A major cutaneous burn injury produces multiple physiologic derangements; however, the mechanisms by which burn trauma produces distant organ injury and dysfunction remain unclear. As early as 1966, Baxter and colleagues described the presence of myocardial depressant factors in the serum of patients with burn trauma over more than 40% of the body surface area (1). Ferrara and colleagues described similar cardiodepressive properties of lymph collected from the area of the cutaneous burn injury (2), and we recently found that mesenteric lymphatic diversion ablated burn-related myocardial contractile depression (3). Although the precise nature of cardiodepressive factors present in the systemic circulation of burn subjects remains unclear, a number of potential candidates have been proposed. Mediators such as thrombin, histamine, complement products, leukotrienes, proinflammatory cytokines (including TNF-α, IL-1β, IL-2, IL-6, IL-8), nitric oxide, and PGE2 degradation products have been shown to mediate some aspect of cellular injury after burn trauma (4–10). Increased serum levels of inflammatory cytokines have been described in experimental and clinical burn injury, but others have reported a significant rise in circulating proinflammatory cytokines only when trauma is complicated by sepsis (11–13). Our previous studies have shown that experimental burn trauma over 40% of total body surface area (TBSA) increased circulating TNF-α, IL-1β, and IL-6 levels and promoted a rise in systemic antiinflammatory cytokine IL-10 levels; however, postburn circulating cytokine levels were significantly lower than those measured in tissue compartments and are significantly lower than systemic cytokine levels measured in sepsis (14–18).
The cardiodepressive effects of inflammatory cytokines are well recognized, and strategies that neutralize or inhibit TNF-α synthesis have been shown to provide considerable cardioprotection in experimental burn trauma (19, 20). However, the difficulty of defining the contributions of multiple mediators, catecholamines, activated complement, clotting factors, and cytokines present in the intact burn subject led us to develop an in vitro model to examine the effects of burn serum on myocardial inflammatory responses and function in an environment free of the neurohumoral and endocrine responses that occur in the intact burn subject. In this model, the addition of burn serum (10% by volume) to isolated perfused hearts or to primary cardiomyocyte cultures allowed us to examine myocardial contractile function as well myocardial inflammatory responses in vitro. This experimental approach allowed us to examine several components of burn cardiomyocyte secretion of inflammatory cytokines as well as myocardial mechanical function.
MATERIALS AND METHODS
Experimental animal model
Adult male Sprague-Dawley rats (320–350 g) were used in the present study. Animals obtained from Harlan Laboratories (Houston, TX) were conditioned in house for 5–6 days after arrival with commercial rat chow and tap water available at will. All studies performed were reviewed and approved by The University of Texas Southwestern Medical Center’s Institutional Review Board for the care and handling of laboratory animals and conformed to all guidelines for animal care as outlined by the American Physiological Society and the National Institutes of Health.
Rats were anesthetized lightly with methoxyflurane 18–20 h before the burn experiment, and body hair on the side, back, and neck was closely clipped. The neck region was treated with a surgical scrub, and the left carotid artery was exposed, and a polyethylene catheter (PE-50) inserted into the artery was advanced retrogradely to the level of the aortic arch. In addition, a polyethylene catheter (PE-50) placed in the right external jugular vein was used to administer fluids and drugs. All catheters were filled with heparinized saline, exteriorized, and secured at the nape of the neck. Eighteen hours after catheter placement, animals were deeply anesthetized (methoxyflurane), secured in a constructed template device, and the surface of the skin exposed through the aperture in the template was immersed in 100°C water for 10 s on the back and upper sides. Use of the template produced a well circumscribed burned area, avoided injury to the abdominal organs, and accomplished full-thickness dermal burns over 40% of the total body surface area. Exposure to this water temperature in adult rats has been shown previously by our laboratory to destroy all underlying nerves, to produce a transient rise in internal body temperature (<0.5° for 30–45 s) after exposure to the 100°C water, and to avoid injury to underlying organs. Sham burn rats were subjected to identical preparation, except that they were immersed in room temperature water to serve as controls. Immediately after immersion, rats were dried, returned to individual cages, and each external jugular catheter connected to a swivel device (BSP99 Syringe Pump, Braintree Scientific, Inc., Braintree, MA) for fluid administration (lactated Ringer’s solution, 4 mL/kg/% burn with one half of the calculated volume given during the first 8 h postburn and the remaining volume given over the next 16 h postburn). The total volume of Ringer’s given over the first 24 h postburn was 50–56 mL. Buprenorphine (0.5 mg/kg) was given every 8 h during the postburn period. Burned rats did not display discomfort or pain, moved freely about the cage, and consumed food and water within 20 minutes after recovering from anesthesia. In the sham burn group, the external jugular vein was cannulated, and lactated Ringer’s was given to maintain catheter patency (0.2mL/kg/h). Twenty-four hours after burn trauma (or sham burn for controls), blood was collected from the arterial catheter to prepare serum for use in cell/organ studies. All serum samples from sham and burned rats were screened for circulating levels of TNF-α, IL-1β, IL-6, IL-10, and nitric oxide (ELISA). Serum LPS levels were measured courtesy of Xoma Corporation (Berkley, CA).
