Altered sodium channel-protein associations in critical illness myopathy
© Kraner et al.; licensee BioMed Central Ltd. 2012
Received: 16 May 2012
Accepted: 30 July 2012
Published: 30 August 2012
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© Kraner et al.; licensee BioMed Central Ltd. 2012
Received: 16 May 2012
Accepted: 30 July 2012
Published: 30 August 2012
During the acute phase of critical illness myopathy (CIM) there is inexcitability of skeletal muscle. In a rat model of CIM, muscle inexcitability is due to inactivation of sodium channels. A major contributor to this sodium channel inactivation is a hyperpolarized shift in the voltage dependence of sodium channel inactivation. The goal of the current study was to find a biochemical correlate of the hyperpolarized shift in sodium channel inactivation.
The rat model of CIM was generated by cutting the sciatic nerve and subsequent injections of dexamethasone for 7 days. Skeletal muscle membranes were prepared from gastrocnemius muscles, and purification and biochemical analyses carried out. Immunoprecipitations were performed with a pan-sodium channel antibody, and the resulting complexes probed in Western blots with various antibodies.
We carried out analyses of sodium channel glycosylation, phosphorylation, and association with other proteins. Although there was some loss of channel glycosylation in the disease, as assessed by size analysis of glycosylated and de-glycosylated protein in control and CIM samples, previous work by other investigators suggest that such loss would most likely shift channel inactivation gating in a depolarizing direction; thus such loss was viewed as compensatory rather than causative of the disease. A phosphorylation site at serine 487 was identified on the NaV 1.4 sodium channel α subunit, but there was no clear evidence of altered phosphorylation in the disease. Co-immunoprecipitation experiments carried out with a pan-sodium channel antibody confirmed that the sodium channel was associated with proteins of the dystrophin associated protein complex (DAPC). This complex differed between control and CIM samples. Syntrophin, dystrophin, and plectin associated strongly with sodium channels in both control and disease conditions, while β-dystroglycan and neuronal nitric oxide synthase (nNOS) associated strongly with the sodium channel only in CIM. Recording of action potentials revealed that denervated muscle in mice lacking nNOS was more excitable than control denervated muscle.
Taken together, these data suggest that the conformation/protein association of the sodium channel complex differs in control and critical illness myopathy muscle membranes; and suggest that nitric oxide signaling plays a role in development of muscle inexcitability.
Critical illness myopathy (CIM) is the most common cause of severe weakness in patients in the intensive care unit [1, 2]. One of the hallmarks of the acute phase of CIM is paralysis due to loss of muscle’s electrical excitability [3, 4]. To determine the mechanism underlying loss of excitability, we utilize a rat model of CIM in which corticosteroid treatment is combined with denervation to mimic treatment of critically-ill patients with corticosteroids and neuromuscular blocking agents [5, 6]. In the rat model, inactivation of sodium channels is the cause of inexcitability [7, 8]. A major contributor to the increase in inactivation is a hyperpolarized shift in the voltage dependence of inactivation of the adult (NaV 1.4) and embryonic (NaV 1.5) skeletal muscle sodium channel isoforms expressed in CIM muscle . As the rat model of CIM occurs in genetically normal rats, this hyperpolarized shift in sodium channel gating must be due to post-translational modification of sodium channels or an alteration in sodium channel association with other proteins that modify gating.
The goal of this study was to identify biochemical changes that could potentially underlie the hyperpolarized shift in the voltage dependence of sodium channel inactivation in CIM. We found changes in several candidates, although at least one change appeared compensatory rather than causative. We identified an increase in the amount of neuronal nitric oxide synthase (nNOS) associated with the sodium channel in CIM as a change that might underlie the hyperpolarized shift in the voltage dependence of inactivation.
