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Sepsis-Induced Channelopathy in Skeletal Muscles is Associated with Expression of Non-Selective Channels

Balboa, Elisa∗,†; Saavedra-Leiva, Fujiko∗,†; Cea, Luis A.; Vargas, Aníbal A.; Ramírez, Valeria; Escamilla, Rosalba∗,§; Sáez, Juan C.∗,§; Regueira, Tomás

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doi: 10.1097/SHK.0000000000000916



Sepsis can lead to complex clinical manifestations due in part to uncontrolled host immune response to infections (1). It is a highly prevalent condition with an elevated mortality, ranging from 15 to 60%, depending on severity (1). Skeletal muscles are affected by the systemic inflammatory response during acute and late sepsis and represent one sepsis associate organ dysfunction (2). However, the cause and relative importance of muscle dysfunction during sepsis is not yet elucidated, mainly due to the lack of knowledge on possible molecular targets that could be used for designing rational therapeutics procedures. This condition is clinically known as sepsis associated intensive care unit acquired weakness (ICUAW) (3), which is associated with sarcolemmal injury in both, respiratory and peripheral muscles (3).

The characteristics of sepsis associated ICUAW are: reductions in force-generating capacity, which compromises diaphragm and peripheral muscle contractile performance in several animal models of sepsis (4). Muscle atrophy is observed in prolonged models of sepsis and is described as skeletal muscle wasting that results from increased proteolysis over protein synthesis (5, 6). The latter is likely to result from altered cytoplasmic Ca2+ concentration found consistently high in sepsis (7). Muscle altered mitochondrial bioenergetic function, namely mitochondrial dysfunction (MD), was obtained both in human and in animal models (8). Oxidative stress has also been implicated in the pathogenesis of sepsis-induced muscle MD and a mitochondrial-specific NO synthase (mtNOS) has been described as relevant (9).

Muscle becomes electrically unexcitable (10), suggesting that an acquired channelopathy may participate in the pathogenesis of ICUAW during sepsis. Recent studies in skeletal muscle atrophy have elucidated the critical role of de novo expression of poorly selective membrane channels including connexin hemichannels (Cx HCs) and P2X7 receptors (P2X7Rs) (11–13). Accordingly, myofibers deficient in connexin43 (Cx43) and Cx45 expression show drastic reduction in denervation-induced atrophy (11) or complete absence of muscle atrophy induced by glucocorticoids (12). Connexins (Cxs) are encoded by a gen family of 21 members in rodents. They form oligohexamers called hemichannels (HCs), which are precursors of gap junction channels (14). Cx HCs are permeable to ions (Na+, K+, and Ca2+) (15, 16). These HCs are expressed in different cell types found in diverse tissues including heart, brain, lung, and kidney (14), but they are not expressed by differentiated skeletal myofibers. However, Cx HCs are expressed in myofibers of denervated muscles (11). Although several previous reports have proposed a relevant role for Cx HCs in inflammation most of them are on organs different from skeletal muscles and they normally express Cxs (17). However and to our knowledge, the possible expression of Cx HCs in skeletal muscles during sepsis has not been reported.

We hypothesized that Cx HC expression is detrimental during sepsis and a relevant initial step in the development of sepsis-associated ICUAW. Using a characterized model of sepsis with evident skeletal muscle atrophy, in the present work, we show that skeletal muscles express de novo Cxs 39, 43, and 45 as well as P2X7Rs, which permeabilize the sarcolemma. Since these channels are poorly selective, we propose that they could partially if not completely explain the channelopathy known to affect skeletal muscle during sepsis. The possible consequences on other relevant sepsis-induced skeletal muscles changes including bioenergetics, mitochondrial membrane potential (MMP) and cellular and mitochondrial redox state are discussed. These findings could be relevant for future interventions and developing novel therapeutic strategies for the prevention of sepsis-induced skeletal muscle atrophy.



HEPES, leupeptin, pepstatin, phenylmethylsulphonyl fluoride, N-benzyl-ptoluene (BTS), sulfonamide, collagenase type I, suramin and ethidium (Etd+) bromide were from Sigma-Aldrich (St. Louis, Mo). DMEM/F12 culture medium, fetal bovine serum albumin, FURA-2AM, Mitosox, MitoTracker red and MitoTracker green were purchased Thermo Fisher Scientific (Waltham, Mass). Tramadol was obtained from Genfar (Gentilly, France). Previously characterized anti-Cx43, anti-Cx45, and anti-Panx1 antibodies were used (18). Anti-Cx39 and P2X7 receptor (P2X7R) antibodies were obtained from Santa Cruz Biotechnology Inc (Dallas, Tex) and Abcam (Cambridge, UK), respectively. Cy2- or Cy3-conjugated goat anti-rabbit IgG antibodies were purchased from Jackson Immuno Research (Indianapolis, Ind).


