphorylation [9,10]. The reversible phosphorylation of enzymes of carbohydrate and lipid catabolism has been well studied in hibernation [11–13] and provides a means for the rapid change in metabolic flux that underlies both the transition into torpor and the high rates of thermogenesis that drive re-warming of the body during arousal. Protein phosphorylation is dynamic and is modulated by the action of protein kinases and protein phosphatases. In recent studies, we have studied protein kinases in mammalian hibernation [14–18].

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Organ-specific changes in kinase activities and protein kinase functional adaptation to the temperature range (i.e., 2–37 C) that characterizes euthermia and torpor were investigated. However, comprehensive studies on protein phosphatases from mammalian hibernators have yet to be conducted. Protein phosphatases play key roles in regulating diverse physiological events such as cell division, metabolism, motility, transcription and translation. Protein tyrosine phosphatases (PTPs)1 catalyze the dephosphorylation of proteins on tyrosine residues. The PTPs are broadly classified as both membrane receptor and non-receptor PTP families. The protein serine/threonine phosphatases are classified into three broad categories, namely type-1 phosphatase (PP1), several type-2 phosphatases (including PP2A, PP2B, and PP2C), and additional PP2A-like phosphatases (such as PP4, PP5, PP6, and PP7) [19]. Several holoenzyme forms of PP1 are known. These contain the PP1 catalytic subunit (PP1c) associated with different regulatory subunits: glycogen-bound PP1 consists of PP1c plus a glycogen binding subunit in muscle (PP1GM) [20] and PP1GL in liver [21], myosin-associated PP1 consists of PP1c bound to a myosin targeting subunit [22], and nuclear PP1 consists of PP1c associated with NIPP-1 (nuclear inhibitor of PP1) [23]. The role of each regulatory subunit type is to target PP1c to a specific subcellular location and to modulate PP1c activity toward a selective pool of substrates. For example, glycogen-associated PP1 is a key regulator of glycogen metabolism, being the major phosphatase that dephosphorylates the enzymes of glycogen degradation (i.e., glycogen phosphorylase) and synthesis (i.e., glycogen synthase) [20]. Members of the type-2 phosphatases are subdivided on the basis of their metal ion requirements: spontaneously active (PP2A), Ca2+/calmodulin-dependent (PP2B), and Mg2+-dependent (PP2C) [19]. Research has revealed prominent roles for PP2A in cell cycle regulation, translation initiation, apoptosis, cell morphology and development [24]. Several holoenzyme forms of PP2A have been characterized with the core enzyme consisting of a 36 kDa catalytic subunit (PP2Ac) complexed with a regulatory A subunit. This core dimer can also be associated with additional B subunits. Unlike the PP1 and PP2A phosphatases, members of the PP2C family are monomeric, lacking regulatory subunits. The PP2Cs contribute to the generalized maintenance of stress-induced protein kinase (SAPK) cascades in an inactive state under non-stressed conditions [25,26]. SAPKs are activated by various extracellular stimuli, including environmental stresses and inflammatory cytokines.

1

Abbreviations used: PTPs, protein tyrosine phosphatases; PP1, type-1 phosphatase; PP2A, PP2B, and PP2C, type-2 phosphatases; PP1c, PP1 catalytic subunit; PMSF, phenylmethylsulfonyl fluoride; TPCK, tosylphenylchloroketone; PEG, polyethylene glycol; PVDF, polyvinylidene difluoride; AMPK, AMP-activated protein kinase.

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The present study explores the tissue-specific responses of protein phosphatases, PP1, PP2A, PP2C and PTPs, during hibernation in Richardson’s ground squirrels in order to determine the role of these key regulators of reversible protein phosphorylation. In addition, PP1c from ground squirrel liver was characterized to address whether its intrinsic properties or its regulation by other factors modulate PP1 activity in a temperature-dependent fashion to affect desirable metabolic changes during hibernation. Materials and methods Animals and chemicals Adult Richardson’s ground squirrels, Spermophilus richardsonii, were obtained in the foothills of the Rocky Mountains near Calgary, Alta. All animals were individually housed at the animal care facility of the University of Calgary, Calgary, Alta. and maintained at 22 C on a fall (10L:14D) photoperiod. Ground squirrels were induced to hibernate as previously described [17]. Control animals were housed at 22 C prior to sacrifice. Tissues from both euthermic and hibernating squirrels were immediately excised, immersed in liquid nitrogen, and subsequently stored at 80 C. [c-32P]ATP (3000 Ci/mmol) was obtained from New England Nuclear (Montreal, PQ). Okadaic acid was from Calbiochem (La Jolla, CA) and microcystin–agarose was from Upstate Biotechnology Inc. (Lake Placid, NY). All other chemicals, column materials, and enzymes were purchased either from Sigma or Boehringer Mannheim. Rabbit polyclonal antibodies recognizing the carboxy-terminus of rat a, d, or c1 PP1c isozymes were a kind gift of Dr. Emma Villa-Moruzzi (University of Pisa, Italy). Goat anti-rabbit antibody coupled to horseradish peroxidase was purchased from Santa Cruz Biotechnology Inc. (Santa Cruz, CA). The ECL kit and reflection autoradiography film were from Mandel Scientific (Guelph, ON). 32P-labeled glycogen phosphorylase a was prepared as described previously [27].

