1

AMPKα1 deletion shortens erythrocyte lifespan in mice: Role of oxidative stress

Shaobin Wang1,3, George L. Dale2, Ping Song1, Benoit Viollet4,5,6, Ming-hui Zou1,3,

1Division of Endocrinology and Diabetes, 2Division of Hematology, Department of Medicine, 3Department of Physiology, University of Oklahoma Health Science Center, Oklahoma City, OK

73104, USA 4Institut Cochin, Paris, France

5University Paris Descartes, CNRS (UMR 8104), Paris, France

6Inserm, U567, Paris, France Address correspondence to: Ming-Hui Zou, M.D. Ph.D., BSEB 325, Section of Endocrinology and Diabetes, Department of Medicine, University of Oklahoma Health Science Center, Oklahoma City, OK 73104, USA. Phone: (405) 271-3974; Fax: (405) 271-3973; E-mail: ming-hui-zou@ouhsc.edu AMP-activated protein kinase (AMPK) is an energy sensor essential for maintaining cellular energy homeostasis. Here we report that AMPKα1 is the predominant isoform of AMPK in murine erythrocytes and mice globally deficient in AMPKα1 (AMPKα1-/-), but not those lacking AMPKα2, had markedly enlarged spleens with dramatically increased proportions of Ter119-positive erythroid cells. Blood tests revealed significantly decreased erythrocyte and hemoglobin levels, with increased reticulocyte count and elevated plasma erythropoietin concentration, in AMPKα1-/- mice. The lifespan of erythrocytes from AMPKα1-/- mice was less than that in wild-type littermates, and the levels of reactive oxygen species (ROS) and oxidized proteins were significantly increased in AMPKα1-/- erythrocytes. In keeping with the elevated oxidative stress, treatment of AMPKα1-/- mice with the antioxidant, tempol, resulted in decreased reticulocyte counts and improved erythrocyte survival. Furthermore, the expression of Foxo3a and ROS scavenging enzymes was significantly decreased in erythroblasts from AMPKα1-/- mice. Collectively, these results establish an essential role for AMPKα1 in regulating oxidative stress and lifespan in erythrocytes.

INTRODUCTION

Reactive oxygen species (ROS) are ubiquitously generated in living cells, including erythrocytes, and overwhelming evidence indicates that they play key roles in essential cellular functions. Oxidative stress, defined as a pathologic state characterized by increased ROS production or decreased ability to detoxify ROS, plays a causative role in tissue injury in many disease conditions, including cardiovascular diseases, neurological disorders, cancers, and aging.(1,2) Mature erythrocytes, with their absence of protein synthesis and high oxygen carrying capacity, are particularly susceptible to oxidative damage because they are rich in heme iron and oxygen, which can spontaneously generate H2O2 and lipid peroxides(3). Under normal conditions, red blood cells (RBCs) have effective defense mechanisms against oxidative stress. When the production of ROS overwhelms erythrocyte antioxidant systems, or antioxidant systems becomes defective, excessive ROS cause oxidative damage to the cytoplasmic membrane and associated cytoskeleton, in the mature red cell. The consequences of oxidative damage in erythrocytes are often manifested as decreased deformability and splenic sequestration. Uptake of abnormal red cells in the spleen results in a decreased lifespan of RBCs leading to anemia(4). Thus, disruption of this oxidant–antioxidant equilibrium can markedly shorten the lifespan of erythrocytes, as reported in mice

http://www.jbc.org/cgi/doi/10.1074/jbc.M110.102467The latest version is at JBC Papers in Press. Published on April 14, 2010 as Manuscript M110.102467

Copyright 2010 by The American Society for Biochemistry and Molecular Biology, Inc.

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

2

deficient for peroxiredoxin I(5), peroxiredoxin II(6), SOD1(7), or SOD2(8). Defects in these enzymes, which are critical to the oxidative stress response, have also been implicated in human diseases that involve acute or chronic hemolysis(9). The enzymopathy most commonly responsible for hemolytic anemia is deficiency of glucose-6-phosphate dehydrogenase (G6PD), which converts NADP to NADPH. NADPH is required for maintenance of glutathione (GSH), a major component of the cellular ROS detoxification system. The absence of these antioxidant protections can result in moderate hemolysis(10); thus, a normal erythrocyte lifespan depends on an adequate antioxidant response(11).

AMP-activated protein kinase (AMPK)(12) is a serine/threonine kinase, conserved from yeast to humans, that regulates energy homeostasis and metabolic stress. Mammalian AMPK is a heterotrimer consisting of three subunits designated α, β, and γ. The α subunit contains a kinase domain, and can exist as either an α1 or α2 isoform. AMPK is activated under conditions that elevate the AMP/ATP ratio, such as glucose deprivation, hypoxia, and muscle contraction(13). Recent evidence also suggests that AMPK may have a much wider range of functions. For instance, it is involved in the regulation of mitochondrial biogenesis(14) and angiogenesis(15). We previously demonstrated that pathologically relevant concentrations of peroxynitrite (ONOO-), a potent oxidant formed by the reaction of nitric oxide and superoxide anions (O2•-), are capable of activating AMPK independently of changes in the AMP/ATP ratio(16), and further, that the activation of AMPK can inhibit intracellular O2•- production in human umbilical vein endothelial cells (HUVECs)(17). Overall, AMPK appears to be essential in maintaining redox homeostasis. Whether AMPK controls oxidative homeostasis in erythrocytes and is therefore critical for their survival, is not clear. Here, we report that AMPKα1 depletion triggers oxidative stress and reduces the lifespan of erythrocytes by

lowering the expression of Foxo3a-mediated antioxidant enzymes.

EXPERIMENTAL PROCEDURES

Reagents and animals

Rabbit anti-AMPKα antibody was obtained from Cell Signaling Technology (Beverly, MA). Rabbit anti-AMPKα1 and Rabbit anit-AMPKα2 antibodies were obtained from Bethyl Laboratories, Inc. (Montgomery, TX). Mouse anti-GAPDH antibody was from Abcam Inc. (Cambridge, MA). Other chemicals and organic solvents of the highest available grade were obtained from Sigma-Aldrich. AMPKα1-/- mice and AMPKα2-/- mice were described elsewhere(18,19). Tempol (4-hydroxy-2,2,6,6-tetramethylpiperidinyloxy) was provided to the animals for the indicated times (see Results) at a concentration of 1 mM in drinking water. Mice were handled in accordance with study protocols approved by the Institutional Animal Care and Use Committee of the University of Oklahoma Health Science Center, Oklahoma.