The present study was performed with approval by the bioethics committee of Pontificia Universidad Católica de Chile (N° 150323001). Male mice wild-type (WT) C57BL6 of 8 weeks of age were kept in standard housing conditions with a 12-h: 12-h dark:light cycle and with food and water ad libitum. Mice were under cecal ligature and puncture for 7 days. For this, mice were anesthetized using isoflurane (Baxter Healthcare, Guayama, Puerto Rico). The abdomen was shaved, and a midline incision (0.5 cm) was made below the diaphragm. The cecum was isolated, tied and punctured once with a 21-Gauge needle (0.8 × 38 mm), and a small amount of cecal material extruded. The cecum was returned to the abdomen and we added 200 μL of PBS 1X to ensure sepsis. The incision was closed with 20 mm Nylon surgical suture (Tagumedica S.A, Santiago, Chile). After the surgery, all mice were administered orally with tramadol (30 mg/kg) for 7 days every 24 h. All mice were verified their weight every day. Sham controls were subjected to the same surgical protocol and cecal isolation, but the cecum was neither tied nor punctured.

Mouse IL-6 immunoassay

Mouse IL-6 was measured in 50 μL of plasma using the Mouse IL-6 Quantikine ELISA Kit (R&D Systems Inc, Minneapolis, Minn) according to the manufacturer's instructions.

Histological analysis

Freshly dissected tibialis anterior (TA) were embedded in tissue mounting solution OCT (Andes import, Chile) and fast frozen in liquid-nitrogen-cooled isopentane (Merck, Germany). Serial cryostat sections of 16 μm thickness were obtained and placed on glass slides (75 × 25 mm, B&C, Germany) and fixed for 10 min with 4% paraformaldehyde (Electron Microscopy Sciences, USA) for immunofluorescence or cross-sectional area (CSA) analysis.

Strength of mice: inverted screen test

To notice differences in mice strength before and after the CLP procedure, we proceeded as described previously (19). The mice were placed in the center of a wire mesh screen. The screen was rotated to an inverted position, 20 cm to 30 cm above a quilted surface and starts the stop clock. The time was stop and wrote when the mouse fall off or when the time reached 60 s. The time was converted in score as follows: 0–10 s = 1; 11–25 s = 2; 26–60 s = 3 and 60+ s = 4.

Cross-sectional area (CSA)

The CSA of skeletal muscle fibers was measured as described previously (11). Briefly, cross-sections were fixed with 4% (wt/vol) paraformaldehyde and stained with H&E. The CSA was evaluated by using offline analyses by ImageJ software (National Institutes of Health, Bethesda, Md).


To detect different proteins in cross sections (16 μm) of TA muscles were blocked for 1 h in blocking solution (1% BSA, 50 mM NH4Cl, 0.01% Triton x-100, 1X PBS, pH 7.4) and then incubated at 4°C overnight with diluted primary anti-Atrogin1 (1:300), anti-Cx43 (1:300), or anti-Cx45 (1:300) antibodies. Samples were washed four times with PBS 1X and then incubated with an appropriate dilution of Cy2- or Cy3-conjugated goat anti-rabbit IgG antibodies (1:300) for 1 h on a dark room. Samples were washed four times with PBS 1X and one time with distilled water and mounted with Fluoromount-G, with DAPI (Electron Microscopy Science, Hatfield, Pa) on glass slides. The images were acquired in microscope Nikon Eclipse Ti, and the fluorescence intensity quantification was done using ImageJ.