Assay of ‘‘active’’ and ‘‘total’’ PP1 activity PP1 activity was assayed with 32P-labeled glycogen phosphorylase a (GPa) substrate, one unit (U) of activity being defined as the amount of enzyme that catalyzes the release of 1 nmol 32P-phosphate per min at 23 C. Unless otherwise indicated, the standard assay mixture contained in a 50 lL volume: 20 mM Tris–HCl, pH 7.0, 0.1 mM EGTA, 10 mM bmercaptoethanol, 5 mM theophylline, and 2.5 nM okadaic acid. Assays were initiated by the addition of 32P-labeled GPa and terminated by the addition of 100 lL ice-cold 25% (w/v) trichloroacetic acid/10 mM phosphoric acid. After incubating in an ice bath for 5 min, all samples were centrifuged and then 100 lL of each supernatant was removed and counted in an LKB 1900CA liquid scintillation counter. Two control assays were included and 32P released under these conditions was subtracted as a blank: (1) zero time control, assays were stopped immediately by the addition of TCA/phosphoric acid solution; and (2) minus enzyme control, buffer replaced the enzyme extract. Both controls gave similar results. Activities were measured both in concentrated tissue extracts (1:3 w/v homogenization) and after pre-treatment of tissue extracts with trypsin. Measurement of PP1 activity in concentrated tissue extracts provides an approximate measure of ‘‘active’’ phosphatase at physiological concentrations of modulating proteins and other factors [28]. However, whereas PP1c is relatively resistant to trypsinolysis, pre-treatment with trypsin removes regulatory proteins that would otherwise inhibit phosphatase activity and so gives a measure of ‘‘total’’ phosphatase present [29]. In this procedure, frozen tissue samples were homogenized with 2–3 volumes of 50 mM imidazole–HCl, pH 7.4, containing 0.25 M sucrose, 4 mM EDTA and 0.5 mM dithiothreitol. Homogenates were centrifuged at 1000g for

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3 min and the supernatant was saved. One aliquot was immediately assayed for ‘‘spontaneously active’’ PP1 activity. A second aliquot (100 lL) was incubated with 1 mg/mL trypsin and 1 mM MnCl2 for 10 min (25 C) and then digestion was stopped by the addition of soybean trypsin inhibitor to a concentration of 10 mg/mL. Dilution buffer (50 mM Tris, pH 7.0, 0.1 mM EGTA, 15 mM b-mercaptoethanol, 2 mg/mL bovine serum albumin) was added to prepare dilutions of 1:50 v/v for liver and 1:10 v/v for all other tissues. The diluted sample was then assayed for ‘‘total’’ PP1 activity.

EDTA, 2 mM EGTA, 10 mM b-mercaptoethanol, 0.1 mM sodium orthovanadate), plus 0.6 M KCl, 0.5% v/v Triton X-100, and protease inhibitors: 1 mM benzamidine, 1 mM PMSF, 20 lg/mL aprotinin, 10 lg/ mL tosylphenylchloroketone (TPCK), 5 lg/mL leupeptin, and 10 lg/mL soybean trypsin inhibitor and then centrifuged at 15,000g for 15 min. Supernatant was removed and polyethylene glycol (PEG) 8000 was added to 12% (w/v). After resting on ice for 20 min, the suspension was centrifuged at 15,000g for 15 min, and then the supernatant was discarded. The pellet was resuspended in the original volume of buffer A containing protease inhibitors. All purification steps were performed at 5 C.

Assay of type-2 protein phosphatase activities DEAE-cellulose chromatography The activities of type-2 protein phosphatases were assayed with a Serine/Threonine Phosphatase Assay Kit (Promega; Madison, WI). Frozen tissue samples were homogenized 1:10 (w/v) in ice-cold homogenization buffer (50 mM Tris, pH 7.0, 0.1 mM EGTA, 10 mM bmercaptoethanol) with 1 mM phenylmethylsulfonyl fluoride (PMSF), 1 lg/mL leupeptin, and 1 lg/mL aprotinin. Homogenates were centrifuged at 20,000g for 20 min. Supernatants were desalted on small columns of Sephadex G-25 to remove low molecular weight metabolites and endogenous inorganic phosphate that would interfere with the subsequent detection of protein phosphatase derived inorganic phosphate [30]. The PP2A assay mixture contained 50 mM imidazole–HCl, pH 7.2, 0.5 mM EGTA, 0.5 mM EDTA, 100 lM substrate hexapeptide (RRApTVA) and 0.02% v/v (b-mercaptoethanol). PP2C activity was measured against the same substrate peptide but in the presence of 1 lM okadaic acid and 10 mM MgCl2. Reactions were initiated by the addition of tissue extract, incubated for 10 min at 22 C, and terminated by addition of 50 lL malachite green dye reagent. After 30 min incubation, absorbance (600 nm) was measured with a Dynatech MR5000 Microplate reader and Biolynx Data Capture software. A blank reaction was performed simultaneously to control for phosphate release in the absence of substrate peptide. Initial trials determined the optimum amount of homogenate used in each assay and demonstrated that reaction rates were linear over the reaction time. One unit of activity is defined as the amount of enzyme that catalyzes the release of 1 nmol phosphate per min at 23 ± 1 C.

The resuspension was loaded onto a DEAE-cellulose (DE-52) column (2.5 · 8 cm) equilibrated in buffer B (50 mM Tris–HCl, pH 7.2, 0.1 mM EDTA, 10 mM b-mercaptoethanol, 10% v/v glycerol). The column was washed extensively with buffer B, and then developed with a linear gradient of 0–750 mM NaCl. Fractions (2 mL) were collected and assayed for PP1 activity. Fractions with activity were pooled, placed in dialysis tubing, and concentrated on solid PEG 8000.

