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Rinnakkaistallenteet Terveystieteiden tiedekunta
2019
Functional and biochemical responses of skeletal muscle following a
moderate degree of systemic iron loading in mice
Liang, C
American Physiological Society
Tieteelliset aikakauslehtiartikkelit
© The American Physiological Society All rights reserved
http://dx.doi.org/10.1152/japplphysiol.00237.2018
https://erepo.uef.fi/handle/123456789/7557
Downloaded from University of Eastern Finland's eRepository
Original Article
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Functional and biochemical responses of skeletal muscle following a moderate degree of 3
systemic iron loading in mice
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Chen Liang1, Marisa C. Mickey1, Candace N. Receno1, Mustafa Atalay2, and Keith C. DeRuisseau1
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1Department of Exercise Science, Syracuse University, 820 Comstock Ave, Room 201 WB,
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Syracuse, NY 13244, USA
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2Institute of Biomedicine, Physiology, University of Eastern Finland, P.O. Box 1627, 70211, Kuopio,
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Finland
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Author Contributions:
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C.L., M.A., and K.C.D. conceived and designed research; C.L., M.C.M, and C.N.R. performed
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experiments; C.L. and K.C.D. analyzed data; C.L., M.A., and K.C.D interpreted results of experiments;
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C.L. and K.C.D prepared figures; C.L. and K.C.D drafted the manuscript; C.L., C.N.R., M.C.M, M.A.,
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and K.C.D. edited and revised the manuscript; C.L., M.C.M, C.N.R., M.A., and K.C.D. approved the final
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version of the manuscript.
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Running Head: Moderate iron loading effects on skeletal muscle
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Key Words: oxidative stress; non-heme iron; glutathione; thioredoxin; proteolysis
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Number of Words: 5378
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Total Number of References: 52
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Corresponding Author:
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Keith C. DeRuisseau, Ph.D.
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Syracuse University
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Department of Exercise Science
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820 Comstock Ave, Room 201 WB
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Syracuse, NY 13244
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Tel: 315-443-9698
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Email: kcderuis@syr.edu
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ABSTRACT
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Excessive iron loading may cause skeletal muscle atrophy and weakness due to its free radical generating
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properties. To determine whether a clinically relevant degree of iron loading impairs skeletal muscle
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function, young male mice received injections (i.p.) of iron dextran (4 mg iron/200 µL) or 2 mM D-
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glucose (control) 5 days/week for two weeks (n = 10/group). Systemic iron loading induced an
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approximate 4-fold increase in the skeletal muscle non-heme iron (NHI) concentration. Soleus specific
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tension (1, 30-250 Hz) was lower among iron-loaded animals compared to controls despite similar body
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mass and muscle mass. Soleus lipid peroxidation (4-hydroxynonenal adducts) and protein oxidation
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(protein carbonyls) levels were similar between groups. In gastrocnemius muscle, reduced glutathione
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(GSH) and glutathione peroxidase (GPX) activity were similar but glutathione disulfide (GSSG) and
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GSSG/GSH ratio were greater in iron-loaded muscle. A greater protein expression level of endogenous
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thiol antioxidant thioredoxin (TRX) was observed among iron-loaded muscle while its endogenous
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inhibitor thioredoxin-interacting protein (TXNip) and TRX/TXNip ratio were similar. Glutaredoxin2
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(GRX2), a thiol-disulfide oxidoreductase activated by GSSG-induced destabilization of its iron-sulfur
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[2Fe-2S] cluster, was lower following iron loading. Additionally, protein levels of α-actinin and αII-
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spectrin at 240 kDa were lower in the iron-loaded group. Ryanodine receptor (RyR1) stabilizing subunit
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calstabin1 was also lower following iron loading. In summary, the contractile dysfunction that resulted
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from moderate iron loading may be mediated by a disturbance in the muscle redox balance and from
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changes arising from an increased proteolytic response and aberrant sarcoplasmic reticulum Ca2+ release.
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NEW & NOTEWORTHY
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While severe iron loading is known to cause muscle oxidative stress and dysfunction, the effects of a
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moderate degree of systemic iron loading on muscle contractile function and biochemical responses
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remain unclear. This study demonstrates that a pathophysiological elevation in the skeletal muscle iron
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load leads to force deficits that coincide with impaired redox status and structural integrity, and lower
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ryanodine receptor-associated calstabin1 in the absence of muscle mass changes or oxidative damage.
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INTRODUCTION
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Iron is a trace element well known for its involvement with various physiological processes that
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include oxygen transport, inflammatory response, energy metabolism, and DNA synthesis (48). While the
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total body iron is principally localized to the erythron, a sizeable amount of iron (10-15%) is contained
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within skeletal muscle that is needed for heme and non-heme iron (NHI) containing proteins including
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myoglobin, cytochromes and other enzymes required for energy metabolism (18). Although the study of
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skeletal muscle iron metabolism has largely centered on the effects of iron deficiency, recent work is
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drawing attention to metabolic and functional aberrations that may result from an increased muscle iron
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status. Notably, associations between an elevated skeletal muscle iron level with conditions including
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aging-associated muscle atrophy (2, 16, 21, 25, 51, 52), diabetes (22), hemochromatosis (23), and obesity
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(34) were reported. Moreover, the overall increase in muscle iron under these various conditions ranged
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between 2 to 7-fold above values observed in healthy controls. Thus, accumulated iron within this range
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may play an important role in skeletal muscle functional impairment and contribute to aging-associated
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muscle dysfunction and pathophysiology of certain diseases.