All animal protocols were approved by the Institutional Animal Care and Use Committee at Wright State University. Critical illness myopathy was induced by a combination of denervation and dexamethasone treatment. Briefly, rat muscle was denervated by removing a 10-mm segment of the sciatic nerve in isoflurane-anesthetized adult female Wistar rats (250 to 350 g body weight). Buprenorphine was administered subcutaneously for postoperative analgesia. Dexamethasone (4 mg kg -1) and antibiotics (0.2 mL of 2.27% Baytril, Bayer, Shawnee Mission, KS, USA) were administered intraperitoneally beginning on the day of denervation and continuing for 7 days. Rats were killed by carbon dioxide inhalation, and the tibialis anterior, soleus, and gastrocnemius muscles removed, weighed, and snap-frozen in liquid nitrogen. For control muscles, non-treated rats from the same group were used and muscles harvested in the same way. All muscles were stored at −80°C until use.
Membranes were prepared from the gastrocnemius muscles as previously reported . Samples were thawed on ice for 10 min and minced finely on a glass plate on ice. Samples were transferred to tubes containing sucrose buffer with protease and phosphatase inhibitors, using 10 volumes of buffer per gram of tissue. The sucrose buffer contained 0.3 M sucrose, 75 mM NaCl, 10 mM EGTA, 10 mM EDTA, 10 mM Tris, pH 7.4, 1 mM phenylmethylsulfonyl fluoride (PMSF), 1 mM iodoacetamide, 0.1 μg/mL pepstatin A, 10 μM leupeptin, 2.5 μM ALLN, and 1 mM sodium fluoride. Samples were homogenized using a PowerGen 700 tissue homogenizer at a setting of 6 for 30 s. The samples were centrifuged in a SS34 rotor at 4,500 g for 15 min. The supernatants were removed and centrifuged again. The final supernatants were centrifuged in a SW55 rotor at 138,000 g for 1 h. The pellets from this last step were resuspended in fresh sucrose buffers with inhibitors and used as the membrane fraction.
For classical purification of sodium channels, membranes were solubilized using NP40 and the sodium channel fraction sequentially purified on DEAE-Sepharose and wheat germ agglutinin-Sepharose columns as described . For all membrane preparation and channel purification procedures, samples, equipment, and buffers were kept at 0-4°C.
Membrane fractions were assayed for protein content using the Lowry protein assay, and equal amounts of protein were loaded for control and CIM samples for each protein assessed (approximately 40 μg protein per lane). For deglycosylation experiments and co-immunoprecipitation experiments, the same amount of membrane protein was used to initiate the experiment; and equal volumes of the final product were loaded for each control and CIM sample analyzed. SDS-PAGE was performed on Criterion 4% to 20% acrylamide gels for most samples; but 5% gels were used for the deglycosylation experiments, and 10% to 14.5% gels were used for analysis of syntrophin and β-dystroglycan co-immunoprecipitations to better resolve the molecular weight region being analyzed. Western blot transfers were carried out in a Criterion transfer apparatus using nitrocellulose as the blotting medium. Following transfer, blots were washed with PBS/0.1% Tween, blocked in 1% I-Block (Tropix) in PBS/0.1% Tween for 1 h; incubated with first antibody for 2 h; washed and incubated with mouse or rabbit secondary antibody conjugated to alkaline phosphatase (Tropix); and visualized using CDP-Star (Tropix) and a Fujifilm LAS-3000 close-caption device camera.
A pan-sodium channel antibody (pan NaV 1.x) was raised against the highly conserved peptide, TEEQKKYYNAMKKLGSKK, corresponding to SP19  at Open Biosystems, ThermoFisher Scientific. The affinity-purified antibody selectively reacted to sodium channel in skeletal muscle and other tissues and could be displaced by the peptide antigen in western blot analysis. The antibody to FGF13 was provided by Dr Geoffrey Pitts (Duke University). The NaV 1.4-specific monoclonal antibody was provided by Dr Robert L Barchi (Thomas Jefferson Medical School), and can be obtained commercially from Sigma (S9568). Other antibodies were obtained from the following commercial sources: FGF12 (Abgent AP6750b), plectin (Epitomics 1399–1), dystrophin (Vector Labs VP-D508), NaV 1.5 (Alomone ASC-005), nNOS (Invitrogen 37–2800), β-dystroglycan (Abcam ab 49515), and syntrophin (Abcam ab11425).