Microcystin–agarose affinity chromatography Fractions containing PP1 activity were incubated with 1 mL of microcystin–agarose. The slurry was allowed to incubate for 5 h with constant mixing. The microcystin–agarose was transferred to a gravity column and washed extensively with buffer B containing 500 mM NaCl. Proteins were eluted with 10 mL of 3 M NaSCN in buffer B and dialyzed overnight at 5 C against buffer B.

Blue dextran chromatography The microcystin–agarose elution was applied to a blue dextran column (1 · 4 cm) equilibrated in buffer B. The column was washed with buffer B and then developed with a linear gradient of 0–750 mM NaCl in buffer B. Fractions with PP1 activity were pooled and concentrated on solid PEG 8000.

Sephacryl S-200 gel filtration Assay of tyrosine phosphatase (PTP) activity PTP activity was assessed with a Tyrosine Phosphatase Assay Kit (Promega; Madison, WI). Soluble and insoluble fractions of ground squirrel tissue homogenates were analyzed with either a general PTP substrate (ENDpYINASL) or a substrate (DADEpYLIPQQG) specific for PTPs containing the SH2-binding site. Frozen tissue samples were quickly weighed and homogenized 1:3 w/v in PTP homogenization buffer (25 mM TrisÆCl, pH 7.4, 2 mM EDTA, 2 mM EGTA, 250 mM sucrose, 10 mM b-mercaptoethanol, 1 mM PMSF, 1 lg/ml aprotinin, 5 mM benzamidine, 1 lg/ml pepstatin, and 1 lg/ml leupeptin). After centrifugation at 13,000g for 30 min, the supernatant (soluble fraction) was centrifuged through Sephadex G-25 columns. For activity in the insoluble fraction, pellets were re-homogenized using a glass pestle in one-third of the original volume (a volume that gave approximately the same protein concentration as in the soluble fraction) of PTP buffer containing 1% v/v Triton X-100 and left on ice for 1 h. After centrifugation as before, the supernatant was desalted through Sephadex G-25 columns and then used for assay. PTP assays were conducted in the presence of 50 mM imidazole–HCl, pH 7.2, 0.5 mM EGTA, 0.5 mM EDTA, 10 mM b-mercaptoethanol, with and without 200 lM substrate peptide (ENDpYINASL or DADEpYLIPQQG) to account for non-specific background phosphatase activity. Phosphate release was measured with the malachite green dye reagent as described earlier. One unit is defined as the amount of enzyme that releases 1 nmol phosphate per minute at 23 C.

Purification of PP1c from ground squirrel liver Frozen liver from euthermic ground squirrels was homogenized 1:5 (w/v) in buffer A (20 mM Tris–HCl, pH 7.2, 20 mM b-glycerophosphate, 2 mM

The concentrated sample was loaded onto a Sephacryl S-200 column (1 · 30 cm) equilibrated with buffer B plus 20% v/v glycerol and 100 mM NaCl. Fractions (0.5 mL) were collected and assayed for PP1 activity. Enzyme native molecular weights were determined by Sephacryl S-200 gel filtration with pyruvate kinase, aldolase, bovine serum albumin, carbonic anhydrase, and cytochrome c as standards. Fractions containing PP1c activity were dialyzed against buffer B with 40% v/v glycerol and stored at 20 C until used for kinetic analysis.

Enzyme kinetics Kinetic properties of purified PP1c were assessed at 37 and 5 C. Temperature was manipulated using a Lab-Line Multi-bloc thermal controller and a cold-water bath. Assay mixtures were pre-incubated for 15 min to allow thermal equilibration before initiating reactions with the addition of PP1c. All assays were conducted in 20 mM imidazole–HCl containing 0.1 mM EGTA and 15 mM b-mercaptoethanol plus various additions. The pH of imidazole–HCl buffers was adjusted at room temperature and allowed to vary with temperature so that the pH at 37 C was 7.2 and the pH at 5 C was 7.5. The pH values were selected to match intracellular pH measured in skeletal muscle of euthermic and hibernating animals as determined by in vivo 31P NMR spectroscopy [31].

Analysis of PP1c isozyme expression The expression of PP1c isozymes in various organs of euthermic ground squirrels was determined by Western blotting with isozyme-specific anti-PP1c antibodies [32]. PP1c was enriched by capture from tissue extracts with microcystin–agarose. Euthermic ground squirrel tissues were homogenized (1:4 w/v) in buffer A containing 0.1% v/v Triton X-100,

J.A. MacDonald, K.B. Storey / Archives of Biochemistry and Biophysics 468 (2007) 234–243 0.6 M NaCl, and protease inhibitors. The tissue extracts were resolved on 12% SDS–polyacrylamide gels and transferred to polyvinylidene difluoride (PVDF) membranes in 25 mM Tris, 192 mM glycine and 20% (v/v) methanol at 110 V for 60 min at 4 C. Non-specific binding sites were blocked with 5% (w/v) non-fat dry milk in TBST (25 mM Tris–HCl, pH 7.4, 150 mM NaCl and 0.05% (v/v) Tween 20). The blots were washed and incubated for 1 h with primary antibodies (1:1000 dilution) in TBST. Then the blots were washed again and incubated for 1 h with horseradish peroxidase-conjugated secondary antibody (1:5000 dilution) in TBST. Blots were developed with enhanced chemiluminescence (GE Healthcare).