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While the precise mechanism(s) by which elevated iron imparts toxic effects within skeletal
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muscle is unknown, it is likely to be mediated via the increased production of reactive oxygen species
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(ROS) (14). Specifically, iron can exist as low molecular weight complexes (i.e., reactive iron or labile
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iron) that are capable of generating free radicals via Fenton chemistry (19). Thus, elevated iron levels
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could contribute to a pathophysiological state in muscle by triggering proteolysis (21, 35) and myonuclear
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apoptosis (45) by inducing ROS production. For example, iron loading is linked to increased skeletal
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muscle free radical generation (3), disturbed redox homeostasis (2), and altered cardiac muscle calcium
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regulation (28, 36), which are factors that can result in reduced muscle contractility. In addition, induction
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of muscle iron overload in mice via intraperitoneal (i.p.) injection of iron dextran led to elevated ROS
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production and reductions in muscle mass and strength (39). However, the total muscle iron level
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achieved in the previous study was much greater than what is typically observed under various
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pathological states. Thus, understanding the impact of a moderate degree of iron loading on skeletal
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muscle function and adaptation responses is clinically relevant.
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The purpose of this study was to determine how a moderate degree of muscle iron loading within
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a physiologically relevant range (i.e., increase of 2 to 7-fold) impacts skeletal muscle functional and
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biochemical responses. It was hypothesized that iron loading would result in muscle contractile
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dysfunction that would coincide with increased oxidative stress, antioxidant expression and activity, and
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impaired structural integrity. It was also postulated that iron loading would result in less calstabin1
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associated with the ryanodine receptor, which would signify dysregulated calcium release from the
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sarcoplasmic reticulum. Identifying potential consequences of an elevated muscle iron status could
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provide greater understanding of how clinical disease conditions and aging adversely affect skeletal
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muscle mass and function.
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METHODS
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Ethical Approval and Experimental Animals
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Male CD-1 mice were obtained from Charles River Laboratories. Mice were housed in groups of four and
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maintained on a 12 hr-12 hr, light-dark cycle under standard laboratory conditions. All mice were
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provided with autoclaved Purina LabDiet 5010 and water ad libitum. The animal use protocol was
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approved by the Syracuse University Institutional Animal Care and Use Committee and all experimental
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procedures followed guidelines established by the American Physiological Society for the use of animals
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in research.
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Experimental Design
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Twenty, 12-week-old male mice were randomly divided into iron treatment or control groups (n =
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10/group). Mice in both groups received 200 μL i.p. injections five days/week for 2 weeks. The iron
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treatment group received 4 mg of iron dextran/injection (Sigma, St Louis, MO) diluted in saline. The iron
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loading protocol was designed to increase skeletal muscle iron concentration to a physiologically relevant
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level shown to be associated with maladaptive states. Control animals received saline i.p. injections
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containing 2 mM D-glucose. On the day of tissue collection, animals were anaesthetized by an i.p.
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injection of Fatal Plus (Vortech Pharmaceuticals Ltd., Dearborn, MI) (80-120 mg/kg). Once in a surgical
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plane of anesthesia the right soleus muscle was quickly excised and utilized for muscle contractile
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experiments. Additional muscles including the left soleus, extensor digitorum longus (EDL), and
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gastrocnemius were harvested and trimmed of excess fat, weighed, frozen in liquid nitrogen and stored at
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−80°C for biochemical analysis. The liver was also harvested, frozen and stored at −80°C. The mice were
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euthanized by removal of the heart and diaphragm.
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Soleus Muscle Contractile Properties
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Following dissection from the animal, the right soleus muscle was immediately transferred to a Krebs-
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Henseleit buffer containing 12 μM d-tubocurarine (Sigma) and aerated with 95% O2 and 5% CO2 (pH =
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7.4). The soleus was secured by the tendons using two lightweight plexiglass clamps (Harvard Apparatus,
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Holliston, MA) and vertically placed between platinum wire stimulating electrodes in an organ bath
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maintained at 25°C. The distal end of the muscle was secured to a fixed Plexiglas rod. The proximal end
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of the muscle was connected to a force transducer (300C; Aurora Scientific, Aurora, Ontario, Canada) and
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the muscle was electrically stimulated to generate a series of twitch forces recorded via a data-acquisition
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system (Dynamic Muscle Control v4.1.6; Aurora Scientific). The muscle was stimulated with a stimulus
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duration of 0.5 ms to obtain optimal length (Lo) by adjusting the muscle length using a micrometer. Once
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Lo was determined the muscle length was measured by a caliper and all force measurements were
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obtained isometrically at Lo. To determine the force-frequency response, the muscle was stimulated at
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frequencies of 1, 15, 30, 50, 80, 120, 150, 250, and 300 Hz using a train duration of 500 ms with a 2-min
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interval between each stimulus. Peak tetanic tension (Po) was identified using a stimulus frequency of 120
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Hz. Following a 2-min recovery, the muscle underwent a fatigue protocol that consisted of repetitive
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stimulation (40 Hz, 0.5 train/s, and 300 ms train duration) for 5 min. Force was recorded continuously up
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to 15 min during the recovery period and data points at 30 s and 1, 2, 5, 10, 15 min were selected to plot
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recovery. At the end of the experiment, the muscles were removed, trimmed of excess fat, weighed, and
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frozen for non-heme iron (NHI) measurement. Muscle force was expressed in absolute, relative (% of
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peak tetanic tension), and/or values normalized to muscle mass or cross-sectional area, which was
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calculated by dividing muscle mass by the product of muscle density (1.06 g/cm3) and fiber length (13).
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The fiber length coefficient of 0.71 was used in the calculation (10).