Skeletal myofibers isolation

Myofibers of mouse flexor digitorum brevis (FDB) muscles were isolated from anesthetized and euthanized mice as described previously (18). Muscles were immersed in cultured medium (DMEM/F12, supplemented with 10% fetal bovine serum) containing 0.2% collagenase type I and incubated for 2.5 h at 37°C. Then, they were transferred to a 15 mL falcon tube containing 3 mL of Krebs buffer (in mM: 145 NaCl, 5 KCl, 3 CaCl2, 1 MgCl2, 5,6 glucose, 10 HEPES-Na, pH 7,4) plus 10 μM BTS (contraction inhibitor, to reduce muscle damage). Muscle tissue was incubated for 2 min with 200 μM suramin, a P2 receptor inhibitor, to prevent the effects of ATP released during the procedure. Then, tissue was gently triturated by passing it 7 times through a wide-tip Pasteur pipette. Myofibers were centrifuged at 1,000 rpm for 10 s (Kubota 8700 centrifuge, Tokyo, Japan) and the sediment was washed with HEPES buffered Krebs saline solution containing 10 μM BTS. Myofibers were gently triturated again by passing it 15 times through a narrow-tip so as to dissociate single myofibers. Dissociated myofibers were centrifuged at 1,000 rpm for 10 s and the sediment was washed and resuspended in HEPES buffered Krebs saline solution containing 10 μM BTS, placed in 1.5 mL Eppendorf tubes until use for further analyses.

Ca2+ signal measurement

The basal intracellular free Ca2+ signal was measured in isolated myofibers using the Ca2+ indicator FURA-2AM and following the manufacturer‘s instructions. Briefly, isolated myofibers were incubated with FURA-2AM (5 μM) in Krebs solution for 50 min at room temperature. Once finished the incubation period, the myofibers were washed with Krebs saline solution without dye, and placed on coverslips to measure the basal free Ca2+ signal (340 nm: 380 nm fluorescence ratio) under a fluorescence microscope Nikon eclipse Ti (Tokyo, Japan) as previously described (13).

Dye uptake assay

Cellular uptake of ethidium (Etd+) was evaluated by time-lapse measurements as described previously (18). In brief, freshly isolated myofibers plated onto plastic culture dishes were washed twice with Krebs saline solution (in mM: 145 NaCl, 5 KCl, 1 CaCl2, 1 MgCl2, 5.6 glucose, 10 HEPES-Na, pH: 7.4). For time-lapse measurements, myofibers were incubated in recording medium containing 5 μM Etd+ and recorded 5 min in control solution and then were recorded 5 min after an HC blocker was added to the control solution (11). The Etd+ fluorescence was recorded in regions of interest that corresponded to nuclei of myofibers using a Nikon Eclipse Ti inverted microscope and NIS-Elements software (Nikon, Tokio, Japan).

Laser confocal imaging

For mitochondrial function analyses the FDB myofibers were exposed to Mitosox (125 nM) a superoxide indicator or MitoTracker Red CMXRos (25 nM) to evaluate MMP in Krebs buffer for 30 min, washed with Krebs, and imaged live. In both cases to measure the total mitochondrial pool, cells were counterstained with MitoTracker Green (250 nM). Images were collected using a NIKON laser scanning confocal microscope. The fluorescence intensities were measured using ImageJ 1.42q software (NIH, Bethesda, Md).

Oxygen consumption

The oxygen consumption of muscle fibers was measured as described previously (20). Fiber bundles from soleus muscle (0.2–0.8 mg dry wet) were separated along their longitudinal axis with a pair of needle-tipped forceps under magnification using an Olympus SZ61 microscope. Bundles were then treated with 50 μg/mL saponin for 30 min in ice and subsequently washed in MIR05. High-resolution O2 consumption measurements were conducted at 30°C with the OROBOROS Oxygraph-2K (OROBOROS Instruments, Innsbruck, Austria).

Western blot analyses

Tibialis anterior (TA) muscle frozen on dry was disrupted in 500 μL of homogenization buffer (2 mM MgCl2, 0.25 M sucrose in 20 mM Tris buffer, pH 7.5) plus proteases inhibitors (5 μg/mL leupeptin, 5 μg/mL pepstatin, and 50 μg/mL phenylmethylsulphonyl fluoride). The homogenates were centrifuged at 3,000 g for 10 min and the supernatants aliquots were used to measure proteins. Proteins in the supernatants were quantified by bicinchoninic acid method (Thermo Scientific, Waltham, Mass).

Western Blot analyses were performed using 30 μg of protein/lane in 15% denatured polyacrylamide gel and electrophoresed on a nitrocellulose membrane for the detection by primary and secondary antibodies. Membranes were incubated with total OXPHOS Rodent WB antibody cocktail (1:1,000 dilutions) (ab110413, Abcam, Cambridge, UK). After the incubation with primary antibody, the membrane was washed and incubated with peroxidase-conjugated secondary antibodies (1:5,000 dilution; Amersham-Pharmacia, New York, NY), and bound antibody was visualized with ECL detection on Quansys Biosciences equipment. Loading control was made using the GAPDH Loading Control Monoclonal Antibody (MA5-15738, Thermo Scientific). The bands intensities were measured using ImageJ and normalized by GAPDH protein.