Analysis of PP1 subcellular distribution Cytosolic and glycogen/microsomal (liver) or glycogen/sarcoplasmic reticulum (skeletal muscle) fractions were prepared from euthermic and hibernating ground squirrel liver and skeletal muscle [33]. Briefly, liver (1 g) was homogenized in 4 volumes of buffer A containing 250 mM sucrose and centrifuged at 10,000g for 10 min to remove cellular debris. The clarified homogenate was then centrifuged at 100,000g for 60 min. The 100,000 g supernatant (cytosolic fraction) and resuspended pellet (glycogen/microsomal fraction) were applied to microcystin–agarose columns as described earlier. Elutions from microcystin–agarose were transferred to PVDF membrane following SDS–PAGE and probed for PP1c by Western blot with a pan-PP1c antibody. For skeletal muscle, tissue (0.5 g) was homogenized in 3 volumes of buffer A containing 250 mM sucrose and then centrifuged at 10,000g for 10 min. The supernatant was removed and stored on ice; the pellet was washed with an additional 3 volumes of buffer A plus sucrose and re-centrifuged at 10,000g for 20 min. The pellet (myofibril fraction) was re-suspended in buffer A containing 0.2% (v/v) Triton X-100 and 0.6 M NaCl. The two supernatants were pooled and centrifuged at 100,000g for 60 min. The supernatant from the ultra-centrifugation step (cytosolic fraction) was removed, and the pellet (glycogen fraction) was resuspended in buffer A. All fractions were kept on ice until assayed for PP1 activity using 32 P-labelled GPa.

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Table 1 PP1 activity against glycogen phosphorylase in ground squirrel organs Tissue and condition

Total activity

‘‘Active’’ activity

% Active

Liver Euthermic Hibernating

73.8 ± 4.3 56.7 ± 2.4a

49.3 ± 2.8 48.1 ± 3.1

67 ± 7 85 ± 6a

Skeletal muscle Euthermic Hibernating

23.9 ± 0.6 34.4 ± 3.2a

24.3 ± 3.7 21.1 ± 1.3

100 ± 15 61 ± 9a

Kidney Euthermic Hibernating

35.4 ± 3.8 26.8 ± 2.2a

27.2 ± 2.7 8.60 ± 1.0a

77 ± 16 32 ± 6a

Brain Euthermic Hibernating

16.7 ± 2.1 32.2 ± 2.5a

10.5 ± 0.6 18.7 ± 1.1a

63 ± 12 58 ± 8

Heart Euthermic Hibernating

16.0 ± 1.3 12.5 ± 2.0

17.5 ± 0.8 4.3 ± 0.6a

100 ± 13 34 ± 10a

Activities are expressed as U/g wet weight tissue at 23 C. Data are means ± SEM for n = 4–6 separate determinations. Hibernating values are significantly different from those in corresponding euthermic tissues (Student’s t-test, two-tailed). a P < 0.05.

isoform expression. Skeletal muscle and kidney expressed significant amounts of all three PP1c isoforms. All tissues except liver contained significant amounts of PP1cc1.

Data and statistics Data on organ phosphatase activities were assessed by ANOVA with post hoc analysis (Dunnett’s test, two-tailed). Statistical testing of kinetic differences between assay temperatures for purified PP1c was performed with the Student’s t-test, two-tailed.

Results PP1 activity using GPa as the substrate was determined for five organs from euthermic and hibernating ground squirrels (Table 1). The highest total PP1 activity was found in liver with values that were 4.6-fold higher than the activities found in brain and heart. During hibernation, total PP1 activities decreased in liver and kidney by 23% and 24%, respectively, whereas activity increased in muscle (by 44%) and brain (by 93%), but did not change in heart. The spontaneously active component of PP1 dropped significantly during hibernation in heart and kidney (by 75% and 68%, respectively), increased in brain (by 82%), and was unchanged in muscle and liver. The percentage of PP1 activity present in the ‘‘active’’ form changed significantly in four out of five tissues during hibernation, rising in liver but decreasing in muscle, heart and kidney. Western blotting was used to characterize the PP1c isozymes expressed in different ground squirrel organs (Fig. 1). The data suggest tissue-specific patterns of PP1c

Fig. 1. Expression of PP1ca, PP1cd, and PP1cc1 proteins in various organs of euthermic ground squirrels. Microcystin–agarose was used to enrich PP1 in extracts from heart, skeletal muscle, liver, brain, and kidney. A sample of recombinant bovine PP1cc1 was used as a blotting control (+ve). Samples of column eluates containing 40 lg total protein were separated on 12% SDS–PAGE gels. This was followed by immunoblotting using the indicated antibodies and detection of a 37–39 kDa immunoreactive band by ECL chemiluminescence system with HRP-conjugated goat anti-rabbit secondary antibody.

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Fig. 2. PP1c levels in glycogen particle (G) and cytosolic (C) fractions of liver from euthermic and hibernating ground squirrels. Microcystin– agarose was used to enrich PP1c from the glycogen and cytosolic fractions. Care was taken to ensure equal total protein loading (40 lg) between the euthermic and hibernating samples. Immunoblotting with the anti-PP1c antibody and detection were as described in Materials and methods.