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Muscle and Liver Non-heme Iron
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NHI was measured according to the method described by Rebouche et al. (40). Briefly, muscle and liver
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samples (~10 mg) were homogenized (1:40; wt:vol) on ice in ultrapure water (18.6 MΩ) using a Dounce
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homogenizer. An equal volume of protein precipitation solution containing 1 N HCL and 10%
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trichloroacetic acid was added to the tissue homogenates and mixed in 1.5 mL polypropylene tubes.
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Samples were vortexed and placed on a heat-block at 95°C for 1 hr. Following incubation, the samples
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were allowed to cool, vortexed, and centrifuged 8,200 x g for 10 min. The supernatant (100 μL) was
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aliquoted and added to an equal volume of chromagen solution containing 0.508 mM ferrozine, 1.5 M
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sodium acetate, and 0.1% thioglycolic acid, and blank solution containing 1.5 M sodium acetate, and 0.1%
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thioglycolic acid. The absorbance was measured at 562 nm (PowerWave HT, BioTek Instruments,
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Winooski, VT) following a 30-min incubation. NHI concentration was calculated from a standard curve
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(0, 2, 4, 6, 8, and 10 μg/mL) made by mixing protein precipitation solution with atomic absorption iron
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standard (Sigma).
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Western Blotting
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Soleus and gastrocnemius muscles were homogenized (1:20; wt:vol) in RIPA Lysis buffer (Santa Cruz
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Biotechnology, Dallas, TX) and centrifuged at 10,000 x g for 10 min at 4°C. The protein content of the
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soluble fraction was assessed by using the RC/DC protein assay (Bio-Rad, Hercules, CA). Proteins (~40
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μg) were individually separated by SDS-PAGE via 12% polyacrylamide gels containing 0.1% SDS.
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Following electrophoresis, the proteins were transferred to nitrocellulose membranes using a Mini-
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Protean Trans-Blot module (276 mA for 2 hr) or an iBlot system (Invitrogen, Thermo Fisher Scientific,
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Waltham, MA). The membranes were then blocked in PBS-Tween buffer containing 5.0% skim milk
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protein and 0.05% Tween-20 and incubated with a primary antibody directed against 4-hydroxynonenal
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adducts (4-HNE, Alpha Diagnostics, San Antonio, TX,RRID:AB_2629282), thioredoxin (TRX, Cayman
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Chemical, Ann Arbor, MI, RRID:AB_2725744), thioredoxin-interacting protein (TXNip, MBL
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international, Woburn, MA, RRID:AB_592934), catalase (CAT, CalBiochem, San Diego, CA,
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RRID:AB_10726597), copper-zinc superoxide dismutase (Cu-ZnSOD, Novus Biologicals, Littleton, CO,
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RRID:AB_925658), manganese superoxide dismutase (MnSOD, Cayman Chemical, RRID:AB_1213308),
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αII-spectrin (Santa Cruz, RRID:AB_671135), glutaredoxin2 (GRX2; MyBiosource, San Diego, CA, Cat#
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MBS2525062), α-actinin (GeneTex, Irvine, CA, RRID:AB_385929), ryanodine receptor (RyR1; Thermo
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Fisher Scientific, RRID:AB_2254138), calstabin1 (Thermo Fisher Scientific, RRID:AB_2102731), or
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GAPDH (Cell Signaling Technology, Danvers, MA, RRID:AB_561053) overnight at 4°C. This step was
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followed by incubation with a horseradish peroxidase–antibody conjugate (1:1,000-1:5,000) directed
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against the primary antibody for 1 hr at RT. The membranes were washed and subsequently treated with
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chemiluminescent reagents (Bio-Rad) and quantified using the ChemiDoc MP imaging system (Bio-Rad).
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Protein Carbonyls
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Protein carbonyls were measured as a biomarker of protein oxidation by using a commercially available
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OxyBlot assay kit (Millipore, Billerica, MA). Briefly, muscle samples (~10 mg) were derivatized to 2,4-
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dinitrophyenylhydrazone (DNP) and neutralized by 2 M Tris base in 30% glycerol. Following
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electrophoresis using 12% polyacrylamide gels, the proteins were transferred to nitrocellulose membranes.
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Protein bands were visualized and quantified using the same procedure as described above.
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Immunoprecipitation
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Gastrocnemius muscle homogenate was centrifuged (10,000 x g) at 4°C. Supernatants (10 µg/µL) were
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removed and incubated with protein A/G PLUS-Agarose beads (Santa Cruz) and mouse IgG (Santa Cruz)
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for 1 hour at 4°C to minimize nonspecific binding. After brief centrifugation the supernatants were
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removed and the beads were discarded. RyR1 was immunoprecipitated from the pre-cleared supernatants
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using an anti-RyR1 antibody (6 µg) incubated for 4 h followed by 24 h incubation with protein A/G
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PLUS-Agarose beads at 4°C. Following centrifugation the supernatant was removed and the beads were
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washed three times with PBS buffer. After adding Laemmli sample buffer to the beads, the sample was
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heated and loaded onto SDS/PAGE (4-20% gradient) precast gels (Bio-Rad). Proteins were transferred to
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nitrocellulose membranes using a Mini-Protean Trans-Blot module (276 mA for 2 hr) and a Western blot
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was then performed on the membranes using RyR1 and calstabin1 (Thermo Fisher Scientific) primary
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antibodies as described above. Calstabin1 expression was expressed relative to RyR1 protein expression
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of the same sample.
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Superoxide Dismutase Activity
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Total superoxide dismutase (SOD) activity was measured using a commercially available assay kit
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(Sigma). Muscle homogenate (1:20; wt:vol) was mixed with RIPA Lysis buffer (Santa Cruz) and
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centrifuged at 10,000 x g for 10 min at 4°C. Supernatant (20 μL) was added to 200 µL of WST Working
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Solution and 20 µL of Enzyme Working Solution, or 20 µL of Dilution buffer, and incubated at 37°C for
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20 min. Absorbance were determined at 450 nm by using a spectrometer (PowerWave HT, BioTek
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Instruments).