Data for each condition are summarized in column bar graph as means + standard error (SEM) or in a box-and-whisker graph; where the number of mice per treatment group is indicated in the legend of every figure. The data were evaluated by Student t test (two group comparison) and one/two-way ANOVA test (multiple comparisons) followed by Tukey (parametric distribution) and Kruskal–Wallis (non-parametric distribution) as post-hoc tests. Analyses were carried out using GRAPHPAD software. Significantly differences between group were considered when P < 0.05.


CLP-induced sepsis reduces body weight and muscle strength and induces skeletal muscle atrophy

Abundant evidence indicates that sepsis induces a myopathy characterized by reductions in muscle force-generating capacity, loss of muscle mass, and altered bioenergetics (21). However, the mechanisms leading to sepsis-induced skeletal muscle dysfunction are not well understood. In order to study these mechanisms we standardized the cecal ligation and puncture (CLP) model (22) in adult male mice. These mice showed a decrease in body weight reaching maximal values between days 3 and 6 after CLP and showed a tendency to recovery at day 7 (Fig. 1A). At day 7th blood levels of interleukin-6 (IL-6) increase by 400% in septic animals whereas they were almost undetectable in control mice (Fig. 1C). Moreover, the muscle strength evaluated at day 7 was reduced by 25% in mice treated with CLP compared with control mice (Fig. 1B).

Fig. 1:
Cecal ligation and puncture (CLP)-induced sepsis reduces weight and strength.

To study whether at day 7 post-CLP treatment muscle showed atrophy we measured the gastrocnemius (GS) muscle weight and the cross sectional area (CSA) of tibialis anterior (TA) muscle fibers. CLP-induced sepsis reduced the muscle weight by about 10% (Fig. 2A) and the CSA was also reduced by about 25% (Fig. 2B).

Fig. 2:
CLP-induced sepsis reduces ratio between gastrocnemius muscle (GS) weight and mouse weight, reduces the cross-sectional area (CSA) and increases Atrogin 1 and Murf immunoreactivity.

To evaluate the possible involvement of CLP-induced protein degradation via ubiquitin proteasome pathway in loss of muscle mass, we evaluated the presence of Atrogin-1 and Murf, two molecular markers of this catabolic pathway (5). Immunofluorescence analysis of TA muscle sections revealed strong Atrogin-1 and Murf immunoreactivity in myofibers of TA muscles from CLP treated mice whereas the reactivity of these proteins was absent or very low, respectively, in muscles of control mice (Fig. 2, C and D). Then, we decided to test our hypothesis at day 7 post-CLP.

Connexins (39, 43, and 45), and P2X7R are de novo expressed and are mainly located at the sarcolemma of skeletal myofibers in mice exposed to CLP

Previous studies have shown that denervated skeletal muscles show de novo expression of Cxs 39, 43, and 45 HCs in addition to P2X7R and upregulation of pannexin1 (Panx1), which play a relevant role in muscle atrophy (11). But there is no evidence of expression of these membrane channels forming proteins in skeletal muscles of mice exposed to CLP. To evaluate this possibility, we evaluated the reactivity of Cxs 39, 43, and 45 and Panx1 and P2X7R as well in slices from TA muscle. All three Cxs and P2X7R were detected in myofibers of CLP but not in control mice (Fig. 3, A and B). In addition, Panx1 was detected in both CLP and in control mice and the reactivity was not significantly changed in muscles of mice exposed to CLP (Fig. 3, A and B). All proteins analyzed were mainly detected in the contour of the myofibers likely to correspond to the sarcolemma.

Fig. 3:
CLP-induced sepsis induces de novo expression of Cxs 39, 43 and 45 and P2X7 receptor but does not affect Panx1 levels and increases the basal cytoplasmic Ca2+ signal and membrane permeability.

CLP-induced sepsis increases sarcolemma permeability and increases the cytoplasmic Ca2+ concentration

Considering that Cx HCs allow flow of ions like Na+ and Ca2+(13) and also has been seen that Cx43 and Cx45 are determinants in denervation induced atrophy (13), we decided to evaluate the intracellular Ca2+ signal using the radiometric fluorescent probe FURA-2 in freshly isolated myofibers from control and CLP mice. The basal Ca2+ signal, represented by the 340/380 ratio with FURA-2 of isolated myofibers from flexor digitorum brevis (FDB) muscles of CLP mice, increased 25% compared with that of myofibers of control mice (Fig. 3C).