Western blot Freshly heparinized blood was collected from wild type, AMPKα1-/- and AMPKα2-/- mice. Blood samples were centrifuged at 1,000 g for 10 min. The plasma and buffy coat were removed by aspiration. The pellets were suspended in 3 volumes of phosphate-buffered saline (PBS), pH 7.4, and then passed through a column of α-cellulose and microcrystalline cellulose (1:l, wt/wt) to remove platelets and white blood cells (WBC), as described (20). Washed RBCs were then collected, and lysed in 25 mM Tris•HCl pH 7.6, 150 mM NaCl, 1% NP-40, 1% sodium deoxycholate, 0.1% SDS, supplemented with 0.25 mg/ml phenylmethylsulfonylfluoride, 1× protease inhibitor, and 1× phosphatase inhibitor cocktails (Calbiochem) (0.15 ml lysis solution per 1 × 107 cells). An equal volume of 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, 112.5 mM NaCl, 37.5 mM Tris-HCl (pH 7.4) was added, and cleared extracts were denatured,

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

3

electrophoresed (25 µg), and blotted. For chemiluminescence, HRP-conjugated secondary antibodies (Jackson ImmunoResearch Laboratories Inc.) and SuperSignal West Dura reagent (Pierce) were used. Histological and hematological analyses

10-12 weeks male and female mice were weighed and sacrificed. Selected organs, including spleen, liver, kidney, heart, and femur, were removed and weighed to calculate an organ index (organ weight/body weight x 100). Tissues processed for microscopic evaluation were fixed in 10% neutral-buffered formalin, embedded in paraffin, sectioned, mounted on slides, and stained with hematoxylin and eosin (H&E) or Prussian blue. Bones were decalcified before fixing in formalin. Tissue sections were examined with an Olympus AX70 microscope with UPlanFI 20x/0.50 lens or 40x/0.85 lens, blood smears with oil-immersion UPlanApO 100x/1.30 lens, and an Qimaging Retiga Exi digital CCD camera

(QImaging Co.) using QCapture software (v. 2.98.0). Peripheral blood samples were harvested in EDTA coated micro tubes (Aktiengesellschaft Co. Germany) by retro-orbital sinus bleeding and analyzed with a HEMAVET 950 Veterinary Hematology Analyzer (Drew Scientific Inc., TX) using mouse species program. Blood smears were stained with Giemsa or new methylene blue, and analyzed microscopically. Flow cytometry analysis

Mice spleens were mechanically dissociated through a 70-µm strainer and washed with cold PBS containing 2% fetal calf serum. Splenocyte single-cell suspensions were double stained with antibodies against fluorescein isothiocyanate-conjugated CD71 (CD71-FITC) and phycoerythrin-conjugated erythroid antigen (Ter119-PE; BD Bioscience, CA). Flow cytometry was performed using a FACSCalibur (BD Bioscience, CA), and FACS data was analyzed using Summit v4.3 software. Reticulocytes were assayed using a Retic-

Count Reticulocyte Reagent System (BD Biosciences, CA). Erythrocyte lifespan determination

In vivo biotinylation followed by FACS analysis were performed according to Suzuki and Dale with modifications(21,22). Briefly, RBCs were labeled in vivo by intravenous injection of 30 mg/kg NHS-LC-biotin (N-succinimidyl-6-[biotinamido] hexanoate; Thermo Fisher Scientific, IL). Small samples (5µl) of peripheral blood were collected from tail vein one hour after biotin labeling and stained using streptavidin-PE. Staining was followed by FACS analysis to ensure that at least 95% of red cells were labeled with biotin. Blood samples were analyzed at 7-day intervals to quantify biotin-labeled cells remaining in the circulation. Erythropoietin (EPO) ELISA. EPO level in blood plasma was determined using the Quantikine Mouse EPO ELISA kit from R&D Systems, Inc. ( Minneapolis, MN) according to manufacturer’s protocol. Erythrocyte reinfusion and clearance test

In vivo RBC tracking was performed as reported(23,24) with modification. Briefly, RBCs of donor mice were biotin-labeled in vivo by tail vein injection of NHS-LC-biotin, as described above. One hour after biotin infusion, blood was collected from donor mice in tubes containing heparin as an anticoagulant. A small aliquot of labeled RBCs was incubated with streptavidin-phycoerythrin and

analyzed by flow cytometry to assure that at least 95% of the blood cells were labeled. Blood was washed and resuspended in sterile saline, and 100 µl of RBC suspension was infused into each recipient mouse by tail vein injection. Initial postinfusion blood samples were obtained after 30 minutes and analyzed by flow cytometry. The typical initial percentage of biotin-labeled RBCs (recorded as Day 0 %) was greater than 5% in recipient mice. Seven days after blood reinfusion, the remaining labeled red cells were determined (recorded as Day 7 %). The

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

4

red cell clearance rate was calculated as: Clearance rate = ([Day 7 % - Day 0 %]/Day 0 %) x 100%. Protein oxidation detection

The OxyblotTM protein oxidation detection kit (Millipore, MA), based on a previously described method(25), was used as described by the manufacturer. Briefly, RBCs lysates from wild-type mice and AMPKα1-/- mice were derivatized with dinitro phenyl hydrazine (DNP), and derivatized samples (20 µg protein/sample) were separated by SDS-PAGE on 12% gels. Proteins were transferred to nitrocellulose membranes, and membranes were blocked and immunoblotted using OxyBlot Kit methods and reagents. Bands were visualized with chemiluminescence (captured on film) and analyzed densitometrically. Membranes stained with Red Ponceau S (Sigma Chemical) were used as loading controls.

Intracellular ROS measurement

Red cell intracellular ROS measurement was performed essentially as reported(8). RBCs were washed and resuspended in staining buffer (BD Bioscience, CA) and loaded with 5 µM CM-H2DCFDA (Invitrogen) in the dark for 30 minutes at 37oC in a 5% CO2 atmosphere. Intracellular fluorescent products were measured immediately by flow cytometry. Osmotic fragility assay The osmotic fragility of erythrocytes in freshly collected blood from AMPKα1-/- and wild-type mice was measured with modification (26). Briefly, RBCs were harvested into heparin coated tubes and suspended in varying concentrations of NaCl, incubating samples at room temperature for 10 min, and then centrifuging at 1500 × g for 10 minutes to sediment unlysed cells and stroma, 200 µl supernatant was collected, the hemoglobin concentration of each sample was measured by Synergy HT Multi-Detection microplate reader (BioTek Instrument, Inc) at 540 nm , with appropriate controls. The lyses

percentage of RBCs was calculated from the absorbance, and a fragility curve was generated by plotting varying salt concentrations versus hemolysis. Fluorescence-activated cell sorting (FACS) and quantitative reverse transcriptase-polymerase chain reaction (RT-PCR)

Bone marrow cell suspensions were stained with antibodies against CD71-FITC and Ter119-PE (BD bioscience, CA) before sorting on an Influx Cell Sorter. Total RNA in sorted cells was extracted using an RNeasy Mini Kit (Qiagen), and cDNA was synthesized from total RNA using an iScript cDNA Synthesis Kit (BioRad, CA), as described by the respective manufacturers. Real-time quantitative RT-PCR was performed using iQ SYBR Green SuperMix and the MyiQ Single-Color Real-Time PCR Detection System (BioRad, CA). Primer sequences are presented in Supplemental Data. Quantitative RT-PCR results were analyzed according to the 2-∆∆ct method(27) using expression of the β-actin gene as a housekeeping control. Statistical Analysis Two-tail Student's t-test was conducted or two-way ANOVA followed by Bonferroni post hoc analyses was used to determine statistical differences between various experimental and control groups. P-value < 0.05 was considered statistically significant.