Some gels were stained rather than processed for western blots. For analysis of phosphoproteins, Peppermint Stick Phosphoprotein Molecular Weight Standards (Invitrogen), which include both phosphoprotein and non-phosphoprotein standards, were included on the gels as controls. After electrophoresis, gels were fixed in 50% methanol and 10% acetic acid for 30 min, transferred to the Pro-Q Diamond Phosphoprotein Stain (Invitrogen) for 90 min, then transferred to Destain (Invitrogen) for 30 min. The gel was de-stained twice more for 30 min; then washed twice in ultrapure water. The gel was imaged on a flatbed fluorescent scanner (Fujifilm FLA-5100). After the phosphoprotein stain, the gel was immediately processed for all-protein stain with SYPRO Ruby Protein Stain (Invitrogen) overnight. The gel was de-stained with 10% MeOH and 7% acetic acid for 45 min, washed twice in ultrapure water, and imaged on the fluorescent scanner. The containers with the gels were covered with foil to prevent photo-bleaching of the stains.
For proteins that were purified using gels, the gels were not fixed prior to the SYPRO stain. After the water wash, these gels were imaged on a UV light-box, their pictures taken with a digital camera, and the bands of interest excised and sent to the proteomics facility.
For deglycosylation with PNGase F (New England Biolabs), 100 μg of control or CIM membranes was adjusted to 80 uL volume; then 20 uL of 5X denaturation mix (5 mM PMSF, iodoacetamide, NaF, 12.5 μM ALLN, 0.5 μg/mL pepstatin A, 2.5% SDS, and 0.2 M DTT) was added and the sample incubated at 65°C for 20 min. Following denaturation and cooling to room temperature, 20 uL of 10% NP40, 20 μL of 500 mM sodium phosphate (pH 7.5), 59 μL water, and 1 μL (500 units) of PNGase F were added to each sample and the samples incubated at 37°C for 1 h. Samples were quenched by addition of gel sample buffer. For neuraminidase (New England Biolabs) treatments, 10 μg of control or CIM membranes were adjusted to 100 μL with 10 μL of 500 mM sodium citrate (pH 6.0), 10 μL of 10% NP40, 5 μL neuraminidase (250 units), and water. Samples were heated at 37°C for 4 h. As a control, 10 μg of fetuin was treated in the same way. Samples were quenched by addition of gel sample buffer. For all samples, mock treatments without enzyme were also carried out. Samples were analyzed by western blot with NaV 1.4-specific antibody.
The mass spectrometric analysis was performed on an optimized proteomics platform as previously reported . Briefly, protein gel bands were washed and subjected to standard in-gel trypsin digestion. Digested peptides were analyzed by capillary reverse-phase liquid chromatography coupled with tandem mass spectrometry (LC-MS/MS), in which the eluted peptides were analyzed by an MS survey scan followed by 10 data-dependent MS/MS scans on an LTQ-Orbitrap ion trap mass spectrometer (Thermo Scientific). Acquired MS/MS spectra were then searched against the rat reference database of the National Center for Biotechnology Information with differential modifications on serine/threonine/tyrosine (+79.9663 Da); then filtered by matching scores and mass accuracy to reduce protein false discovery rate to less than 1% using a concatenated reversed database . Furthermore, we manually validated MS/MS spectra of matched phosphopeptides using the following criteria: (1) the presence of signature phosphate neutral losses (−49 for doubly charged or −32.7 for triply charged) for serine/threonine phosphorylation; (2) the strong intensity (usually the top peak) of the neutral loss peak; (3) possible ambiguity of modification site assignment; and (4) comparison of the MS/MS spectra of modified peptides with the corresponding unmodified counterparts.