Since Cx43 and Cx45 HCs are permeable to Ca2+(15) and we found both expressed in myofibers of CLP mice, we evaluated the presence of functional Cx HCs in freshly isolated myofibers. As previously reported (11, 13), we found very low Etd+ uptake in control myofibers (Fig. 3D). In contrast, myofibers of CLP mice presented about 4-fold higher dye uptake as compared with control myofibers and were blocked to control values by 200 μM lanthanum ions (La3+) (Fig. 3D), a Cx HC and P2X7R blocker. These results suggest that sepsis increases sarcolemma permeability through connexin HCs and P2X7Rs.

CLP-induced sepsis leads to mitochondrial dysfunction in skeletal muscle

Intracellular free Ca2+ concentration is important for the correct functioning of mitochondria such as energy production for cell activity and determining cell fate by triggering or preventing apoptosis (23). To determine whether the increased basal Ca2+ signal was related to the mitochondrial function in CLP mice, we measured the mitochondrial oxygen consumption using permeabilized muscle fibers from control and CLP mice. The mitochondrial oxygen consumption was found lower in muscles of CLP than in control mice (Fig. 4A). This finding is not explained by a reduction in the amount of complexes (Fig. 4, B and C), since there is no significant difference in the normalized abundance of the differences mitochondrial complexes between the control and the sepsis model.

Fig. 4:
CLP-induced sepsis leads to decreased oxygen consumption rate.

In addition, we also evaluated the mitochondrial membrane potential (MMP) using the fluorescent probe Mitotracker red (MTRed) that stains mitochondria in live cells and its accumulation is dependent upon membrane potential. The MMP was 3-fold lower in myofibers of CLP mice compared with that of myofibers from control mice, which was evident by the reduction in red fluorescence in myofibers of CLP mice compared with control myofibers (Fig. 5, A and B). The reduction in red fluorescence was not due to a reduced number in mitochondria because similar staining with Mitotracker green was detected in myofibers of CLP and control mice (Supplementary Fig. 1, and the intensity of MtRed fluorescence was normalized by MtGreen fluorescence.

Fig. 5:
CLP-induced sepsis leads to decreased mitochondrial membrane potential and increased superoxide production.

Since high cytoplasmic Ca2+ induces mitochondrial ROS production in cells (23), we evaluated mitochondrial superoxide production using Mitosox. As expected, the myofibers from CLP mice presented a 4-fold increase in red fluorescence than myofibers of control mice, indicating higher levels of superoxide (Fig. 5, C and D). Again, the number of mitochondria was similar in both conditions and the intensity of Mitosox fluorescence was normalized by MtGreen fluorescence (Fig. 5).


In the present work, we characterized a mouse model that presented the commonly course of sepsis observed in clinic. As expected, septic mice presented systemic inflammation, loss of strength, activation of metabolic pathways of muscle degradation and atrophy. In this model, myofibers were found to de novo express three Cxs and P2X7R, which are putative candidates to explain the channelopathy that characterizes the dysfunction of skeletal muscles in sepsis.

At day 7 post-CLP the high IL-6 concentration detected in the blood of mice reflected a generalized inflammatory condition. This cytokine has been associated previously with the induction of the expression of several metabolic pathways for muscle wasting (24). Accordingly, we observed reduction in strength, muscle mass and CSA, characteristics of muscle atrophy, a phenomenon described also in human sepsis in critical illness (4). Therefore, this animal model of sepsis used in the present work could be useful to identify new molecular targets to design strategies to prevent and treat early ICUAW.

In agreement the above proposal, we found that skeletal myofibers of septic mice express Cxs 39, 43, and 45 as well as P2X7R, which were absent in normal muscles as reported previously (11, 13, 18). These proteins were detected in the surrounding of the cells strongly suggesting that were located in the sarcolemma. Accordingly, the sarcolemma permeability to Etd+ of myofibers from septic mice was much higher than that of control mice. The Etd+ increase of myofibers from septic mice was completely blocked by La3+. Since Cx HCs, but not Panx1 channels or P2X7Rs, are blocked by La3+(25), it is likely that the referred increase in sarcolemma permeability was primarily mediated by Cx HCs.