RESULTS: AMPKα1 is the predominant isoform in murine erythrocytes.

We found that AMPKα1 is the predominant isoform of AMPKα in murine erythrocytes, as demonstrated by western blots. Using an antibody specific for AMPKα1, we confirmed the expression of AMPKα1 in mice RBCs (Figure 1A). Because no recognizable bands were seen on the blot with the same loading with an antibody specific for AMPKα2, we proceeded to use an antibody that recognizes both AMPKα1 and AMPKα2 isoforms. As

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

5

shown in Figure 1A, AMPKα was largely absent in erythrocytes from AMPKα1-/- mice, whereas there was no significant difference of the expression of total AMPKα between AMPKα2-/- and wild-type erythocytes. Genetic deletion of AMPKα1, but not AMPKα2, in mice causes splenomegaly and moderate anemia.

Next, we determined if genetic deletion of AMPKα1 or AMPKα2 affected the pathology of organs, including hearts, kidneys, and spleens. AMPKα1-/- mice exhibited splenomegaly (Figure 1B and 1C), whereas body weights and the weights of other major organs were not significantly changed (Supplemental Figure 1); this phenotype was not observed in AMPKα2-/- mice. Hematological tests showed moderate anemia in AMPKα1-/- mice, as evidence by low erythrocyte number, low hemoglobin content, and increased red cell distribution width (RDW) compared with wild-type mice. Leukocyte and platelet counts were not significantly changed in AMPKα1-/- mice compared with wild-type literate mates (Table 1). In AMPKα2-/- mice, there were no significant differences in erythrocyte or platelet parameters compared with wild-type mice (Supplemental Table 1). Accordingly, subsequent experiments focused on AMPKα1-/- mice. H&E-stained sections of spleens from AMPKα1-/- mice revealed expanded red pulps and an increase in erythroid precursor cells (Figure 1D), indicating a compensatory reaction to anemia. Prussian blue staining showed elevated iron deposits in spleen sections from AMPKα1-/- mice (Figure 1E), indicating increased destruction of erythrocytes in the spleen, and Giemsa staining of blood smears indicated polychromasia, combined macrocytosis and microcytosis, and reticulocytosis in AMPKα1-/- mice (Figure 1F). Collectively, these data indicate that a deficiency of AMPKα1 caused moderate anemia, characterized by a significant decrease in red cell count, hematocrit, and hemoglobin in peripheral blood relative to wild-type

littermate controls. The phenotypic differences between AMPKα1-/- mice and AMPKα2-/- mice likely stem from the distinctively different expression levels of these two isoforms in erythrocytes. Erythropoiesis is increased in AMPKα1-/- mice.

The anemia observed in AMPKα1-/- mice could be due to defective red cell maturation (ineffective erythropoiesis), increased red cell destruction (hemolysis), or a combination of both processes. We first analyzed splenocytes by flow cytometry. Erythroid cells (Ter119+) in the spleens of AMPKα1-/- mice were increased compared to those of wild-type mice (Supplemental Figure 2A), whereas the relative abundance of other cell types in the spleen was not increased (Supplemental Figure 2 B-E). A subpopulation of Ter119high cells was distinguished based on their expression of the transferrin receptor (CD71), which decreases with erythroblast maturation. Combining Ter119 and CD71 expression distinguishes four subpopulations of erythroid cells -- Ter119medCD71high, Ter119highCD71high, Ter119highCD71med, and Ter119highCD71low corresponding to increasingly mature erythroblasts(28). Using this approach, we found a dramatic expansion of the early erythroblast population (Ter119highCD71high) and increased proerythroblast population (Ter119mediumCD71high) in AMPKα1-/- mice (Figure 2A). Further analysis the Ter119high erythroblasts arbitrarily based on the forward scatter (FSC) parameter and CD71 expression level. Ter119high erythroblasts can be subdivided to three groups: Ery.A (Ter119highCD71highFSChigh) are basophilic, Ery.B (Ter119highCD71highFSClow) are late basophilic and polychromatic, and Ery.C (Ter119highCD71lowFSClow) are orthochromatic erythroblasts and reticulocytes increasingly mature erythroblasts according to the reports(29,30) indicated that the major elevated population was late basophilc and polychromatic cells (Supplementary Figure 4).

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

6

Next, using new methylene blue-stained blood smears (data not shown) and flow cytometry with thiazole orange staining (Figure 3A), we determined the number of reticulocytes, and found that the reticulocyte count in AMPKα1-/- mice was 3-4-fold higher than that in wild-type mice (Figure 3B). The increased reticulocyte levels were correlated with elevated plasma levels of erythropoietin (EPO) (Figure 3C). These data indicate that erythropoiesis was elevated in AMPKα1-/- mice, possibly reflecting a compensatory reaction to the moderate anemia observed in these mice. Erythrocyte lifespan is shortened in AMPKα1-/- mice.

The normal lifespan of circulating RBCs, which is determined by their clearance from the peripheral circulation (predominantly by the spleen), has between reported to be approximately 120 days and 40 days in humans and mice, respectively(31). To determine more directly whether the observed anemia was due to decreased production of mature RBCs or increased destruction of RBC in the circulation, we measured erythrocyte lifespan using direct in vivo biotin labeling. To this end, wild-type and AMPKα1-/- mice were injected with NHS-LC-biotin to label RBCs, and cell lifespan was determined by flow cytometric analysis of circulating, biotinylated RBCs. The time required for 50% of labeled erythrocytes to be lost from wild-type mice (half-life) was about 24 days, consistent with previous reports(5,22). However, the half-life of labeled erythrocytes decreased to about 14 days in AMPKα1-/- mice (Figure 4A).

To further demonstrate the destruction of erythrocytes in AMPKα1-/- mice, we performed a blood cross-transfusion experiment. Wild-type mice received either biotin-labeled wild-type erythrocytes (WT-WT) or biotin-labeled AMPKα1-/- erythrocytes (KO-WT), and AMPKα1-/- mice received either biotin-labeled wild-type erythrocytes (WT-KO) or biotin-labeled AMPKα1-/- erythrocytes (KO-KO). The result demonstrated that the KO-

WT group had a higher clearance rate of infused erythrocytes than did the WT-WT group and similar results were observed when compared KO-KO with WT-KO group (Figure 4B), suggesting accelerated clearance of erythrocyte in AMPKα1-/- mice. Consistent with these results, Prussian blue-stained spleen sections from AMPKα1-/- mice showed a marked increase in iron deposits. Osmotic fragility is decreased in erythrocytes from AMPKα1-/- mice.