Equal expression of the three PP1c isoforms has been previously reported in mouse skeletal muscle [34]; however, there was variable expression of the three PP1c isoforms in ground squirrel skeletal muscle with high levels of PP1cc1, moderate levels of PP1cd, and low levels of PP1ca. Ground squirrel liver had relatively low expression of PP1c relative to the other tissues; only PP1cc1 and PP1cd isoforms were detectable. Rat liver was also shown to express PP1cd and PP1cc1 with the expression levels in liver lower than those found in other rat tissues [35]. Muscle, brain, and kidney had significant amounts of PP1cd expression whereas heart had very low amounts. Ground squirrel brain showed high levels of PP1cc1 and moderate levels of PP1cd, which is similar to the PP1c expression in rat brain [35]. The subcellular distribution of PP1c was assessed in liver extracts of euthermic and hibernating squirrels. Extracts were separated into cytosolic and glycogen-associated fractions, and PP1c in each fraction was detected by Western blotting with a pan-PP1c antibody. The majority of PP1c was found in the cytosolic fraction isolated from either euthermic or hibernating animals (Fig. 2). While PP1c was associated with the glycogen fraction of euthermic liver, the phosphatase was not detected in the glycogen fraction isolated from hibernating liver. Subcellular distribution of skeletal muscle PP1 activity The subcellular localization of PP1 activity in skeletal muscle (Fig. 3) as assessed in skeletal muscle isolated from either euthermic or hibernating squirrels. There was no difference in the total Triton X-100 extractable PP1 activity found in muscle extracts from euthermic or hibernating animals. In both states, the majority of PP1 activity was localized to the myofibrillar fraction with smaller amounts found in the cytosolic and glycogen fractions. Similar amounts of PP1 activity were associated with the glycogen fraction in both hibernating and euthermic conditions. Hibernation had a significant effect on enzyme distribution in the cytosolic and myofibrillar fractions; PP1 activity in the cytosolic fraction increased from 3.0 ± 0.2% of total in euthermia to 12.4 ± 5.4% in hibernation. Correspondingly, the percentage of total PP1 activity associated with

Fig. 3. Distribution of PP1 activity against glycogen phosphorylase a as the substrate in subcellular fractions of skeletal muscle from euthermic and hibernating ground squirrels. Enzyme activity is expressed as a percentage relative to activity of the euthermic enzyme in the crude homogenate. Subcellular fractions are (H) Triton X-100 extracted crude homogenate, (C) cytosolic supernatant, (G) glycogen particles, and (M) Triton X-100 washed myofibrils. Data are means ± SEM, n = 4 independent determinations. *Significantly different from the corresponding euthermic value, P < 0.05.

the myofibrils decreased from 76.6 ± 2.2% in euthermia to 67.4 ± 4.2% in hibernation. PP2A and PP2C phosphatase activities The effect of hibernation on the activities of type-2 protein phosphatases in ground squirrel organs is shown in Table 2. In euthermic squirrels, PP2A activity was highest in brain and heart at 6.6 mU/mg protein and was lowest in liver at 0.74 mU/mg protein. Hibernation had no effect on the PP2A activity measured in liver, muscle and heart. However, activities in kidney, brown adipose and brain Table 2 PP2A and PP2C activities in ground squirrel organs Tissue

Euthermic

Hibernating

PP2A Liver Skeletal muscle Kidney Brown adipose Brain Heart

0.74 ± 0.13 0.91 ± 0.16 1.53 ± 0.13 1.60 ± 0.14 6.53 ± 0.15 6.60 ± 0.93

0.73 ± 0.06 1.09 ± 0.19 0.95 ± 0.09a 0.99 ± 0.02a 4.60 ± 0.03a 5.92 ± 0.05

PP2C Liver Skeletal muscle Kidney Brown adipose Brain Heart

0.29 ± 0.06 ND 0.60 ± 0.04 0.28 ± 0.04 0.56 ± 0.07 0.20 ± 0.05

0.48 ± 0.03a ND 0.66 ± 0.05 0.36 ± 0.06 1.02 ± 0.11a 0.19 ± 0.02

Activities are expressed as mU/mg protein where 1 U is the release of 1 nmol phosphate per min at 23 C. Data are means ± SEM, n = 4–6 separate determinations. a Values are significantly different from the corresponding euthermic values (Student’s t-test, two-tailed), P < 0.05. ND—PP2C activity was not detected in skeletal muscle.

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were significantly lower in hibernating animals, 62–70% of the values found in the euthermic condition. PP2C activities in ground squirrel tissue are also shown in Table 2. In euthermic animals PP2C was highest in brain and kidney at approximately 0.6 mU/mg protein and lowest in heart (0.2 mU/mg protein); activity was not detected in skeletal muscle. PP2C activities in brain and liver increased during hibernation by 80% and 70%, respectively. Protein tyrosine phosphatase activities The effects of hibernation on protein tyrosine phosphatase activities in the soluble and insoluble fractions of ground squirrel tissues were assessed with two different substrates. Substrate 1 (ENDpYINASL) is a peptide used for the detection of a wide variety of PTPs and represents a highly conserved motif (DYINA) found in the catalytic domains of most PTPs. Intracellular PTPs reportedly exhibit a higher Vmax and affinity for this substrate than do receptor PTPs [36]. Substrate 2 (DADEpYLIPQQG) corresponds to the autophosphorylation site found in EGFR (amino acids 988–998); it is a substrate for PTPs which contain the SH2 (src-homology) binding site. Thus, the values obtained with substrate 1 can be used to indicate general effects on tyrosine dephosphorylation capacities of organs, whereas activities using substrate 2 more specifically represent only PTPs that contain the SH2-binding site. When assayed with substrate 1, total PTP activities in the soluble fraction rose during hibernation in brain and liver, increasing by 53% and 47%, respectively (Fig. 4a). Kidney and heart PTP activities remained unchanged. PTP activities in the insoluble fraction (Fig. 4b) were increased in liver and skeletal muscle, increasing by 15% and 210% of the control values, respectively. Assays conducted with substrate 2 (DADEpYLIPQQG) also showed increased PTP activities in ground squirrel tissues during hibernation. In the soluble fraction (Fig. 4a), PTP activity rose significantly during hibernation in muscle and brain (by 110% and 115%, respectively). In the insoluble fraction (Fig. 4b), PTP activity was increased during hibernation in three tissues; a 2.8-fold increase in skeletal muscle, a 28% increase in brain, and a 67% increase in liver were noted. Activity in heart was unchanged. Purification and properties of PP1c from ground squirrel liver PP1c from liver of euthermic ground squirrels was purified 3700-fold with a final yield of 14% (Table 3). The final specific activity of the purified enzyme was 40,588 U/mg protein, and a homogeneous preparation was confirmed by a single protein band seen on SDS– PAGE after silver staining (Fig. 5). The SDS–PAGE gel of the purified enzyme also indicated a molecular weight of 37 kDa, which was consistent with the molecular weight of 37 ± 1 kDa (n = 3) determined by the Sephacryl S-200