For each sample analyzed (control, CIM, control/peptide, CIM/peptide), 200 μL of protein G dynabeads (Invitrogen) was incubated with 40 μg of pan-NaV 1.x antibody - and for the peptide controls, 40 μg of blocking peptide - for 3 h to 4 h, rotating end-over-end. During this incubation, 3.0 mg of membrane protein was prepared for each sample. In a total of 1.8 mL volume, the sample was solubilized in a buffer containing 20 mM KPO4 (pH 6.5), 180 mM KCl, 1 mM EGTA, 0.5 mM MgCl2, 1 mM PMSF, 1 mM iodoacetamide, 1 mM NaF, 2.5 μM ALLN, 10 μM leupeptin, 0.1 μg/ml pepstatin A, and 1% NP40. Samples were vortexed, incubated on ice for 1 h, centrifuged at 13,700 x g in a refrigerated microfuge for 30 min, and the supernatants recovered for the immunoprecipitation. The antibody-bound beads were washed twice with PBS/0.5% NP40 and incubated with the solubilized channel protein, rotating end-over-end, overnight. The next morning, the beads were collected at the side of the tube using a magnet, and the supernatants removed. The beads were washed five times with the same buffer used for solubilization, except NP40/asolectin was used in the buffer instead of NP40. The samples were eluted from the beads with gel sample buffer. Equal volumes of sample were loaded for each sample (con IP, CIM IP, con IP/pep, CIM IP/pep), and starting membranes were used as a positive control (con memb, CIM memb). Six separate immunoprecipitations were carried out. Prior to eluting with sample buffer, all procedures were maintained at 0°C to 4°C.
Medial gastrocnemius muscles from control or CIM rats were removed, fixed in 4% paraformaldehyde for 1 h, cryoprotected in 15% sucrose solution overnight, and frozen in liquid nitrogen. Ten-μm-thick cross-sections were cut. The same rabbit FGF12 and mouse monoclonal nNOS antibodies used in western analysis were used for staining. For dystrophin staining of the FGF12 samples, the same mouse monoclonal used in western analysis was used, but for analysis of the mouse nNOS-stained samples, a rabbit dystrophin was used. Labeling of mouse monoclonal antibodies was visualized using a Dylight 488-conjugated donkey anti-mouse secondary antibody (Jackson ImmunoResearch Laboratories). Labeling of rabbit antibodies was visualized using a rhodamine-conjugated donkey anti-rabbit antibody (Jackson ImmunoResearch Laboratories). Images were obtained using a Fluoview FV 1000 confocal microscope and an X60 oil objective (Olympus Optical).
Wild type and nNOS knockout adult mice (B6.129 S4-Nos1 tm1Plh /J, Jackson Labs, 25 g to 30 g body weight) were denervated by removing a 0.5-mm segment of the left sciatic nerve in the upper thigh under isoflurane anesthesia (2% to 3% inhaled). Buprenorphine was administered subcutaneously for postoperative analgesia. Mice were sacrificed on days 3 or 7 by carbon dioxide inhalation. The extensor digitorum longus (EDL) muscle was dissected tendon to tendon; then muscle fibers were labeled with 10 μM 4-Di-2-ASP, and imaged using an upright epifluorescence microscope during recording of action potentials as previously described . For all experiments, the recording chamber was continuously perfused with solution containing (in millimoles per liter) NaCl, 118; KCl, 3.5; CaCl2, 1.5; MgSO4, 0.7; NaHCO3, 26.2; NaH2PO4, 1.7; and glucose, 5.5 (pH 7.3-7.4, 20-22°C) equilibrated with 95% O2 and 5% CO2.
Western blots were quantified using the software supplied with the Fujifilm LAS-3000 close-caption device camera. For western blots with multiple samples of control and CIM samples (for NaV 1.4, NaV 1.5, pan-NaV 1.x, FGF12, and FGF13), the average of the control was used as the 100% standard. All individual control and CIM samples were calculated relative to this number, and errors shown are SEM. Statistical comparison between control and CIM were carried out using Student’s t-test. For quantification of the co-immunoprecipitation, the CIM was calculated relative to the control (set at 100%) for each individual protein in each co-immunoprecipitation. The expression in CIM relative to control for all six co-immunoprecipitations was analyzed by Student’s t-test, and errors shown are SEM. Muscle excitability was compared by calculating the percent of inexcitable fibers in each muscle. At least six fibers were studied in each muscle. The percent for each muscle was compared between control and nNOS-null mice using Student’s t-test with n as the number of muscles studied.