To uncover the possible cause of why erythrocytes from AMPKα1-/- mice had an increased clearance rate, we first determined the osmotic fragility of erythrocytes. As depicted in Figure 5A, AMPKα1-/- deficient erythrocytes were significantly more resistant to hypotonic lysis than were wild-type red cells (Figure 5A), suggesting that they had a more rigid and less deformable cell membrane. Phenylhydrazine-induced mortality is increased in AMPKα1-/- mice.

Phenylhydrazine (PHZ), a potent inducer of oxidative stress in vivo and is capable of denaturing hemoglobin and hemolyzing red cells (32). To test PHZ tolerance, we injected wild-type and AMPKα1-/- mice with 100 mg/kg PHZ. As shown in Figure 5B, five of six PHZ-treated AMPKα1-/- mice died within 24 hours, and the sixth died within 48 hours after treatment. In contrast, all six wild-type mice receiving PHZ survived (Figure 5B). These results imply that AMPKα1-/- mice are hypersensitive to PHZ-induced oxidative stress. Intracellular ROS levels and protein oxidation are increased in AMPKα1-/- erythrocytes.

Next, we determined if increased levels of protein aggregates were due to increased oxidative stress in erythrocytes. Protein oxidation can be measured indirectly by western blotting for carbonyls that result from the reaction of side chains of lysine, proline, threonine, or arginine with

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

7

ROS(25),(33) Intracellular ROS concentrations measured under baseline conditions and after challenging with exogenous H2O2 (50 µM) showed that ROS levels were elevated in erythrocytes from AMPKα1-/- mice compared to wild-type mice under both conditions (Figure 6A). In agreement with the observed elevation in ROS concentrations, oxidized protein levels in erythrocytes from AMPKα1-/- mice were significantly increased (~7-fold) as measured by 2,4-dinitrophenylhydrazine (DNPH) derivatized carbonyl immunochemical staining (Figure 6B, and 6C). Chronic administration of Tempol prolongs the lifespan of erythrocytes in AMPKα1-/- mice.

To further evaluate the potential affect of ROS on shortened red cell survival, we fed tempol, a potent antioxidant, to wild-type and AMPKα1-/- mice. Tempol treatment had no effect on reticulocyte index or erythrocyte lifespan in wild-type mice. However, it significantly increased the lifespan of erythrocytes in AMPKα1-/- mice, but decreased the reticulocyte index (Figure 7A and B). 4 month after Tempol treatment, the spleen index was significantly decreased in treatment groups compared with no treatment AMPKα1-/- controls (Figure 7C).

Expression of the Foxo3a transcription factor and antioxidant genes are decreased in erythroblast cells from AMPKα1-/- mice.

Finally, we investigated the molecular basis of increased oxidative stress in the erythrocytes from AMPKα1-/- mice by measuring the mRNA levels of several antioxidant genes known to play a key role in erythrocytes. Total RNA was extracted from sorted CD71highTer119high erythroblast cells from wild-type and AMPKα1-/- mice, and analyzed by quantitative real-time RT-PCR. As shown in Figure 7D, the expression of the Foxo3a transcription factor was significantly reduced in the erythroblast cells from AMPKα1-/- mice compared to that in wild-type mice; the mRNA levels of Sod2,

catalase, and glutathione peroxidase (GPx-1) were also reduced.

DISCUSSION:

In this study, we provide the first

evidence to suggest that AMPKα1, but not AMPKα2, maintains the oxidant–antioxidant balance in erythrocytes by controlling Foxo3a transcription factor-mediated antioxidant enzyme expression. We further show that genetic deletion of AMPKα1 causes oxidative stress, resulting in a shortened erythrocyte lifespan, RBC hemolysis, and anemia in vivo. The most important find of this study we found that erythrocytes predominantly express AMPKα1 and genetic deletion AMPKα1 lead to anemia in vivo. The anemia could be caused by defective red cell maturation (ineffective erythropoiesis), increased red cell destruction (hemolysis), or a combination of both processes. Our data clearly indicate that the anemia observed in AMPKα1-/- mice is likely the result of increased erythrocyte removal by the spleen. This conclusion is supported by several lines of evidence. First, the rate at which labeled erythrocytes were eliminated from the circulation, was markedly higher in AMPKα1-/- mice than in wild-type littermates (Figure 4A). Consistent with this result, the Prussian blue-stained sections of spleen showed marked increases in iron deposits in AMPKα1-/- mice. Second, reticulocytes were increased in AMPKα1-/- mice and the increased reticulocyte levels were correlated with elevated levels of plasma EPO. In addition, Ter119highCD71high erythroblasts in the spleen were also dramatically expanded (Figure2A), and a histological examination of bone sections of AMPKα1-/- showed no significant deficiency (Supplemental Figure 3). We noticed that there was a decrease in the late erythroblast population (Ter119 high CD71 low cells ) in spleen compared AMPKα1-/- mice compared with wild type controls, meanwhile the early erythroblast population (Ter119 medium CD71 high and Ter119 high CD71 high) was increased (Figure

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

8

2A,2B). The result suggested the maturation of Ter119 highCD71high stage to Ter119 high CD71 low stage is the limiting step for anemia compensation in AMPKα1-/- mice and that the enhanced loss of abnormally fragile RBCs from the body, exceeds the rate of recruitment of the late erythroblast population in spleen. Furthermore, the increased reticulocyte count accompanied by anemia, is highly suggestive of peripheral hemolysis in these animals. In line with our observations, Foller et al have reported genetic deletion AMPKα1 lead to anemia (34). Importantly, our results suggest that elevated intracellular ROS level play important role for the shorted lifespan of red cells, which supported by our FACS data, protein oxidation immunoblot results, Phenyl hydrazine (PHZ) treatment result and more importantly, our results showed that in vivo treatment of the mice with the antioxidant can significantly, even not completely, reverse the phenotype. Taken together, these results suggest that the shortened erythrocyte lifespan in AMPKα1-/- mice is due to elevated destruction of circulating erythrocytes within the spleen due to factors intrinsic to the erythrocytes.