Fig. 4. Effect of hibernation on the activities of protein tyrosine phosphatases (PTPs) against ENDpYINASL or DADEpYLIPQQG substrate peptides in (a) soluble and (b) insoluble (membrane-associated, Triton-extracted) fractions of tissues from control (white bars) and torpid (black bars) Richardson’s ground squirrels. Data are U/mg protein, means ± SEM, n = 5 independent determinations, one PTP Unit (U) being defined as the amount of enzyme that catalyzes the release of 1 nmol phosphate per minute at 23 C. *Significantly different from the corresponding control value by the Student’s t-test, P < 0.05.

gel filtration step of the purification. These data confirm that the purified PP1c was a monomer and was absent of any associated regulatory proteins. The purified enzyme was most likely PP1ca as it was the dominant isoform detected by Western blotting in ground squirrel liver. Purified ground squirrel PP1c was not activated by divalent cations Ca2+, Mn2+, or Mg2+ (all chloride salts) over a

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Table 3 Purification of PP1c from ground squirrel liver Step

Total protein (mg)

Total activity (U)

Specific activity (U/mg)

Fold-purification

% Yield

Crude PEG precipitation DEAE-cellulose Microcystin–agarose Blue dextran Sephacryl S-200

4677 3487 429 0.99 0.63 0.17

50,700 81,500 34,200 21,500 17,000 6900

11 23 80 21,717 26,984 40,588

1 2 7 1974 2453 3690

100 161 67 42 33 14

PP1c activity was measured at 23 C against 32P-labelled GPa in 20 mM imidazole–HCl, 0.1 mM EGTA, 15 mM b-mercaptoethanol. PP1c activities are expressed in Units (U) where 1 U is 1 nmol of 32P-phosphate released per min.

Table 4 Temperature effects on nucleotide inhibition of purified ground squirrel PP1c I50 (mM) ATP ADP AMP IMP

5 C

37 C

6.3 ± 0.3 12.1 ± 1.6 >25 7.5 ± 0.5

0.32 ± 0.002* 0.44 ± 0.04* 8.2 ± 0.5* 10.3 ± 0.1*

Data are means ± SEM for n = 3 separate enzyme preparations. * Significantly different from the corresponding 5 C value, P < 0.01.

Fig. 5. Purification of PP1c from ground squirrel liver. SDS–PAGE with silver staining of samples taken from different steps in the purification. Lanes are (A) molecular weight standards, (B) Microcystin–agarose, and (C) Sephacryl S-200 gel filtration.

concentration range from 0 to 10 mM. Purified PP1c was inhibited by okadaic acid with an I50 value of 65 ± 7 nM (n = 3), and the enzyme showed 100% inhibition in the presence of 1 lM okadaic acid. This is reasonable indication that the purified phosphatase was not contaminated by PP2A or PP2C. Okadaic acid has little inhibitory activity against PP2B (calcineurin) or other Ser/Thr phosphatases (PP4, PP5, PP6, and PP7) and virtually no effect on PP2C or protein tyrosine phosphatases [19]. Purified ground squirrel PP1c was characterized with respect to inhibition by nucleotides at both hibernating and euthermic body temperatures (Table 4). All inhibition constants were significantly higher at the lower assay temperature. Purified PP1c was potently inhibited by ATP and ADP at 37 C, with I50 values of 0.32 ± 0.02 mM and 0.44 ± 0.04 mM, respectively. The two adenylates were also inhibitory at 5 C but I50 values were over 20-fold

higher. IMP was inhibitory but at non-physiological levels (I50 = 10.3 mM at 37 C and 7.5 mM at 5 C). Ground squirrel PP1c was only weakly inhibited by AMP at either temperature. The pH curves for ground squirrel PP1c assayed at euthermic and hibernating body temperatures are shown in Fig. 6. The pH optimum at 37 C was quite narrow and centered at pH 6.9 whereas the optimum pH range at 5 C was broader, stretching from pH 5.8 to 7.5. The effect of temperature on ground squirrel PP1c activity is shown as an Arrhenius plot in Fig. 7. A distinct break in

Fig. 6. The effect of pH on purified ground squirrel liver PP1c activity at 5 C (s) and 37 C (d). Data are means ± SEM for n = 3 separate enzyme preparations. PP1c activity is expressed relative to the maximum activity at the optimal pH.