To isolate cardiomyocytes, rats received an i.p. injection of heparin (2000 units) 20–30 min before sacrifice. The rats were decapitated, hearts were harvested and placed in a petri dish containing room temperature heart medium [113 mM NaCl, 4.7 mM KCl, 0.6 mM KH2PO4, 0.6 mM Na2HPO4, 1.2 mM MgSO4, 12 mM NaHCO3, 10 mM KHCO3, 20 mM d-glucose, 0.5× MEM (minimum essential medium), amino acids (50×, Gibco/BRL 11130-051), 10 mM Hepes, 30 mM taurine, 2.0 mM carnitine, and 2.0 mM creatine], which was bubbled constantly with 95% O2–5% CO2. Hearts were cannulated via the aorta and perfused with heart medium at a rate of 12 mL/min for a total of 5 min in a nonrecirculating mode. Enzymatic digestion was initiated by perfusing the heart with digestion solution, which contained 34.5 mL of heart medium described above plus 50 mg of collagenase II (Worthington 4177, Lot MOB3771), 50 mg BSA (bovine serum albumin), Fraction V (Gibco/BRL 11018-025), 0.5 mL trypsin (2.5%, 10×, Gibco/BRL 15090-046), 100 M CaCl2, and 40 mM BDM (2,3-butadedione monoxime). Enzymatic digestion was accomplished by recirculating this solution through the heart at a flow rate of 12 mL/min for 20 min. All solutions perfusing the heart were maintained at a constant temperature of 37°C. At the end of the enzymatic digestion, the ventricles were removed and mechanically disassociated in 6 mL of enzymatic digestion solution containing a 6-mL aliquot of 2× BDM/BSA solution (3 mg BSA, Fraction V to 150 mL of BDM stock, 40 mM). After mechanical disassociation with fine forceps, the tissue homogenate was filtered through a mesh filter into a conical tube. The cells adhering to the filter were collected by washing with an additional 10-mL aliquot of 1× BDM/BSA solution (prepared by combining 100 mL of BDM stock, 40 mM; 100 mL of heart medium described above; and 2 g of BSA, Fraction V). Cells were then allowed to pellet in the conical tube for 10 min. The supernatant was removed, and the pellet was resuspended in 10 mL of 1× BDM/BSA. The cells were washed and pelleted further in BDM/BSA buffer with increasing increments of calcium (100 μM, 200 μM, 500 μM, to a final concentration of 1000 μM). After the final pelleting step, the supernatant was removed, and the pellet was resuspended in MEM (prepared by adding 10.8 g 1× MEM, Sigma M-1018, 11.9 mM NaHCO3, 10 mM Hepes, and 10 mL penicillin/streptomycin, 100×, Gibco/BRL 1540-122 with 950 mL MilliQ water); total volume was adjusted to 1 L. At the time of MEM preparation, the medium was bubbled with 95% O2–5% CO2 for 15 min, and the pH was adjusted to 7.1 with 1 M NaOH. The solution was then filter sterilized and stored at 4°C until use. At the final concentration of calcium, the cell viability was measured, and cell suspensions with greater than 85% viability were used for subsequent studies. Myocytes with a rod-like shape, clear defined edges, and sharp striations were prepared with a final cell count of 5 × 104 cells/mL/well (16, 18, 20, 21).
Primary cardiomyocytes were pipetted into microtiter wells (5 × 104 cells/well), and burn serum (or sham serum) was added to each microtiter well (10% by volume). Separate aliquots of cells were pretreated for 30 min with either recombinant bacterial permeability increasing protein (rBPI) (15 μg/5 × 104 cells, Xoma Corporation, Berkley, CA) or anti-TNF-α (10 μM/5 × 104 cells, Endogen, Woburn, MA) before burn serum challenge. Cells were then incubated (CO2 incubator at 37°C for either 1, 2, 3, or 18 h); at the end of the incubation period, supernatants were collected to measure creatine kinase (CK) as an index of cell injury (CK Kit, procedure No47-UV, Sigma Chemical Co., St. Louis, MO), and inflammatory cytokines secreted into the supernatant were measured by ELISA. Cell viability and morphology were examined. Additional aliquots of myocytes were loaded with either Fura-2AM or SBFI to measure myocyte calcium and sodium, respectively.