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Glutathione Peroxidase Activity
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Glutathione peroxidase (GPX) activity was measured from muscle homogenate (1:20; wt:vol) with tert-
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butyl hydroperoxide as substrate mixed with buffer containing 50 mM Tris-HCL and 0.1 mM EDTA in
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the presence of 1 mM NADPH, 2.5 mM GSH, and 1.5 U/ml GSSG reductase (47). Activity was
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determined by monitoring the reduction in absorbance per minute according to the oxidation of NADPH
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at 340 nm (PowerWave HT, BioTek). Activity was normalized to protein concentration of the sample and
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expressed as units per mg protein.
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Glutathione
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Total glutathione (TGSH) was measured using supernatant (10 μL) from muscle homogenate (1:20;
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wt:vol) centrifuged at 10,000 x g for 5 min at 4°C. Supernatant (20 μL) was diluted 10-fold and reacted
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with 0.25 mM 5,5′-dithiobis-(2- nitrobenzoic acid), 0.4 mM NADPH, and 0.6875 U/ml glutathione
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reductase (Sigma). GSSG was determined from muscle homogenate supernatant prepared in 5%
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metaphosphoric acid, and derivatized with 2% 2-vinylpyridine and triethanolamine. TGSH and GSSG
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were determined by monitoring the formation of 2-nitro-5-thiobenzoic acid at 412 nm (PowerWave HT,
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BioTek). The concentrations of TGSH and GSSG were obtained by linear regression from the standard
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curves and GSH was derived from subtracting GSSG from TGSH.
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Statistical Analyses
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All measures were analyzed using independent sample t-test to compare group differences. The
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assumptions of normality and homogeneity of variance were verified using Shapiro-Wilk test and
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Levene’s test, respectively. Box-Cox transformation was used if the assumption of normality was violated.
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A repeated-measures analysis of variance (ANOVA) was used to assess for the analysis of contractile
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function changes across different frequency levels. Mauchly’s criterion was used to determine whether
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data violated the assumption of homogeneity of variance, and Greenhouse-Geiser corrected significance
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values are reported alternatively when violation occurred. Test statistics of overall multivariate
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significance was achieved using Wilk’s Lambda. Area under the curve was calculated for force-frequency,
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fatigue, and recovery data using the trapezoidal integral method. Data are graphically depicted by boxplot
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in which whiskers represent the 5th and 95th percentiles and mean represented by a plus sign (+). A
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significance level of P < 0.05 was selected for all analyses, which were performed using SPSS v. 24.0
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(IBM Corp., Armonk, NY). Tabular data are reported as mean ± SD.
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RESULTS
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Non-heme Iron Level in Muscle and Liver
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The NHI level of muscles and liver is reported in Table 1. To ensure that the iron injection protocol
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resulted in a moderate increase in skeletal muscle iron status, we examined the NHI level of the soleus,
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EDL, and gastrocnemius muscles, in addition to the liver. NHI level of the muscles and liver from the
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iron-loaded mice was significantly greater than control (P < 0.01) values. Specifically, 10 days of iron
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injections resulted in a 3.7 to 4-fold increase in the muscle iron level. The liver NHI level was 5.2-fold
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greater compared to the value of the control group.
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Body Mass (BM) and Muscle Mass (MM) Characteristics
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Animal BM and MM values are displayed in Table 2. Body mass (P = 0.40) and soleus muscle mass (P =
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0.55) were not significantly different between iron and control groups. Soleus MM/BM ratio (P = 0.39)
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was also not significantly different between groups.
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Soleus Muscle Contractile Properties
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Soleus muscle twitch and tetanus contractile properties are displayed in Table 3. There was no significant
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difference in absolute (P = 0.34) or normalized (P = 0.13) twitch force of muscles between iron-loaded
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mice and controls. Also, no significant difference was observed for time to peak tension (P = 0.16), or
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half-relaxation (P = 0.21) time between groups. However, when normalized to muscle mass (P = 0.02)
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and calculated physiological cross-sectional area (P = 0.003), maximal isometric tetanic tension of
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muscles from iron-loaded animals was significantly lower than controls.
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To assess whether a moderate increase in muscle iron level compromised muscle function, we
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also determined the force-frequency response of the soleus muscle (Fig. 1). There was a significant
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interaction between group and stimulation frequency (P = 0.001). Muscle from iron-loaded mice
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displayed lower force values at stimulation frequencies tested at 1 Hz and between 30 to 250 Hz (P <
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0.05). Also, the area under the curve for normalized tension was significantly lower (P = 0.006) for iron-
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loaded mice compared with controls.
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The response of the soleus muscle to fatiguing contractions and recovery is shown in Fig. 2.
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There was an expected progressive reduction in force generation during the stimulation protocol. No
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significant difference between groups was observed for area under the curve during fatigue (P = 0.63).
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Following the fatigue-inducing protocol, muscle force gradually recovered over the 15-min recovery
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period. The area under the curve during recovery was not significantly different between groups (P =
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0.74).
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Oxidative Damage and Antioxidant Enzyme Measures
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Oxidative damage was determined by measurement of 4-HNE adducts and protein carbonyls. Analysis
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and representative images of oxidative damage levels from gastrocnemius muscle are shown in Fig. 3.
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Levels of 4-HNE adducts were not significantly different between iron-loaded and control groups (P =
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0.99). Protein carbonyls were also not different between groups (P = 0.57).
