Inhibition of monocyte chemoattractant protein-1 prevents diaphragmatic inflammation and maintains contractile function during endotoxemia

Experiments were performed in 8- to 10-week-old male C57BL/6 mice (Charles River Laboratories, Saint-Constant, QC, Canada). All procedures were approved by the institutional animal care and ethics committee, in accordance with the guidelines issued by the Canadian Council on Animal Care. The mice were anesthetized with a mixture of ketamine (130 μg/g) and xylazine (20 μg/g) prior to sacrifice.

To evaluate the direct effects of MCP-1 on cytokine expression by diaphragmatic muscle cells, primary diaphragmatic muscle cell cultures were established [9] by using single living muscle fibers to isolate myoblast precursors (satellite cells). In brief, excised diaphragm muscle strips were subjected to collagenase digestion and trituration to liberate individual fibers. The individual fibers were transferred into Matrigel-coated (Becton Dickinson, Franklin Lakes, NJ) plates. Diaphragmatic myoblasts were expanded in growth medium (20% fetal bovine serum, 10% horse serum, 1% chick embryo extract in DMEM) until attaining approximately 75% confluence. The cultures were then placed in differentiation medium (2% fetal bovine serum, 10% horse serum, 0.5% chick embryo extract in DMEM) to induce myoblast fusion into differentiated myotubes. All experiments were performed on day 5 of maintenance in differentiation medium. Diaphragmatic myotubes were washed with DMEM before stimulation with recombinant murine MCP-1 (100 ng/ml).

To determine the effects of MCP-1 on NF-κB activity in muscle cells, myoblasts were simultaneously transfected with a NF-κB-driven firefly luciferase reporter plasmid (pNF-κB; Clontech, Mountain View, CA) and a constitutively active thymidine kinase promoter-driven Renilla luciferase plasmid (pRL-TK; Promega, Madison, WI), as previously described [24]. In this system, the constitutively active Renilla luciferase serves as an internal control to adjust for any differences in transfection efficiency. For these studies, we used the C2C12 skeletal muscle cell line (ATCC, Manassas, VA) rather than primary skeletal muscle cells, as the latter are known to be resistant to standard transfection techniques [25]. C2C12 myoblasts (5 × 10⁵) were seeded onto 60-mm plates and transfected the following day at approximately 50% confluence, by using Lipofectamine 2000 (Invitrogen, Carlsbad, CA). On day 5 in differentiation medium, the cells were stimulated with murine MCP-1 (100 ng/ml) (R&D Systems, Minneapolis, MN), and the activity levels of both forms of luciferase (firefly and Renilla) were quantified by using the Dual-Luciferase Reporter Assay System (Promega). Light emission was measured in an L_max 384 luminometer (Molecular Devices, Downingtown, PA), and the results are expressed as the ratio of firefly (reflecting NF-κB activity) to Renilla luciferase activities in relative light units.

A commercial ELISA kit for murine MCP-1 (R&D Systems, Minneapolis, MN) was used to measure serum and tissue MCP-1 protein levels in duplicate, according to the manufacturer's instructions. Serum was collected by cardiac puncture, and total protein was extracted from the diaphragm, tibialis anterior, liver, and lung. Frozen tissue samples were homogenized in lysis buffer (1% Triton X-100, 50 mM HEPES (pH 8.0), 150 mM NaCl, 10% glycerol, 2 mM EDTA, 1.5 mM MgCl₂, 10 μg/ml aprotinin, 10 μg/ml leupeptin, 1 mM phenylmethylsulphonyl fluoride, 1 mM sodium orthovanadate). Homogenates were centrifuged 10 minutes at 10,000 rpm, and the supernatant protein content measured with Bradford assay (BioRad Laboratories, Hercules, CA).

To measure mRNA expression levels of MCP-1 and its receptor CCR2, IL-1α, IL-1β, and IL-6, total RNA from tissue or cell cultures was extracted by using Trizol reagent (Invitrogen) according to the manufacturer's protocol. ³²P-labeled, anti-sense RNA probes were synthesized from commercially available Multi-Probe-Template sets (BD Biosciences, San Diego, CA). Riboprobes were hybridized overnight at 56°C with 10 μg of sample RNA, according to the manufacturer's instructions. Protected RNA fragments were separated by using a 5% polyacrylamide gel and analyzed with autoradiography. For each RNA probe, all experimental groups were run on a single gel to allow quantitative comparisons. The bands representing mRNA content were quantified by using an image-analysis system (FluorChem 8000; Alpha Innotech, San Leandro, CA), and the signals normalized to the L32 housekeeping gene as a loading control.