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Fig. 7. Arrhenius plot of ground squirrel PP1c activity versus reciprocal temperature (K). PP1c activity was measured with GPa as the substrate in 20 mM imidazole–HCl, 0.1 mM EGTA, 15 mM b-mercaptoethanol. The pH was defined to be 7.2 at 23 C and then allowed to self-adjust with changing temperature. The assay mixtures were pre-incubated for 15 min to allow temperature equilibration before initiating the reaction with the addition of purified PP1c. Data are means ± SEM for n = 3 separate enzyme preparations; where absent, SEM bars are contained within the symbol.

the plot was seen at 9 C. The Ea for the reaction above 9 C was 59 ± 2 kJ/mol and below 9 C was 2.4-fold higher at 144 ± 5 kJ/mol. Discussion Coordinated regulation of glucose and lipid homeostasis occurs during mammalian hibernation [11–13]. Catabolism during hibernation is reorganized so that most organs depend on aerobic lipid oxidation for their energy needs. The primary mechanism responsible for carbohydrate sparing is reversible protein phosphorylation. Protein phosphatases are the cellular antagonists of protein kinases, and they match their protein kinase counterparts in specificity and complexity of signaling. Our results demonstrate that tissue-specific changes in type-1 (PP1), type-2 (PP2A, PP2C), and tyrosine (PTP) phosphatase activities are associated with hibernation. These data suggest the importance of both serine/threonine- and tyrosine protein phosphatases in the metabolic reorganization that is required to support torpor. During hibernation, even though animals stop eating, glycogen phosphorylase is suppressed to spare carbohydrate reserves as opposed to starvation where glycogen is rapidly depleted. Liver glycogen phosphorylase is activated via phosphorylation by cAMP-dependent protein kinase (PKA) following increases in [cAMP] and by allosteric effectors. The decrease in glycogen phosphorylase activity observed in liver during hibernation could occur via three mechanisms: (1) by actively increasing the dephosphorylation potential (i.e., PP1 activity) relative to the phosphory-

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lation potential (i.e., PKA activity), (2) by actively suppressing the phosphorylation potential relative to unchanging dephosphorylation potential, or (3) by altering the levels of allosteric substances that accelerate (modulate) the conversion of glycogen phosphorylase a to the inactive b form (i.e., caffeine, glucose, and AMP). Liver cAMP levels were unchanged in hibernation as was the percentage of PKA present as the free catalytic form [16]. Thus, the changes in liver PP1 activity observed in this study (Table 1) are consistent with altered dephosphorylation potential regulating glycogen metabolism in hibernation. The active forms of PP1 are generally associated with various subcellular structures. The specificity in localization is mediated by targeting subunits that direct PP1c to particular locations, enhance its activity toward certain substrates and confer important regulatory properties upon it [37]. This concept has been best established for the glycogen-bound enzymes in skeletal muscle (PP1GM) and liver (PP1GL). The majority of PP1c in euthermic ground squirrel liver was found in the cytosol with a minor amount associated with the glycogen fraction. The small amount of total cellular PP1c associated with glycogen in the euthermic state is not unexpected as the total cellular levels are quite high, and only a small subset of the total cellular complement of PP1c is believed to be associated with the glycogen particle. This glycogen-associated PP1c could promote glycogen storage in the pre-hibernating state. Generally, glycogen-associated PP1 activity (as the PP1GL holoenzyme) has greatly enhanced selectivity towards glycogen synthase while activity towards glycogen phosphorylase is suppressed [21]. The reduction in glycogen-associated PP1c hibernation may halt both the synthesis and degradation of glycogen in the hypometabolic state. This finding is also consistent with altered PP1 dephosphorylation potential contributing to a reduction in glycogenolysis and the preservation of carbohydrate reserves during torpor. Ground squirrel PP1c purified from liver showed the characteristics of a type-1 protein phosphatase and lacked properties that are typical of type-2 phosphatases such as activation by cations and inhibition by okadaic acid at concentrations less than 1 nM. Ground squirrel PP1c was moderately inhibited by okadaic acid with an I50 value of 65 nM; this value is similar to that reported for the mammalian enzyme from other sources (10–20 nM) [19]. Enzyme properties of the purified PP1c were examined at temperatures characteristic of euthermia (37 C) and hibernation (5 C). Temperature-dependent mechanisms of enzyme control are believed to contribute to the regulation of metabolic rate and other key cellular processes as the body temperature of hibernators can drop by 30 C or more [10,16]. Our analysis of the pH dependency of PP1c activity indicated that PP1c would retain its full catalytic potential over a wide range of intracellular pH values in torpid tissue. The data suggest that PP1c activity would not be affected by conditions of relative acidosis that exist in the torpid state [38]. In addition, our examination of