While the amount of NaV 1.4 α subunit remained constant, its migration was altered. One potential explanation for altered migration of the NaV 1.4 α subunit is an alteration in the level of glycosylation. The NaV 1.4 α subunit is known to be highly glycosylated [16, 17]. Membranes from control or CIM tissue were treated with one of two enzymes, PNGase F, which removes N-linked glycosylation at the ASN-linkage  or a recombinant neuraminidase, which removes terminal sialic acid moieties . The samples were then probed in a western blot with the NaV 1.4-specific antibody. As shown, removal of the entire carbohydrate sugar tree at the N-linkage completely abolished migration differences between the control and CIM samples, while treatment with neuraminidase did not (Figure 1C). As a positive control for neuraminidase function, we treated fetuin (a protein with a large number of sialic acid moieties), under the same conditions and found a shift in migration (Figure 1D). Taken together, these data indicate that a focused removal of terminal sialic acid moieties is insufficient to account for the change in migration of NaV 1.4.
In the NaV 1.2 brain sodium channel, a complex interplay of phosphorylation of sites within the I-II loop and the III-IV loop carried out by protein kinase A and C reduce peak sodium currents . Skeletal muscle sodium channels are not as highly phosphorylated as their brain and neuronal counterparts , but phosphorylation could affect gating of sodium channels in CIM. NaV 1.4 is phosphorylated by protein kinase A in vitro at a single site . In the NaV 1.5 channel, phosphorylation of S1505 in the III-IV loop by protein kinase C both reduces peak current and shifts inactivation gating in the hyperpolarizing direction .
We compared biochemical properties of control and CIM sodium channels to find candidates that might account for the hyperpolarized shift in inactivation gating seen in the acute phase of CIM. We identified several biochemical changes in sodium channel in CIM, but the most promising candidates appeared to be alterations in sodium channel-associated proteins in CIM. In particular, nNOS is a promising candidate that was more associated with sodium channels from CIM muscle. In mice lacking nNOS, the normal reduction in excitability following denervation was greatly reduced. These data are consistent with the possibility that increased association of nNOS with sodium channels is involved in triggering loss of muscle excitability in CIM.
We identified an increase in NaV 1.5 in CIM muscle such that it is approximately 28% of the entire channel population. This is similar to our earlier estimate of 21% obtained by measuring current densities . In our previous study, both the TTX-insensitive (NaV 1.5) and TTX-sensitive (NaV 1.4) channels demonstrated similar hyperpolarizing shifts in inactivation gating in CIM , so increased expression of NaV 1.5 per se cannot be responsible for the shift.
A second change identified in membranes from CIM muscle was reduced glycosylation of the NaV 1.4 sodium channel. Removal of the entire carbohydrate ‘tree’ at the asparagine-linkage eliminated the molecular weight difference between the control and CIM channel. However, selective removal of sialic acid moieties with neuraminidase did not eliminate the size difference. Given that the NaV 1.4 channel is known to have multiple carbohydrate trees [16, 17], the simplest explanation for these observations is that some but not most of the carbohydrate trees are removed in CIM, removing some but not most of the sialic acids. Previous work shows that removal of sialic acid from sodium channels shifts inactivation gating in a depolarizing direction [16, 17, 31]. This is opposite of the hyperpolarizing shift we observed in CIM . Thus, removal of some carbohydrate trees may be a compensatory mechanism that moves the voltage dependence of inactivation towards more depolarized potentials.
One change we found in CIM muscle that could underlie the hyperpolarized shift in the voltage dependence of inactivation was an alteration in the composition of the dystrophin protein associated complex (DAPC) as summarized in Figure 5D. This figure is based not only on work in this paper, but also on work carried out by other investigators that identified the components of the DAPC (reviewed in ). In our hands, the DAPC appeared to dissociate more easily in control samples such that all components (except sodium channels) were present at higher levels in the CIM samples. This was especially true for β-dystroglycan and nNOS, which are present at much higher levels in CIM CoIPs. These observations suggested to us that the sodium channel-DAPC complex is bound more tightly in CIM, perhaps indicating that the sodium channel and cytoskeleton are in a different and more strongly ‘locked’ conformation in the disease. Alternatively, the constituent members of the DAPC may be dynamically regulated. In either case, the presence of the important signaling protein nNOS in the DAPC of CIM muscle suggests that NO signaling through this protein could contribute to the altered inactivation gating in CIM.