The specialized structure of the venous system of the red pulp gives the spleen a unique capacity to remove abnormal erythrocytes. RBCs must cross the narrow interendothelial slits in walls of the venous sinuses in order to reenter the venous system, a process that requires erythrocytes to undergo remarkable deformation(35,36). Alterations in membrane characteristics can affect cell survival. Perhaps not surprisingly, abnormal structure and deformability of the erythrocyte membrane is observed in many clinical red cell disorders(37), and reduced erythrocyte deformability plays an important role in shortened erythrocyte survival in many types of hemolytic anemia(38). Consistent with this interpretation, we found that the erythrocytes from AMPKα1-/- mice had markedly decreased osmotic fragility, an indication of increased rigidity of erythrocytes.

The clearance of abnormal RBC occurs primarily in spleen by two methods: first, the rigid red cells would be trapped by specific venous sinuses structure of red pulps. Second, the macrophage in the spleen would recognize the specific surface markers such as phosphatidylserine (PS) exposure or CD 47 on the abnormal or foreign infused RBCs (39,40). Unlike the data shown in Figure 4A, the KO-KO combination did not have higher clearance rate than that in WT-WT. This apparent conflict between Figure 4A and 4B can be reconciled by abnormal macrophage function in spleen to recognize infused RBCs. Indeed, macrophages isolated from AMPKα1 knockout mice had significantly decreased ability to recognize and phagocytes fluorescence-labeled oxidized-LDL in vivo (Zhang M., unpublished data).

Due to low metabolic rate in red cells, these cells are relatively not sensitive to nutrient depletion but are very sensitive to oxidative stress because of the high physiological hemoglobin level (9,41).The deformability of erythrocytes is also affected by oxidative stress (42-44). Excessive ROS in erythrocytes causes damage to the cytoplasmic membrane and associated cytoskeleton in the mature red cell, effects that manifest as decreased deformability of RBCs and splenic sequestration. In this study, we found increased oxidative damage in erythrocytes from AMPKα1-/- mice, as evidenced by increased ROS levels, increased protein oxidation , decreased RBC survival, and decreased osmotic fragility. This phenotype is akin to that observed in mice deficient for the antioxidant enzymes, peroxiredoxin I (5), peroxiredoxin II(6) , Sod1(7), or Sod2(8). That the antioxidant defenses in AMPKα1-/- mice are defective is also supported by PHZ-tolerance experiments, which showed that AMPKα1-/- mice were hypersensitive to PHZ-induced oxidative stress compared to wild-type littermates. Most importantly, we found that chronic administration of the antioxidant, tempol, markedly prolonged the

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

9

lifespan of erythrocytes and ameliorated anemia in AMPKα1-/- mice. Foller et al have reported genetic deletion AMPKα1 lead to anemia (34).

The specialized structure of the venous system of the red pulp gives the spleen a unique capacity to remove abnormal erythrocytes. RBCs must cross the narrow interendothelial slits in walls of the venous sinuses in order to reenter the venous system, a process that requires erythrocytes to undergo remarkable deformation(35,36). Alterations in membrane characteristics can affect cell survival. Perhaps not surprisingly, abnormal structure and deformability of the erythrocyte membrane is observed in many clinical red cell disorders(37), and reduced erythrocyte deformability plays an important role in shortened erythrocyte survival in many types of hemolytic anemia(38). Consistent with this interpretation, we found that the erythrocytes from AMPKα1-/- mice had markedly decreased osmotic fragility, an indication of increased rigidity of erythrocytes.

The deformability of erythrocytes is also affected by oxidative stress (42-44). Excessive ROS in erythrocytes causes damage to the cytoplasmic membrane and associated cytoskeleton in the mature red cell, effects that manifest as decreased deformability of RBCs and splenic sequestration. In this study, we found increased oxidative damage in erythrocytes from AMPKα1-/- mice, as evidenced by increased ROS levels, increased protein oxidation , decreased RBC survival, and decreased osmotic fragility. This phenotype is akin to that observed in mice deficient for the antioxidant enzymes, peroxiredoxin I (5), peroxiredoxin II(6) , Sod1(7), or Sod2(8). That the antioxidant defenses in AMPKα1-/- mice are defective is also supported by PHZ-tolerance experiments, which showed that AMPKα1-/- mice were hypersensitive to PHZ-induced oxidative stress compared to wild-type littermates. Most importantly, we found that chronic administration of the antioxidant, tempol, markedly prolonged the lifespan of

erythrocytes and ameliorated anemia in AMPKα1-/- mice. In supporting this concept, our published studies have demonstrated that AMPKα2 deletion causes excessive oxidative stress and atherosclerosis in vivo (17,45). Taken together, our results strongly imply that AMPKα1 deletion increases the removal of erythrocytes in vivo, likely via increased levels of intracellular ROS.

Another important finding of this study is that Foxo3 might be an important target of AMPK in erythrocytes and AMPK-FOXO-3 axis is essential in maintaining redox homeostasis. Earlier study from Tothova and colleagues (46) has reported that significant elevation of ROS in the Foxo3-deletion hematopoietic stem cell population, as compared with wild type controls, which is correlated with decreased expression of anti-oxidant genes including catalase, MnSOD GADD45 et al. In the study carried out by Marinkovic D. et al (47), they found that animals deficient in the Foxo3 have shortened lifespan of erythrocyte together with decreased expression of ROS scavenging enzymes including catalase, SOD1, SOD2, and glutathione peroxidase 1 in erythroblast. Consistently, FOXO3 deficient mice died rapidly when exposed to erythroid oxidative stress-induced conditions and antioxidant treatment can significantly improve the lifespan of Foxo3-/- erythrocyte and corrected the reticulocyte index in these mice. Our observations are in line with earlier reports that FOXO3 is a downstream target of AMPK(48), and that AMPK activity, mediated at least partly by Foxo transcription factors, is required for the extension of lifespan in Caenorhabditis elegans by dietary restriction(49). During the preparation of this paper, Colombo et al(50) reported that AMPKα1 regulates the status of antioxidants in HUVECs and that Foxo3a is involved in this process. How AMPKα1 regulates Foxo3 transcription factors remains unknown and warrants further investigation.

In summary, our results demonstrate that AMPKα1 is important in regulating the expression of genes involved in

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

10

antioxidative defense in red blood cells, genetic deletion of AMPKα1 resulted in oxidant-antioxidant imbalance in RBCs, and increased intracellular ROS level caused elevated protein oxidation and increased osmotic fragility. The elimination of those

fragile erythrocytes by spleen exceeded the recruitment of RBCs leads to anemia phenotype in AMPKα1-/- mice (Figure 7E). Our research suggested that AMPK activation might be a therapeutic target for treating erythrocyte disorders.

FOOTNOTES

We thank Paul Friese for assistance with flow cytometry analyses. We are grateful to Mrs. Sima Asfa and Melissa for their excellent technical support. This work was supported in part by NIH grants (HL079584, HL074399, HL080499, HL089920, and HL096032), and by research awards from the American Diabetes Association, Juvenile Diabetes Research Foundation, Oklahoma Center for Advancement of Science and Technology, and a Travis Endowed Chair in Endocrinology, University of Oklahoma Health Sciences Center. Dr. M.H. Zou is a recipient of a National Established Investigator Award from the American Heart Association.