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temperature effects on PP1c activity (i.e., Arrhenius plots) indicates that torpor would not promote an accumulation of phosphorylated proteins in the hibernating animal since the temperature effects on protein phosphatases mimic those for protein kinases [16]. Thus, PP1c appears to be no more susceptible to temperature and/or pH change than some of its counterpart protein kinases. The data presented herein also indicate that ground squirrel PP1c is less likely to be affected by changing energetic status in hibernation. The sensitivity to adenylates (inhibition by ATP, ADP, and AMP) was reduced when ground squirrel PP1c was assayed at hibernating temperatures. In typical mammalian systems, AMP binds to glycogen phosphorylase a and prevents its dephosphorylation by PP1 [39]. Substances that accelerate the conversion of phosphorylase a to b (i.e., glucose and caffeine) abolish the inhibitory effects of AMP. Interestingly, the phosphorylase phosphatase activity of ground squirrel PP1c was only affected by high non-physiological levels of AMP, and this rather weak inhibition was unaffected by either caffeine or theophylline (data not shown). It would appear then that, at physiological [AMP], ground squirrel PP1c can dephosphorylate glycogen phosphorylase a whether or not AMP is present since physiological levels of AMP are not high enough to inhibit the process. The data suggest a unique relationship between the two proteins that limits AMP inhibition and should therefore aid in the conservation of liver glycogen stores during hibernation. Hallenbeck and colleagues [40] have previously shown a reduction in PP2A activity during hibernation in thirteenlined ground squirrels. In their study, PP2A activity was significantly reduced in both hibernating brain and liver as compared to active controls. Furthermore, the decrease in PP2A activity was believed to result from increased expression of the inhibitor of PP2A (I2PP2A) because increases in the level of the catalytic subunit of PP2A were also observed. We found a similar decrease in brain PP2A activity during hibernation, but changes in liver PP2A activity were not observed. The activities of PP2A and PP2C can be modulated by extracellular signals and can thereby modulate signal transduction pathways via the SAPK cascades [25,26,41,42] and the AMP-activated protein kinase (AMPK) cascade [43,44]. PP2A and PP2C have been shown to inactivate the AMPK cascade in vitro; however, in isolated hepatocytes, okadaic acid had no effect on the dephosphorylation of acetyl-CoA carboxylase, implying that PP2C is responsible for dephosphorylation of AMPK in liver [42]. The AMPK cascade has been described as a cellular fuel gauge that is activated by cellular stresses that deplete ATP [45]. By detecting low fuel situations AMPK protects cells by switching off ATP-consuming pathways (e.g. fatty acid synthesis and sterol synthesis) and switching on alternative pathways for ATP generation (e.g. fatty acid oxidation). A recent study of the AMPK cascade in hibernating ground squirrels found a significant decrease in AMPK activity in torpid liver [15]. AMPK exerts the major control over fatty acid

synthesis, and suppression of the cascade by increased PP2C activity during hibernation is not unexpected since the animals are not synthesizing fatty acids. In conclusion, the present study demonstrates that alterations in the activities of ground squirrel protein phosphatases occur in response to hibernation in tissue-specific manners. Changes in protein phosphatase activities, in concert with changes in protein kinase activities, suggest the relative importance of these modules in controlling metabolic function and cellular processes during hibernation. Furthermore, our analysis of PP1c suggests that the ground squirrel enzyme is capable of regulating cellular processes at low temperatures. Acknowledgments Appreciation is extended to M. de la Roche for assistance with tissue collection and J.M Storey for editorial assistance. Supported by a research grant (6793) from the Natural Sciences and Engineering Research Council of Canada (NSERC). K.B.S. is recipient of a Canada Research Chair (Tier I). J.A.M. is recipient of a Canada Research Chair (Tier II) and a Scholarship from the Heart & Stroke Foundation of Canada. References [1] L.C.H. Wang, T.F. Lee, in: M.J. Fregley, C.M. Blatteis (Eds.), Handbook of Physiology: Environmental Physiology, Oxford University Press, New York, 1996, pp. 507–532. [2] F. Geiser, J. Comp. Physiol. [B] 158 (1988) 25–37. [3] K.B. Storey, Comp. Biochem. Physiol. A Physiol. 118 (1997) 1115– 1124. [4] K.L. Drew, C.L. Buck, B.M. Barnes, S.L. Christian, B.T. Rasley, M.B. Harris, J Neurochem. (2007), doi:10.1111/j.14714159.2007.04675.x, Epub ahead of print. [5] P. Morin Jr., K.B. Storey, Cryobiology 53 (2006) 310–318. [6] V. Velickovska, B.P. Lloyd, S. Qureshi, F. van Breukelen, J. Comp. Physiol. [B] 175 (2005) 329–335. [7] K.B. Storey, in: R.C. Roach, P.D. Wagner, P.H. Hackett (Eds.), Advances in Experimental Medicine and Biology, Kluwer/Plenum Academic Press, New York, 2003, pp. 21–38. [8] H.V. Carey, M.T. Andrews, S.L. Martin, Physiol. Rev. 83 (2003) 1153–1181. [9] K.B. Storey, S. Afr. J. Zool. 33 (1998) 55–64. [10] K.B. Storey, J.M. Storey, in: W. Walz (Ed.), Integrative Physiology in the Proteomics and Post-Genomics Age, Humana Press, New Jersey, 2005, pp. 169–200. [11] S.P.J. Brooks, K.B. Storey, J. Comp. Physiol. [B] 162 (1992) 23–28. [12] K.B. Storey, J. Biol. Chem. 262 (1987) 1670–1673. [13] K.B. Storey, S.P.J. Brooks, J. Comp. Physiol. [B] 162 (1992) 23–28. [14] C.P. Holden, K.B. Storey, Arch. Biochem. Biophys. 358 (1998) 243– 250. [15] S. Horman, N. Hussain, S.M. Dilworth, K.B. Storey, M.H. Rider, Comp. Biochem. Physiol. B. Biochem. Mol. Biol. 142 (2005) 374– 382. [16] J.A. MacDonald, K.B. Storey, J. Comp. Physiol. [B] 168 (1998) 513– 525. [17] J.A. MacDonald, K.B. Storey, Int. J. Biochem. Cell Biol. 37 (2005) 679–691. [18] H. Mehrani, K.B. Storey, Neurochem. Int. 31 (1997) 139–150. [19] T. Mustelin, Methods Mol. Biol. 365 (2007) 9–22. [20] B.J. Toole, P.T. Cohen, Cell. Signal. 19 (2007) 1044–1055.

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