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Analysis and representative Western blot images of key antioxidant proteins from gastrocnemius
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are shown in Fig. 4 and Fig. 5. The protein expression level of TRX (P = 0.01) was significantly greater
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in the iron-loaded group compared with controls. Similar levels of TXNip (P = 0.50) and TRX/TXNip (P
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= 0.16) were observed between the two groups. However, GRX2 was lower (P = 0.04) in iron-loaded
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muscle compared with controls. CAT (P = 0.99), Cu-ZnSOD (P = 0.70) and MnSOD (P = 0.47) protein
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expression levels were similar between iron animals and controls. Total SOD activity was also similar
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between groups (P = 0.85).
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Analysis of antioxidant enzymes and activity related to glutathione metabolism are shown in Fig.
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6. While GSH (P = 0.39) and GPX activity (P = 0.82) were similar between two groups, muscle from
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iron-loaded mice displayed a significantly greater GSSG (P = 0.03) and GSSG/GSH ratio (P = 0.03).
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Muscle Tissue Structure Integrity
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The levels of αII-spectrin and α-actinin were assessed as an indication of myocyte structural integrity. As
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illustrated in Fig. 7, αII-spectrin at 240 kDa was significantly lower in the iron group compared with
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controls (P = 0.02). No differences were observed in its cleavage products at 150 kDa (P = 0.41) or 120
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kDa (P = 0.11). Similar to αII-spectrin, α-actinin levels were also lower in the iron-loaded gastrocnemius
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muscle (P = 0.04).
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Calcium Release Channel RyR1 and Calstabin1
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As illustrated in Fig. 8, immunoprecipitation of RyR1-associated calstabin1 was lower in the muscle of
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iron-loaded mice compared with controls (P = 0.04).
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DISCUSSION
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The iron loading regimen used in the present study caused a moderate elevation in the muscle NHI level
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that was associated with diminished muscle force production without changes in muscle mass. The results
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suggest that oxidative damage may not be a contributing factor to the muscle dysfunction since no
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changes were observed in lipid peroxidation or protein oxidation measures. However, changes in GRX2
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and GSH-TRX system in response to iron loading suggests conditions of an unfavorable redox status. The
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lower expression level of cytoskeletal proteins in iron-loaded muscle may also be reflective of increased
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proteolysis and reduced muscle structural integrity, which could be an additional factor responsible for
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reduced force production. A lower expression of RyR1-associated calstabin1 induced by iron loading also
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suggests aberrant sarcoplasmic reticulum (SR) Ca2+ release. The implications of these findings are
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discussed in the sections below.
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Muscle Iron Concentration following the 10-day Iron Loading Regimen
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The degree of iron loading achieved in the muscles was within the range of iron levels reported for
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various pathological conditions. Huang et al. (22, 23) reported a 2.1-fold greater skeletal muscle NHI and
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a 2.3-fold greater ferritin level of Hfe-/- mice, which is a model of hemochromatosis. Rats treated with
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STZ, a Type-I diabetes animal model, showed a 1.4-fold increase in muscle iron content (43). Using MRI,
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obese patients exhibited 1.2-fold greater paravertebral muscle iron level than non-obese individuals (34).
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Moreover, our group previously reported a 2 to 7-fold greater NHI concentration of the plantaris over the
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progression of aging in rats (16). Our current findings are consistent with those of others (51) whereby
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gastrocnemius muscle iron concentration was 3.4-fold higher in 32-month-old rats as compared with
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young animals. Thus, the moderate increase in muscle iron level achieved in this study is likely to be
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reflective of states that are clinically relevant.
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A high degree of muscle iron loading was previously shown to reduce endurance capacity and muscle
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strength in mice (39). However, the high degree of muscle iron loading that was achieved in the previous
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study appears to be beyond what is typically observed in pathological conditions. During our data
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collection, a study was published that examined cell signaling and muscle atrophy responses to moderate
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iron loading (24). Daily iron injections (10 mg iron dextran) for one week resulted in a 2.0-fold greater
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gastrocnemius iron level compared with controls. Notably, iron-loaded muscle displayed atrophy and
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reduced phosphorylation of Akt-forkhead box O3 signaling. Since these findings were divergent to our
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current data we conducted a pilot experiment to closely replicate part of the experimental design. Briefly,
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12-week-old male CD-1 mice were randomly divided into control (2 mM D-glucose in saline i.p.; n = 4)
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and 10 mg iron dextran (i.p.; n = 5) groups. All mice received a daily injection for seven consecutive days
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and gastrocnemius muscles were harvested 24-hrs following the last injection. In contrast to results of the
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previous report, we observed muscle NHI concentration of mice receiving 10 mg iron to be 5.1-fold
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greater than controls, which was greater than the iron level achieved by injecting 4 mg of iron for 10-days
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based on results of this current study. Thus, differences in the methodology used to assess tissue iron may
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explain why our findings differ from those of Ikeda et al. (24).
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Iron Loading Effects on Muscle Contractile Properties
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Muscles from iron-loaded mice exhibited reduced force production. A negative correlation between
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maximal specific tension and soleus NHI concentration (r = -0.807; P < 0.001) was also observed;
387
indicating a strong link between excessive muscle iron and impaired maximal force generation. The
388
reduction in normalized force values across a broad range of stimulation frequencies occurred in the
389
absence of differences in muscle mass. Our data are consistent with previous data obtained in vivo, in
390
which iron overloaded mice generated lower force on a maximal strength test. Furthermore, the
391
progressive loss of force over repeated trials was greater among the iron overloaded animals (39).