REFERENCES 1. Zou, M. H. (2007) Prostaglandins Other Lipid Mediat 82, 119-127 2. Castro, L., and Freeman, B. A. (2001) Nutrition 17, 161, 163-165 3. Johnson, R. M., Goyette, G., Jr., Ravindranath, Y., and Ho, Y. S. (2005) Free

Radic Biol Med 39, 1407-1417 4. Mohandas, N., and Gallagher, P. G. (2008) Blood 112, 3939-3948 5. Neumann, C. A., Krause, D. S., Carman, C. V., Das, S., Dubey, D. P.,

Abraham, J. L., Bronson, R. T., Fujiwara, Y., Orkin, S. H., and Van Etten, R. A. (2003) Nature 424, 561-565

6. Lee, T. H., Kim, S. U., Yu, S. L., Kim, S. H., Park, D. S., Moon, H. B., Dho, S. H., Kwon, K. S., Kwon, H. J., Han, Y. H., Jeong, S., Kang, S. W., Shin, H. S., Lee, K. K., Rhee, S. G., and Yu, D. Y. (2003) Blood 101, 5033-5038

7. Iuchi, Y., Okada, F., Onuma, K., Onoda, T., Asao, H., Kobayashi, M., and Fujii, J. (2007) Biochem J 402, 219-227

8. Friedman, J. S., Lopez, M. F., Fleming, M. D., Rivera, A., Martin, F. M., Welsh, M. L., Boyd, A., Doctrow, S. R., and Burakoff, S. J. (2004) Blood 104, 2565-2573

9. Cimen, M. Y. (2008) Clin Chim Acta 390, 1-11 10. McMullin, M. F. (1999) J Clin Pathol 52, 241-244 11. Tsantes, A. E., Bonovas, S., Travlou, A., and Sitaras, N. M. (2006) Antioxid

Redox Signal 8, 1205-1216 12. Hardie, D. G., Scott, J. W., Pan, D. A., and Hudson, E. R. (2003) FEBS Lett

546, 113-120 13. Kahn, B. B., Alquier, T., Carling, D., and Hardie, D. G. (2005) Cell Metab 1,

15-25 14. Winder, W. W., Holmes, B. F., Rubink, D. S., Jensen, E. B., Chen, M., and

Holloszy, J. O. (2000) J Appl Physiol 88, 2219-2226 15. Ouchi, N., Shibata, R., and Walsh, K. (2005) Circ Res 96, 838-846

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

11

16. Zou, M. H., Hou, X. Y., Shi, C. M., Kirkpatick, S., Liu, F., Goldman, M. H., and Cohen, R. A. (2003) J Biol Chem 278, 34003-34010

17. Xie, Z., Zhang, J., Wu, J., Viollet, B., and Zou, M. H. (2008) Diabetes 57, 3222-3230

18. Jorgensen, S. B., Viollet, B., Andreelli, F., Frosig, C., Birk, J. B., Schjerling, P., Vaulont, S., Richter, E. A., and Wojtaszewski, J. F. (2004) J Biol Chem 279, 1070-1079

19. Viollet, B., Andreelli, F., Jorgensen, S. B., Perrin, C., Geloen, A., Flamez, D., Mu, J., Lenzner, C., Baud, O., Bennoun, M., Gomas, E., Nicolas, G., Wojtaszewski, J. F., Kahn, A., Carling, D., Schuit, F. C., Birnbaum, M. J., Richter, E. A., Burcelin, R., and Vaulont, S. (2003) J Clin Invest 111, 91-98

20. Beutler, E., West, C., and Blume, K. G. (1976) J Lab Clin Med 88, 328-333 21. Suzuki, T., and Dale, G. L. (1987) Blood 70, 791-795 22. Hoffmann-Fezer, G., Mysliwietz, J., Mortlbauer, W., Zeitler, H. J., Eberle, E.,

Honle, U., and Thierfelder, S. (1993) Ann Hematol 67, 81-87 23. Bogdanova, A., Mihov, D., Lutz, H., Saam, B., Gassmann, M., and Vogel, J.

(2007) Blood 110, 762-769 24. Ishikawa-Sekigami, T., Kaneko, Y., Okazawa, H., Tomizawa, T., Okajo, J.,

Saito, Y., Okuzawa, C., Sugawara-Yokoo, M., Nishiyama, U., Ohnishi, H., Matozaki, T., and Nojima, Y. (2006) Blood 107, 341-348

25. Levine, R. L., Williams, J. A., Stadtman, E. R., and Shacter, E. (1994) Methods Enzymol 233, 346-357

26. Turgeon, M. L. (2005) osmotic fragility of erythrocyte: Dacie's method. in Clinical hematology: theory and procedures (Turgeon, M. L. ed.), Fourth Edition Ed., Lippincott Williams & Wilkins, Hagerstown. pp 455-458

27. Livak, K. J., and Schmittgen, T. D. (2001) Methods 25, 402-408 28. Zhang, J., Socolovsky, M., Gross, A. W., and Lodish, H. F. (2003) Blood 102,

3938-3946 29. Socolovsky, M., Nam, H., Fleming, M. D., Haase, V. H., Brugnara, C., and

Lodish, H. F. (2001) Blood 98, 3261-3273 30. Liu, Y., Pop, R., Sadegh, C., Brugnara, C., Haase, V. H., and Socolovsky, M.

(2006) Blood 108, 123-133 31. Beutler, E. (2005) Destruction of Erythroyctes. in Williams Hematology

(Marshall A. Lichtman, W. J. W., Ernest Beutler, Kenneth Kaushansky, Thomas J. Kipps, Uri Seligsohn, Josef Prchal ed.), Seventh Edition Ed., McGraw-Hill Medical, New York pp 405-410

32. Itano, H. A., Hirota, K., and Hosokawa, K. (1975) Nature 256, 665-667 33. Stadtman, E. R. (1993) Annu Rev Biochem 62, 797-821 34. Foller, M., Sopjani, M., Koka, S., Gu, S., Mahmud, H., Wang, K., Floride, E.,

Schleicher, E., Schulz, E., Munzel, T., and Lang, F. (2009) FASEB J 23, 1072-1080

35. Cesta, M. F. (2006) Toxicol Pathol 34, 455-465 36. Mebius, R. E., and Kraal, G. (2005) Nat Rev Immunol 5, 606-616 37. An, X., and Mohandas, N. (2008) Br J Haematol 141, 367-375 38. Mohandas, N., Phillips, W. M., and Bessis, M. (1979) Semin Hematol 16, 95-

114

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

12

39. Oldenborg, P. A., Zheleznyak, A., Fang, Y. F., Lagenaur, C. F., Gresham, H. D., and Lindberg, F. P. (2000) Science 288, 2051-2054