To quantify macrophages and neutrophils, skeletal muscle cryosections (5 μm thick) were reacted with monoclonal antibodies directed against either macrophage F4/80 (1:75 dilution) (Abcam, Cambridge, MA) or neutrophil Ly-6G (1:50 dilution) (BD Biosciences). Nonspecific binding sites were blocked by incubating sections for 1 hour with PBS containing 3% BSA and 5% goat serum, followed by goat anti-mouse IgG Fab fragment (1:20 dilution) (Jackson Laboratories, West Grove, PA) for 30 minutes. Biotinylated rabbit anti-rat IgG secondary antibody (1:100 dilution) (Vector Laboratories, Burlingame, CA) was added and revealed by using the Vectastain streptavidin-HRP system (Vector Laboratories) with DAB substrate (Sigma-Aldrich Canada, Oakville, ON, Canada). To quantify inflammatory cell infiltration, the central and adjacent 20 × fields of the tissue were photographed by using a digital camera, and a stereology software package (Image-Pro Plus; Media Cybernetics, Silver Spring, MD) was used to overlay a 275-point grid onto each image (six photographs per muscle). Inflammatory cells were quantified by using a standard point-counting method, in which an abnormal point was defined as falling either on an inflammatory cell or on a myofiber invaded by such cells. The percentage area of inflammation was then calculated by dividing the number of abnormal points by the total number of points falling on the muscle tissue section [26]. The muscle images were selected in random order, with the operator blinded to the identity of the experimental groups.

As an additional index of neutrophil activity within tissues, myeloperoxidase (MPO) activity was determined [27]. In brief, frozen tissues were homogenized in 1 ml ice-cold 50 mM potassium phosphate buffer at pH 6.0. Homogenates were centrifuged at 12,000 g for 15 minutes at 4 degrees Celsius, and the supernatant was discarded. Pellets were resuspended, homogenized, centrifuged, and the pellets were resuspended in buffer. Assays were performed in duplicate on supernatant added to buffer containing 0.167 mg/ml o-dianisidine and 0.0005% H₂O₂. Enzymatic activity was determined spectrophotometrically by measuring the change in absorbance at 460 nm over a 3-minute period. Values are expressed as units per gram of tissue, with each unit representing the change in optical density per minute.

The diaphragm or EDL muscle was surgically excised for in vitro contractility measurements, as previously described [7, 28]. Muscles from the different experimental groups were selected in random order, with the individual performing the contractility measurements being blinded to their identity. After removal from the animal, muscles were transferred into Krebs solution (118 mM NaCl, 4.7 mM KCl, 2.5 mM CaCl₂, 1.2 mM MgSO₄, 1 mM KH₂PO₄, 25 mM NaHCO₃, and 11 mM glucose) chilled to 4°Celsius and perfused with 95% O₂/5% CO₂ (pH 7.4). The muscles were then mounted in a jacketed tissue-bath chamber filled with continuously perfused Krebs solution warmed to 25°Celsius. After a 15-minute thermoequilibration period, muscle length was gradually adjusted to optimal length (L_o, the length at which maximal twitch force is obtained). The force-frequency relation was determined by sequential supramaximal stimulation for 1 second at 5, 10, 20, 30, 50, 100, 120, and 150 Hz, with 2 minutes between each stimulation train. At the end of the experiment, L_o was directly measured with a microcaliper and the muscle blotted dry and weighed. Specific force (force/cross-sectional area) was calculated, assuming a muscle density of 1.056 g/cc and expressed in N/cm².

All data are presented as mean values ± SD (n = 6 per group). Group mean differences were determined with Student's t test, or with one-way or two-way ANOVA with post hoc application of the Tukey test to adjust for multiple comparisons. A statistics software package was used for all analyses (SigmaStat V2.0; Jandel Scientific, San Rafael, CA). Statistical significance was defined as P < 0.05.