392 393
We postulate that the lower force production of iron-loaded mice may have partially been the result of a
394
deterioration in myofibril structural proteins. This is supported by our result of αII-spectrin and α-actinin
395
showing lower expression in the iron-loaded muscle. Notably, these cytoplasmic proteins can be cleaved
396
by calpains, which are Ca2+ dependent non-lysosomal cysteine proteases. αII-spectrin is located within
397
the contractile fibers close to the SR as well as on the plasma membrane in cardiomyocytes (4) and
398
functions in the maintenance of muscle cell structures and force transmission (5). α-actinin belongs to the
399
family of focal adhesion proteins, linking integrin receptors to the actin cytoskeleton. Since integrins are
400
important sites for force transmission through focal adhesions, the lower expression of α-actinin may be a
401
contributing factor to muscle dysfunction (27). Altered release of Ca2+ from the SR could also play a role
402
in mediating impaired muscle force production and structural integrity. Notably, RyR1 channel-
403
stabilizing subunit calstabin1 was lower in the iron-loaded muscles, which suggests elevated iron may
404
lead to increased calcium leak that could disrupt Ca2+ release during stimulated contractions. Down-
405
regulation of calstabin1 and abnormal Ca2+ leak from the SR was reported to contribute to diaphragm
406
muscle weakness following long-term mechanical ventilation (33). Muscle iron loading may have altered
407
β-adrenergic signaling that could explain the down-regulation of calstabin1 (31, 33). Further study would
408
be necessary to clarify the role of the sympathetic nervous system in Ca2+-relevant mechanism(s) of
409
skeletal muscle functional impairment resulting from an elevated iron load.
410
Iron-loaded muscle did not exhibit greater fatigability as hypothesized, nor did it show muscle weakness
411
during the post-fatigue period. This finding conflicts with studies showing increased fatigue and lower
412
endurance capacity among hemochromatosis patients (1, 15). The lack of difference in the fatigue
413
response could be a result of testing the muscle at a temperature lower than that of in vivo conditions.
414
Muscles display greater resistance to fatigue at a lower temperature (41) due to lower ROS production. In
415
light of the lower force production observed in the unfatigued state it may be that the experimental
416
condition led to an underestimation of the impact of iron loading on force generation during fatiguing
417
contractions.
418 419
Iron Loading Altered the Muscle Redox Status, but not Markers of Oxidative Damage or
420
Enzymatic Antioxidant Status
421
It was anticipated that iron loading would result in muscle oxidative damage as previous studies showed
422
an association between iron accumulation and increased ROS production. However, we did not observe
423
greater levels of 4-HNE adducts or protein carbonyls in soleus muscle of iron-loaded mice. This could be
424
explained by their increased removal since the accumulation of oxidatively damaged proteins induces
425
proteolytic activity which in turn facilitates damaged protein catabolism (37). Additional tissue iron could
426
also be stored in ferritin, which is protective against oxidative stress since ferritin protein is able to
427
sequester and stabilize iron. However, the dynamics of iron storage/release from ferritin could still lead to
428
increases in labile iron capable of participating in redox reactions (50). Oxidants including superoxide (8,
429
9, 20) and nitric oxide (42) can release iron from the ferritin molecule in vitro. We also anticipated a
430
greater antioxidant response to counter elevated ROS generation incurred by iron loading. However, iron-
431
loaded animals did not show a greater degree of total SOD activity or a difference in protein expression
432
levels of Cu-ZnSOD, MnSOD, or CAT. In contrast, a thiol antioxidant response was revealed as
433
evidenced by a greater TRX protein expression. Through its disulfide reductase activity the TRX system
434
plays an important role in cellular redox status regulation and antioxidant protection. The response of
435
greater TRX in these mice is consistent with a previous report showing greater TRX protein level and
436
activity in the brain of female rats that were loaded with iron (12). Indeed, TRX is a stress-inducible thiol-
437
containing endogenous antioxidant and redox-regulator, which was shown to be an indicator of oxidative
438
stress in a variety of disease states. For example, the presence of catalytically active iron and induction of
439
oxidative stress in lung-injury models was accompanied by TRX nuclear translocation and its interaction
440
with other redox-sensitive elements (37). It was also reported that TRX was associated with serum ferritin
441
and further increased by iron infusion in hepatitis C virus-infected patients (26). As an endogenous
442
inhibitor of TRX, TXNip expression was unaffected but inconsistent expression of these reciprocal
443
proteins was previously reported (46). Thus, despite greater TRX protein expression in response to an
444
elevated iron level, the unaltered ratio of TRX/TXNip may have limited the contribution of TRX in
445
regulating thiol homeostasis.
446 447
Oxidized glutathione (GSSG) and its ratio to the reduced form (GSH) were greater in iron-loaded muscle,
448
which further supports the occurrence of a shift in antioxidant pools. The greater level of TRX after iron
449
administration may have been a compensatory response to greater GSSG and GSSG/GSH ratio. Moreover,
450
there is cross-talk between the TRX system and GSH metabolism through disulfide yields (38), and
451
GRXs possibly play an important role in their interactions. GRXs are thiol-disulfide oxidoreductases that
452
belong to the TRX family of proteins that are critically involved in regulating redox and iron homeostasis
453
(6). GRX2 is localized to the mitochondria and can accept electrons from GSH and thioredoxin reductase
454
(17). Moreover, GRX2 is a member of the TRX protein family that possesses an iron-sulfur [2Fe-2S]
455
cluster, which may serve as a redox sensor (30). Elevated GSSG/GSH can disrupt the iron-sulfur cluster
456
of dimeric GRX2, resulting in the formation of enzymatically active, iron free monomer of GRX2 (30).