40. Bratosin, D., Mazurier, J., Tissier, J. P., Slomianny, C., Estaquier, J., Russo-Marie, F., Huart, J. J., Freyssinet, J. M., Aminoff, D., Ameisen, J. C., and Montreuil, J. (1997) C R Acad Sci III 320, 811-818

41. Fibach, E., and Rachmilewitz, E. (2008) Curr Mol Med 8, 609-619 42. Snyder, L. M., Fortier, N. L., Trainor, J., Jacobs, J., Leb, L., Lubin, B., Chiu,

D., Shohet, S., and Mohandas, N. (1985) J Clin Invest 76, 1971-1977 43. Mohandas, N., Clark, M. R., Jacobs, M. S., and Shohet, S. B. (1980) J Clin

Invest 66, 563-573 44. Hebbel, R. P., Leung, A., and Mohandas, N. (1990) Blood 76, 1015-1020 45. Wang, S., Zhang, M., Liang, B., Xu, J., Xie, Z., Liu, C., Viollet, B., Yan, D.,

and Zou, M. H. Circ Res 106, 1117-1128 46. Tothova, Z., and Gilliland, D. G. (2007) Cell Stem Cell 1, 140-152 47. Marinkovic, D., Zhang, X., Yalcin, S., Luciano, J. P., Brugnara, C., Huber,

T., and Ghaffari, S. (2007) J Clin Invest 117, 2133-2144 48. Greer, E. L., Oskoui, P. R., Banko, M. R., Maniar, J. M., Gygi, M. P., Gygi, S.

P., and Brunet, A. (2007) J Biol Chem 282, 30107-30119 49. Kops, G. J., Dansen, T. B., Polderman, P. E., Saarloos, I., Wirtz, K. W.,

Coffer, P. J., Huang, T. T., Bos, J. L., Medema, R. H., and Burgering, B. M. (2002) Nature 419, 316-321

50. Colombo, S. L., and Moncada, S. (2009) Biochem J

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

13

FIGURE LEGENDS Figure 1. AMPKα1 is the predominant isoform of AMPK expressed in mouse erythrocytes and its deletion causes splenomegaly. A: AMPKα1 and total AMPKα protein expression in erythrocytes from wild-type, AMPKα1-/-, and AMPKα2-/- mice were detected by immunoblotting. ∆ indicates non-specific bands. B: Representative picture of spleens from wild-type and AMPKα1-/- mice. C: Increased spleen index in AMPKα1-/- mice compared with wild-type mice. Data were expressed as mean ± S.D (**p < 0.01 vs. WT, n = 12 in each group). D: H&E-stained sections of spleen, showing clearly defined white pulp and red pulp in wild-type spleen, and evidence for extramedullary erythropoiesis in AMPKα1-/- spleen. Thick arrow indicates white pulp and thin arrow indicates red pulp E: Prussian blue-stained sections of spleen, showing a marked increase of blue stained iron deposits in spleen sections of AMPKα1-/- mice. F: Geimsa-stained blood smears from wild-type and AMPKα1-/- mice. Figure 2. Erythropoiesis is increased in AMPKα1-/- mice. A. Ter119-PE and CD71-FITC staining of splenocytes from wild-type and AMPKα1-/- mice. B. Bone marrow cells were stained with Ter119-PE and CD71-FITC. R1, proerythroblasts; R2, basophilic erythroblasts; R3, late basophilic and polychromatophilic erythroblasts; R4, orthochromatic erythroblasts. Figure 3. Reticulocyte counts and plasma EPO levels in AMPKα1-/- mice. A-B: Thiazol orange staining followed by flow cytometry and statistical analysis. Data were expressed as mean ± S.D (**p < 0.01 vs. WT, n = 6 in each group). C: Plasma EPO level determined by ELISA. Data were expressed as mean ± S.D (**p < 0.01 vs. WT, n = 10 in each group). . Figure 4 Effects of blood infusion on erythrocyte lifespan and clearance in AMPKα1-/- and wild-type mice. A: A representative of two independent experiments showing decreased lifespan of erythocytes in AMPKα1-/- mice (n = 3 in each group). The dotted line indicated the time for the 50% clearance of NHS-biotin labeled red blood cells in mice B: Clearance of reinfused biotin-labeled blood cells. WT-WT: blood from wild-type donor infused into wild-type recipients; KO-WT: blood from AMPKα1-/- donor infused into wild-type recipients; WT-KO: blood from wild-type donor infused into AMPKα1-/- recipients; KO-KO: blood from AMPKα1-/- donor infused into AMPKα1-/- recipients (n = 5–7 in each group). Figure 5. Osmotic fragility, and survival rate in wild-type and AMPKα1-/- mice following PHZ treatment. A: Decreased osmotic fragility in erythrocytes from AMPKα1-/- mice (n = 5 in each group). The dotted line indicated the concentration of NaCl solution in which 50% of red blood cells were lysed. B: Survival was markedly decreased in AMPKα1-/- mice compared with wild-type littermate controls after PHZ treatment (n = 6 in each group). Figure 6. Increased ROS levels and protein oxidation in erythrocytes of AMPKα1-/- mice. A. Erythrocyte intracellular ROS levels were determined by flow cytometery using a H2DCFDA probe without or with addition of exogenous H2O2 (50 µM). B. Increased protein oxidation in lysates of erythrocytes from AMPKα1-/- mice. A representative results from three independent experiments showing detection of oxidized proteins using DNPH-derived carbonyl immunochemistry staining. C. Quantification of the Quantitative analysis of the oxidative status of wild type and AMPKα1-/- erythrocytes by comparison of signal intensity of lanes in B using AlphaEaseFC software version 4.0.0 (Alpha Innotech Co.). Data were expressed as mean ± S.D (**p < 0.01 vs. WT, n = 3 in each group).

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

14

Figure 7. Chronic administration of tempol prolongs the lifespan of erythrocytes in AMPKα1-/- mice. A: The reticulocyte index was measured after 5 weeks in all groups. B: In vivo tempol therapy improves the lifespan of erythrocytes in AMPKα1-/- mice. Wild-type and AMPKα1-/- mice were provided normal drinking water or water containing tempol (1 mM) for 5 weeks, and the lifespan of in vivo biotinylated erythrocytes was measured. Tempol was administered continuously during the experimental period. C: Selected antioxidant genes were monitored in wild-type and AMPKα1-/- erythroblast cells by quantitative real-time RT-PCR. Gene expression was normalized to β-actin and is expressed as a percentage of the control values. Data were expressed as mean ± S.D (*p < 0.05, **p < 0.01, n = 4 in each group). D: Prolonged provide Tempol (1mM) in drinking water for 4 months, spleen index were measured (**p < 0.01, n = 6 in each group). E: Proposed mechanism illustrating that AMPKα1 is essential for normal erythrocyte lifespan. Table 1. Hematological parameters: analysis of mouse peripheral blood with a HEMAVET 950 Veterinary hematology analyzer.