The protein components that we identified in the DAPC are consistent with those identified by other investigators [26, 27]. In skeletal muscle, the consensus C-termini (S/TXV-COOH) of NaV 1.4 and 1.5 sodium channels bind the PDZ domain of syntrophin at a site overlapping and/or closely adjacent to the binding site for nNOS . Through syntrophin, both sodium channels and nNOS bind the C-terminus of dystrophin . In cardiac muscle, the dynamic nature of this complex was shown by comparative analysis of control vs. syntrophin point mutation that causes Long QT syndrome. The syntrophin point mutation altered the complex constituents, such that the plasma membrane Ca2+ ATPase no longer bound syntrophin. This released inhibition of nNOS, allowed S-nitrosylation of the NaV 1.5 sodium channel, and altered gating .
Dystrophin is part of the muscle cytoskeletal system. In mdx mice, which lack dystrophin, sodium channel inactivation gating is shifted 10 mV more positively than that of control mice . This observation suggests that loss of cytoskeletal components shifts inactivation gating in a depolarizing direction, a finding consistent with our hypothesis that sodium channel inactivation gating is hyperpolarized in CIM because it is more tightly associated with cytoskeletal components. However, acute disruption of cytoskeleton by pressure during formation of seals during patch clamp measurements has been found to trigger hyperpolarized shifts in the voltage dependent of NaV 1.4 and NaV 1.5 activation and fast inactivation [34–37]. Thus, while it is clear that changes in cytoskeleton can have profound effects on the voltage dependence of sodium channel gating, we currently do not know which changes in cytoskeleton will translate into changes in sodium channel gating.
Our finding that nNOS is present at higher levels in the sodium channel-DAPC complex in CIM raises the possibility that increased signaling through NO-dependent pathways contributes to loss of muscle excitability in CIM (see however, ). There are several cell signaling pathways that are regulated by NO, including protein phosphorylation through cGMP-protein kinase  and direct nitrosylation of cysteine or other amino acid side chains, as discussed above for the cardiac NaV 1.5 . We measured phosphorylation changes in CIM and found no overall difference (Figure 2).
To determine whether increased association of nNOS with sodium channels could be involved in inducing inexcitability of muscle, we measured excitability following denervation in control and nNOS-null mice. In rats, denervation alone induces inexcitability in only a minority of fibers, so addition of corticosteroids is necessary to induce inexcitability [8, 30]. In control mice, denervation alone was sufficient to induce inexcitability so it was not necessary to co-administer corticosteroids. In nNOS-null mice, a greater percentage of muscle fibers remained excitable following denervation. There are multiple mechanisms that could account for maintenance of excitability following denervation in the absence of nNOS. Further study will be necessary to determine if the contribution of nNOS to inexcitability is mediated by its association with sodium channels as part of the DAPC.
We surveyed sodium channels and their associated proteins in control versus CIM muscle using a variety of biochemical techniques to identify candidates that could underlie the hyperpolarized shift in inactivation gating/loss of electrical excitability that is characteristic of CIM. While we identified a number of changes in CIM, including increased expression of the NaV 1.5 sodium channel and partial de-glycosylation of the NaV 1.4 sodium channel, it is the change in association of the sodium channels with members of the DAPC that seems most promising as a potential explanation for the shift in inactivation.
Critical illness myopathy
Dystrophin associated protein complex
Ethylene glycol-bis(2-aminoethylether)-N,N,N’,N’-tetraacetic acid
Fibroblast growth factor
Fibroblast growth factor homologous factors
Neuronal nitric oxide synthase
Peptide: N-Glycosidase F
Sodium dodecyl sulfate polyacrylamide gel electrophoresis.
This work was supported by NIH grant number NS040826 (MMR) and quantification of westerns and stained gels were carried out in the Wright State University Proteome Analysis Laboratory.
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