457
However, lower expression of GRX2 resulting from iron loading may have reduced its involvement in
458
redox and iron status control. For example, experimental inhibition of GRX2 in dopaminergic neurons
459
resulted in altered iron regulation, increased level of mitochondrial iron, and reduced iron-sulfur cluster
460
biogenesis and activities of proteins including mitochondrial complex I, mitochondrial aconitase, and
461
cytosolic aconitase (29). Thus, a decreased capacity for FeS cluster synthesis combined with greater iron
462
release from disrupted GRX2 [2Fe-2S] clusters may increase intracellular labile iron levels that could
463
alter iron regulation and ROS production (29). Furthermore, mice deficient in GRX2 showed altered
464
release of ROS from mitochondria isolated from liver and cardiac muscle (11). Overall, the iron-loading
465
impact on the muscle redox status may extend to the GRX system, in which lowered GRX2 expression
466
may link elevated muscle iron status with an additional mechanism of altered ROS generation and iron
467
status regulation.
468 469
Our findings support the notion that muscle iron loading may lead to disrupted skeletal muscle Ca2+
470
regulation. In addition to calstabin1 dissociation, Ca2+ channel remodeling including RyR1 thio-
471
nitrosylation, oxidation, and Ser-2844 phosphorylation was observed in controlled mechanical
472
ventilation-induced diaphragm muscle weakness (33). Thus, an iron-induced redox disturbance may lead
473
to deregulated Ca2+ homeostasis that activates calpain and triggers degradation of cytoskeletal proteins.
474
Moreover, evidence linking iron associated redox balance disturbances with damage to structural proteins
475
has also been documented. For example, an association between TRX and spectrin was reported in
476
HbEβ-thalassemic erythrocytes whereby upregulated TRX corresponded with disrupted cytoskeleton
477
integrity shown by presence of low molecular weight fragments of β-spectrin (7). A greater GSSG/GSH
478
ratio and modification of the spectrin cytoskeleton network was also displayed among patients with
479
Fanconi's anemia (32). In sickle cells and hippocampal neurons, a connection between a high GSSG/GSH
480
ratio and spectrin ubiquitination was reported. Although in this previous report a high GSSG/GSH ratio
481
led to a reduced level of spectrin ubiquitination and turnover, it provides additional support linking altered
482
redox status with the dysregulation of proteolytic capacity (44). Though there is limited evidence of a
483
direct link of α-actinin and the GSH-TRX system, a study showed that thioredoxin reductase facilitates
484
actin-denitrosylation and β2 integrin-specific adherence through focal adhesion kinase in neutrophils (49).
485
Additional work to directly connect iron loading effects on muscle contractile dysfunction by a
486
mechanism involving redox imbalance and membrane cytoskeleton deterioration is warranted.
487
488
Summary and Conclusion
489
This study examined skeletal muscle contractile properties in vitro using a mouse model subjected to a
490
moderate degree of systemic iron loading. Although iron loading did not trigger an atrophic response of
491
the muscles, our data indicate that iron may play an important role in the regulation of muscle functional
492
properties. The contractile dysfunction induced by iron may be mediated by perturbations in redox
493
signaling that is part of a regulatory mechanism that leads to an increased proteolytic response and
494
abnormal SR Ca2+ release. Furthermore, iron loading triggers a thiol antioxidant response that may further
495
alter iron status regulation. Future work could further explore thiol antioxidant interactions and
496
implications of elevated skeletal muscle iron status under various pathophysiological conditions.
497 498
Acknowledgements
499
Funding support for these experiments was provided by the Syracuse University SOE.
500 501
Conflict of Interest
502
The authors declare that they have no conflict of interest.
503
504 505
506 507
508 509
510 511
512 513
514 515
516 517
518 519
520 521
522 523
524
525 526
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670 671
672
673
674 675
676 677
678 679
680 681
682 683
684 685
686 687
688 689
690 691
692 693
694 695
696 697
698 699
700 701
702 703
704 705
706 707
708
Table 1. Tissue NHI level (nmol/gww) and mean ratio
709 710
Control Iron Mean Ratio
n 10 10
Soleus 700.8 ± 202.7 2619.7 ± 663.0** 3.7
EDL 383.6 ± 119.8 1547.9 ± 392.5** 4.0
Gastrocnemius 702.5 ± 351.0 2850.3 ± 937.7** 3.7
Liver 1509.1 ± 376.7 7826.3 ± 261.7** 5.2
Values are means ± SD. n = number of animals. Mean Ratio = Mean NHI of the Iron group relative to the Control group. **Different from Control: P < 0.01.
711 712
713 714 715 716 717 718 719 720 721
722
723
724
725
726
727
728
729
730
731
732
733
734
Table 2. Body mass and muscle mass
735
n BM, g Soleus mass, mg MM/BM Ratio
Control 10 40.3 ± 1.7 9.4 ± 1.0 0.233 ± 0.021
Iron 10 39.5 ± 2.3 9.9 ± 2.2 0.250 ± 0.055
Values are means ± SD. n = number of animals. BM, body mass; MM, muscle mass.