Nonetheless, it remains possible that Cx HCs, P2X7R, and Panx1 channels in an orchestrate fashion permeabilize the membrane since Cx HC and Panx1 channels enable the releasing of ATP to the extracellular milieu where it can activate P2 receptors including P2X7Rs. Moreover, P2X7Rs are permeable to Ca2+ and the increase in intracellular free Ca2+ concentration activates both Cx HCs and Panx1 channels (26). Moreover, Cx43 and Cx45 (26), but not Cx39 (16), are also permeable to Ca2+ increasing the Ca2+ influx and cell overloaded. In agreement the basal intracellular Ca2+ signal of myofibers from septic mice was significantly higher compared with control myofibers. The increase in Ca2+ influx via P2X7Rs and Cx43 and Cx45 HCs could contribute to the activation of the ubiquitin proteasome pathway since levels of Murf and Atrogin 1 were drastically increased in myofibers of septic mice. These findings also strongly suggest that these newly express poorly selective membrane channels could be the cause of drastic reduction in membrane potential as reported long time ago but remained without explanation (27).

Cx HCs and P2X7Rs are novel pathways described in this study might also explain reduced muscle membrane excitability, because the sarcolemma depolarization caused by active Cx HCs and P2X7R could increase the number of inactivated Na+ channels, which together with the lower expression of Na+ channels induced by TNF-α (28) could drastically reduce the excitability of myofibers. In addition, de novo expression of functional Cx HCs and P2X7Rs should critically reduce the electrochemical gradient across the sarcolemma. This means that the intracellular Na+ and Ca2+ signal should increase in myofibers of septic mice and great amounts of ATP leading to a reduction in ADP for new ATP synthesis. All these changes should contribute to a mitochondrial malfunctioning and muscle weakness. Similar alterations in oxidative phosphorylation have been found in animal limb muscles and in muscle samples from septic human patients (8). Muscle of septic patients has been described as having signs of bioenergetic failure, comprising oxidative stress, MD, and ATP depletion (8). In our model, there was a clear decrease in the ability of the cells to use oxygen, namely, MD. Accordingly, we detected a reduction in oxygen consumption that was not explained by a reduced complexes number or concentration, but by a decrease in activity.

The loss of electrochemical gradient across the sarcolemma caused by the expression of functional poorly selective sarcolemma channels should reduce the MMP leading to MD, which is greatly affected by increase in intracellular Na+ or Ca2+ signal. In support to this possibility, it has been demonstrated that elevated intracellular Na+ or Ca2+concentration reduces the MMP (29). The intracellular Ca2+ overload could lead to great generation of reactive oxygen species and activation of the permeability transition pore contributing to reduce the MMP (29). Moreover, a reduced MMP has been found to precede an increase a mitochondrial oxidative stress in a model of sepsis (30) and increased generation of peroxynitrite and superoxide as detected in the present work. Interestingly, these oxidant agents reduce the intracellular redox potential, condition known to activate Cx HCs (31). Hence, they could contribute to maintain active the feed-forward inflammatory mechanism described above where Cx HCs, P2X7Rs, and Panx1 channels participate in an orchestrate way (Fig. 6). This mechanism is known to activate the inflammasome in denervated myofibers or myofibers of animals treated with glucocorticoids two conditions in which infiltrated inflammatory cells are absent but muscle cells generate proinflammatory cytokines (11, 12). Moreover, proinflammatory cytokines also induce the expression of Cx HCs in skeletal myofibers (32) and inflammatory conditions are likely to be additive. From this perspective, it is anticipated that treatment with glucocorticoids in septic patients could worsen the outcome of skeletal muscles and as a member of the MODS muscles could worsen the outcome of septic patients. In addition, it has been reported that using glucocorticoids as treatment does not decrease the mortality in mouse septic models (33) and the activation of glucocorticoids receptor has been associated with a bad outcome during sepsis (34). Nevertheless, IL-6-guided stratification identified a subgroup of septic mice with high risk of death that benefit from dexamethasone treatment (35). Thus, a putative therapeutic design that might improve the prognosis of septic patients could be the use of Cx HC and/or P2X7R blockers to reduce muscle inflammatory response together with glucocorticoid to block the inflammatory response mediated by cells of the innate immune system.

Fig. 6:
Scheme of events that might explain the possible involvement of Cx HCs, P2X7R, and Panx1 channels in sepsis-induced channelopathy.

In conclusion, our study showed that during a relevant model of sepsis, expression of functionally active Cx HCs takes place in the sarcolemma of muscle cells. The appearance of Cx HCs together with P2X7Rs was associated with an increase in intracellular Ca2+ signal, which may mediate MD and muscle atrophy.


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Cecal ligature; connexin; connexon; hemichannel; mitochondrial dysfunction; muscle waste; P2X7 receptor; pannexin

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