To evaluate mRNA expression levels of MCP-1, diaphragms from saline and LPS groups of mice were analyzed with RNase protection assay, as shown in Figure 1a. MCP-1 mRNA was not detected in control diaphragms, but was greatly increased in the diaphragms of septic animals (Figure 1b). Conversely, expression levels of CCR2, the only known receptor for MCP-1, were downregulated in the diaphragm after LPS administration (Figure 1c). The upregulation of MCP-1 mRNA transcript levels was associated with a similar increase in MCP-1 protein content within the septic diaphragm, as shown in Figure 2a. MCP-1 protein levels were also found to be significantly elevated in the serum (Figure 2b), as well as in the lung, liver, and the tibialis anterior muscle (Figure 2c) of LPS-group animals. Interestingly, MCP-1 protein levels were two- to threefold higher in the diaphragm than in the hindlimb muscle (tibialis anterior) under septic conditions.

To determine whether the augmented levels of MCP-1 detected in the septic diaphragm were associated with increased leukocyte infiltration into the muscle, immunohistochemical analysis was performed with antibodies directed against markers for macrophages and neutrophils. As shown in Figure 3, no measurable differences between control and septic diaphragms were found in the numbers of either leukocyte population. This was further confirmed for the neutrophil population by the lack of change in diaphragmatic MPO activity, whereas MPO activity was greatly increased in the lungs of septic animals (Figure 3f).

The ability of MCP-1 to modulate proinflammatory cytokine gene expression in the diaphragm during sepsis in vivo was investigated by pretreating animals with anti-MCP-1 neutralizing antibody. As indicated in Figure 4, transcript levels for IL-1α, IL-1β, and IL-6, as well as for MCP-1 itself, were all significantly lower in the diaphragms of mice that were pretreated with the MCP-1 neutralizing antibody before LPS administration. Therefore, systemic blockade of endogenous MCP-1 in vivo had major effects on the regulation of these proinflammatory genes in the septic diaphragm.

We next sought to determine whether MCP-1 is capable of directly stimulating inflammatory responses in primary diaphragmatic muscle cell cultures examined at 4, 8, and 16 hours after stimulation. Interestingly, despite significant effects of MCP-1 neutralization on the expression of these genes in the septic diaphragm in vivo, the transcript levels of IL-1α, IL-1β, and IL-6 were unaltered by direct MCP-1 stimulation of skeletal muscle cells in vitro (no detectable expression under either unstimulated or stimulated conditions). As shown in Figures 5a and 5b, only MCP-1 itself was significantly upregulated by MCP-1 stimulation in diaphragmatic muscle cells, and this effect was noted at 8 hours after stimulation. Moreover, in keeping with the fact that MCP-1 did not upregulate these classic proinflammatory genes in primary muscle cell cultures, we also did not find any significant influence of MCP-1 treatment on the NF-κB transcriptional activity assay in C2C12 skeletal muscle cells (Figure 5c). Taken together, these results suggest that MCP-1 is capable of acting on skeletal muscle cells to upregulate its own expression, but in a manner not dependent on NF-κB pathway activation.

To evaluate the potential contribution of MCP-1 to the adverse effects of sepsis on the contractile function of skeletal muscles, two different approaches were used. First, to determine whether direct exposure of skeletal muscle fibers to MCP-1 has effects on contractile function, recombinant MCP-1 protein was injected into the EDL muscle. The dose of MCP-1 administered to the EDL was extrapolated from the diaphragmatic MCP-1 content (picograms per muscle weight) at 12 hours after LPS administration, as determined with ELISA and presented earlier in Figure 2. Figure 6a shows that at 12 hours after injection of recombinant MCP-1 into the EDL, a small but statistically significant reduction was noted in the force-generating capacity of the MCP-1-injected EDL muscles relative to the contralateral control (saline-injected) muscles from the same animals. Furthermore, as was the case for septic diaphragms at the same time point after LPS administration (12 hours), the MCP-1-injected EDL muscles did not show any histologic evidence of inflammatory cell infiltration (not detected in either saline- or MCP-1-injected muscles).

Second, to determine whether MCP-1 plays a role in diaphragmatic contractile dysfunction during sepsis, the force-generating capacity of the diaphragm was compared in animals pretreated with anti-MCP-1 neutralizing antibody versus an irrelevant isotype control immunoglobulin. As expected, LPS administration led to a major decrease in diaphragmatic force production 12 hours later. The LPS-induced depression of diaphragmatic force was unaffected by pretreatment with an irrelevant isotype control antibody. In marked contrast, the loss of diaphragmatic force production at 12 hours after LPS administration was greatly alleviated in animals pretreated with anti-MCP-1 neutralizing antibody, as illustrated in Figure 6b. These findings indicate that MCP-1 plays a significant role in the impairment of diaphragmatic function associated with acute endotoxemic sepsis.