736
737 738
739 740
741 742
743 744
745 746
747 748
749 750
751 752
753 754
755 756
757 758
759
760
761
762
763
764
765
766
767
768
769
770
771
772
773
774
775
Table 3. Soleus contractile properties
776
Control Iron
n 7 6
Twitch
Pt, mN 42.5 ± 5.4 37.4 ± 11.0
Pt, N/cm2 4.6 ± 0.7 4.0 ± 0.7
TPT, ms 34.3 ± 5.3 38.3 ± 4.1
1/2RT, ms 50.7 ± 8.4 56.7 ± 7.5
Tetanus
Po, mN 240.0 ± 47.0 222.8 ± 62.7 Po/pCSA, N/cm2 26.2 ± 2.5 22.4 ± 0.3*
Po/MM, mN/mg 25.3 ± 1.8 22.4 ± 1.3*
Pt/Po 0.171 ± 0.021 0.178 ± 0.031 Values are means ± SD. n = number of animals. Pt, peak twitch tension; TPT, time to peak twitch tension; 1∕2RT, half-relaxation time;
Po, maximal isometric tetanic tension; pCSA, calculated physiological cross-sectional area; Pt/Po, twitch-to-tetanus ratio. *Different from Control: P < 0.05.
777 778 779
780
781
782
783
784
785
786
787
788
789
790
791
792
Figure captions
793
Fig. 1. Force frequency response of the soleus muscle obtained from control (n = 7), and iron-loaded (n =
794
8) mice. Values are means ± SD. *Different from Control, P < 0.05.
795 796
Fig. 2. Fatigue and recovery response of soleus muscle. A: Absolute active force during fatiguing
797
contractions and recovery. B: Percentage of initial active force during fatigue and recovery. n = 7 for both
798
control and iron-loaded groups.
799 800
Fig. 3. Oxidative damage in soleus of control and iron-loaded mice. A: Top: Arbitrary optical density of
801
4-HNE adducts expressed relative to GAPDH (n = 9-10 animals/group). Bottom: Representative Western
802
blot image of 4-HNE adducts and GAPDH for one control and iron-loaded muscle, respectively. B: Top:
803
Arbitrary optical density of protein carbonyls (PC) expressed relative to GAPDH (n = 9-10
804
animals/group). Bottom: Representative Western blot image of protein carbonyls and GAPDH for one
805
control and iron-loaded muscle, respectively.
806 807
Fig. 4. Protein levels of TRX system in gastrocnemius of control and iron-loaded mice. A: Top: Arbitrary
808
optical density of TRX expressed relative to GAPDH (n = 9-10 animals/group). Bottom: Representative
809
Western blot image of TRX and GAPDH for one control and iron-loaded muscle, respectively. B: Top:
810
Arbitrary optical density of TXNip expressed relative to GAPDH (n = 9-10 animals/group). Bottom:
811
Representative Western blot image of TXNip and GAPDH for one control and iron-loaded muscle,
812
respectively. C: Arbitrary optical density of TRX/TXNip (n = 9-10 animals/group). D: Top: Arbitrary
813
optical density of GRX2 expressed relative to GAPDH (n = 9-10 animals/group). Bottom: Representative
814
Western blot image of GRX2 and GAPDH for one control and iron-loaded muscle, respectively.
815
*Different from Control, P < 0.05.
816
817
Fig. 5. Antioxidant enzyme protein expression and activity in gastrocnemius of control and iron-loaded
818
mice. A: Top: Arbitrary optical density of CAT expressed relative to GAPDH (n = 9-10 animals/group).
819
Bottom: Representative Western blot image of CAT and GAPDH for one control and iron-loaded muscle,
820
respectively. B: Top: Arbitrary optical density of Cu-ZnSOD expressed relative to GAPDH (n = 9-10
821
animals/group). Bottom: Representative Western blot image of Cu-ZnSOD and GAPDH for one control
822
and iron-loaded muscle, respectively. C: Top: Arbitrary optical density of MnSOD expressed relative to
823
GAPDH (n = 9-10 animals/group). Bottom: Representative Western blot image of MnSOD and GAPDH
824
for one control and iron-loaded muscle, respectively. D: Total SOD enzyme activity (n = 9-10
825
animals/group).
826 827
Fig. 6. Glutathione and GPX activity in gastrocnemius of control and iron-loaded mice. A: GSH (n = 7-8
828
animals/group). B: GSSG (n = 7-8 animals/group). C: Ratio of GSSG/GSH (n = 7-8 animals/group). D:
829
GPX activity (n = 9-10 animals/group). *Different from Control, P < 0.05.
830 831
Fig. 7. Muscle cytoskeleton protein expression in gastrocnemius of control and iron-loaded mice. A: Top:
832
Arbitrary optical density of αII-spectrin expressed relative to GAPDH (n = 9-10 animals/group). Bottom:
833
Representative Western blot image of αII-spectrin and GAPDH for one control and iron-loaded muscle,
834
respectively. B: Top: Arbitrary optical density of α-actinin expressed relative to GAPDH (n = 9-10
835
animals/group). Bottom: Representative Western blot image of α-actinin and GAPDH for one control and
836
iron-loaded muscle, respectively. *Different from Control, P < 0.05.
837 838
Fig. 8. Immunoprecipitation of calstabin1 in gastrocnemius of control and iron-loaded mice. Top:
839
Arbitrary optical density of calstabin1 expressed relative to immunoprecipitated RyR1 (n = 9-10
840
animals/group). Bottom: Representative Western blot image of calstabin1 and RyR1 for one control and
841
iron-loaded muscle, respectively. *Different from Control, P < 0.05.
842
10 15 20 25 30
ific Tension (N/cm2)
*
* *
* *
*
1 2 3 4 5 10 15 20 0.06
0.08 0.10 0.12 0.14 0.16 0.18 0.20 0.22
Time (min)
Force (N)
Control Iron
Fatigue Recovery
40 50 60 70 80 90 100
Percent of initial force (%)
Control Iron
B A
Control Iron 0.0
0.2 0.4 0.6 0.8
4-HNE Adducts/ GAPDH
4-HNE Adducts
Control Iron
0.0 0.1 0.2 0.3 0.4 0.5
PC/ GAPDH
PC