Abbreviations: RBC, red blood cell count; Hb, hemoglobin concentration; HCT, hematocrit; MCV, mean corpuscle volume; MCH, mean corpuscle hemoglobin; MCHC, mean corpuscle hemoglobin concentration; RDW, red cell distribution width; WBC, white blood cell count; NE, neutrophil count; LY, lymphocyte count; MO, monocyte count; EO, eosinophil; BA, basophil count; PLT, platelet; MP, mean platelet volume. All data are expressed as means ± S.D (n = 5 in each group).

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

A.

B.

C.

AMPKα

GAPDH

Wild Type

AMPK α1-/-

Wild Type AMPKα1-/-

D.

E.

F.

Wild Type AMPKα1-/-

**

0.00.20.40.60.81.01.21.41.61.8

Sple

en in

dex

Wang, S., et al., Fig 1.

Wild Type AMPKα1-/- AMPKα2-/-

AMPKα1∆∆

50µm 50µm

100µm 100µm

100µm 100µm

by guest on March 27, 2020 http://www.jbc.org/ Downloaded from

Wild Type AMPKα1-/-B.

Bone Marrow

Ter-119-PE

CD

71-F

ITC

R1=0.93 ±0.34R2=33.16 ±4.92R3=3.23 ±0.62R4=13.24 ±3.88

R1=1.59±0.83R2=37.56 ±1.42R3=1.80 ±0.40R4=5.58 ±2.32

Wild Type AMPKα1-/-A.

Spleen

Ter-119-PE

CD

71-F

ITC

Ter-119-PE

CD

71-F

ITC

R1=0.20±0.08R2=16.96 ±1.35R3=1.63 ±0.04R4=25.37 ±2.19

R1=0.44 ±0.12R2=41.12 ±3.13R3=3.16 ±0.83R4=17.88 ±1.44

CD

71-F

ITC

Ter-119-PE

Wang, S., et al., Fig 2.

by guest on March 27, 2020 http://www.jbc.org/ Downloaded from

0.0

5.0

10.0

15.0

20.0

Ret

icul

ocyt

e(%

)

**

Wild Type AMPKα1-/-

A. B.

C.

**

500

Wild Type AMPKα1-/-0

100

200

300

400

600

EPO

leve

l (pg

/ml)

Wild Type AMPKα1-/-

Wang, S., et al., Fig 3.

by guest on March 27, 2020 http://www.jbc.org/ Downloaded from

A. B.

Days after Biotin

0

10

20

30

40

50

60

70

80

90

100

110

0 7 14 21 28 35 42

Perc

ent o

f lab

eled

cel

l (%

)

Wild TypeAMPKα1-/-

0

5

10

15

20

25

30

35

40

45

WT-WT KO-WT WT-KO KO-KOIn

fuse

d ce

ll cl

eara

nce

(%)

Wang, S., et al., Fig 4.

by guest on March 27, 2020 http://www.jbc.org/ Downloaded from

A. B.

Time after PHZ injection

0

1

2

3

4

5

6

7

day0 day1 day2 day3 day4 day5

Mic

e su

rviv

al n

umbe

r

Wild TypeAMPKα1-/-

0102030405060708090

100110

0.00

0.15

0.25

0.35

0.40

0.45

0.50

0.55

0.60

0.65

0.75

0.85

NaCl concentration (%)

RB

C H

emol

ysis

pers

cent

(%)

Wild TypeAMPKα1-/-

Wang, S., et al., Fig 5.

by guest on March 27, 2020 http://www.jbc.org/ Downloaded from

A. B.

C.

AMPKα1-/-Wild type

Loading

AMPKα1-/-Wild type

**

Cou

nts

H2DCFDA

Wild Type

AMPKα1-/-

H2DCFDA

AMPKα1-/-

Wild Type50 µM H2O2

Cou

nts

Wang, S., et al., Fig 6.

0

2000

4000

6000

8000

10000

12000

14000

16000

18000

Oxi

dize

d Pr

otei

n de

nsity

rel

ativ

e to

Loa

ding

by guest on March 27, 2020 http://www.jbc.org/ Downloaded from

Perc

ent o

f lab

eled

cel

ls(%

)

Days after Biotin

**

* *0

20

40

60

80

100

120

0 7 14 21 28 35 42

Wild TypeWild Type+TEMPOLAMPK α1-/-AMPK α1-/-+TEMPO

C.

B.

* *

**

**

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

Foxo3 Catalase SOD2 GPx-1

Rel

ativ

e m

RN

A L

evel

s

Wild Type AMPKa1-/-

A.

AMPKα1-/-Wild type

p<0.05

NS

0

5

10

15

20

25

Ret

icul

ocyt

ein

dex

(%)

No treatment controlTEMPOL treatment

Wang, S., et al., Fig 7.

D.

0

0.5

1

1.5

2

2.5

p>0.05

p<0.01

Sple

en In

dex

AMPKα1-/-Wild type

No treatment controlTEMPOL treatment

by guest on March 27, 2020 http://www.jbc.org/ Downloaded from

Wang, S., et al., Fig 7.

E.

by guest on March 27, 2020 http://www.jbc.org/ Downloaded from

Shaobin Wang, George L. Dale, Ping Song, Benoit Viollet and Ming-hui ZouAMPK alpha 1 deletion shortens erythrocyte lifespan in mice: role of oxidative stress

published online April 14, 2010J. Biol. Chem. 

  10.1074/jbc.M110.102467Access the most updated version of this article at doi:

 Alerts:

  When a correction for this article is posted• 

When this article is cited• 

to choose from all of JBC's e-mail alertsClick here

Supplemental material:

  http://www.jbc.org/content/suppl/2010/04/14/M110.102467.DC1

by guest on March 27, 2020

http://ww

w.jbc.org/

Dow

nloaded from

VOLUME 285 (2010) PAGES 19976 –19985DOI 10.1074/jbc.AAC119.009647

Correction: AMPK�1 deletion shortens erythrocyte life span in mice: Role of oxidative stress.Shaobin Wang, George L. Dale, Ping Song, Benoit Viollet, and Ming-Hui Zou

Supplemental Fig. 4 was mislabeled. This error has now been corrected and does not affect the results or conclusions of this work. The authorsapologize for the error.

FIGURE S4: CD71/Ter119/FSC histograms for bone marrow and spleen from wild type and AMPK�1�/� mice.

ADDITIONS AND CORRECTIONS

10738 J. Biol. Chem. (2019) 294(27) 10738 –10738

© 2019 by The American Society for Biochemistry and Molecular Biology, Inc. Published in the U.S.A.