Intracellular calcium ([Ca2+]i) and sodium ([Na+]i) measurements—
Myocyte loading with Fura-2 AM was accomplished over 45 min, whereas myocyte loading with sodium-binding benzofurzan isophthalate (SBFI) was accomplished over 1 h at room temperature in the dark. Myocytes were then suspended in 1.0 mM calcium-containing minimum essential medium (MEM) and washed to remove extracellular dye; myocytes were placed on a glass slide on the stage of a Nikon inverted microscope. The microscope was interfaced with Grooney™ optics for epiillumination, a triocular head, phase optics, and 30× phase contrast objective and mechanical stage. Excitation illumination source (300 W compact Xenon arc illuminator) was equipped with a power supply. In addition, this InCyt I.m. 2™ Fluorescence Imaging System (Intracellular Imaging, Cincinnati, OH) included an imaging workstation and Intel Pentium Pro 200 MHz–based PC. The computer controlled filter changer allowed alternation between the 340 and 380 excitation wavelengths. Images were captured by monochrome charge-coupled device (CCD) camera equipped with a TV relay lens. InCyt Im2™ Image software allowed measurement of intracellular calcium and sodium concentrations from the ratio of the two fluorescent signals generated from the two excitation wavelengths (340 nm/380 nm); background was removed by the InCyt IM2TM software. The calibration procedure included measuring fluorescence ratio with buffers containing different concentrations of either calcium or sodium. At each wavelength, the fluorescence emissions were collected for 1-min intervals, and the time between data collections was 1–2 min. Because quiescent or noncontracting myocytes were used in these studies, the calcium levels measured reflect diastolic levels.
Isolated coronary perfused hearts—
Adult control rats were heparinized, and hearts were removed and placed in ice-cold (4°C) Krebs-Henseleit bicarbonate-buffered solution (in mM: 118 NaCl, 4.7 KCl, 21 NaHCO3, 1.25 CaCl2, 1.2 MgSO4, 1.2 KH2PO4, 11 glucose). All solutions were prepared each day with demineralized, deionized water and bubbled with 95% O2–5% CO2 (pH 7.4; Po2 550 mmHg; Pco2 38 mmHg). A 17-gauge cannula, placed in the ascending aorta, was connected via glass tubing to a buffer-filled reservoir (Ismatec, model 7335-30, Cole-Parmer Instrument Co., Chicago, IL) and used to maintain perfusion of the coronary arteries by retrograde perfusion of the aortic stump. Coronary perfusion pressure was measured, and effluent was collected to confirm coronary flow rate. Contractile function was assessed by measuring intraventricular pressure with a H2O-filled latex balloon attached to a polyethylene tube and threaded through the apex of the left ventricular chamber. Peak left ventricular (LV) systolic pressure and LV end-diastolic pressure were measured with a Statham pressure transducer (Model P23ID, Gould Instruments Inc., Oxnard, CA) attached to the balloon cannula, and the rates of LVP rise (+dP/dt) and fall (−dP/dt) were obtained using an electronic differentiator (Model 7P20C, Grass Instruments, Inc., Quincy, MA) and recorded (Grass Model 7DWL8P). Left ventricular developed pressure was calculated by subtracting end-diastolic from peak systolic pressure (22). A Grass PolyVIEW Data Acquisition System was used to convert acquired data into digital form. Either sham burn serum or burn serum was added to an aliquot of Krebs-Henseleit buffer to achieve a final concentration of 10% by volume. Initially, each heart was perfused with serum-free Krebs-Henseleit buffer and stabilized for 10–12 min. Perfusion was then continued with buffer containing either sham burn or burn serum and continued for an additional 40–45 min. All indices of ventricular function were then measured. In an additional group of hearts, rBPI21 (kindly provided by Xoma Corporation, Berkley, CA) was added to the initial serum-free perfusate and circulated through the heart for 10–12 min. Perfusion was then continued with burn serum containing perfusate for 40–45 min; left ventricular function was then determined in rBPI-burned serum–treated hearts.
All values are expressed as mean ± SE of mean. Analysis of variance (ANOVA) was used to assess an overall difference among the groups for each of the variables. Levene’s test for equality of variance was used to suggest the multiple comparison procedure to be used. If equality of variance among the four groups was suggested, multiple comparison procedures were performed (Bonferroni); if inequality of variance was suggested by Levene’s test, Tamhane multiple comparisons (which do not assume equal variance in each group) were performed. Probability values less than 0.05 were considered statistically significant (analysis was performed using SPSS for Windows, Version 7.5.1).
To determine whether burn trauma per se produced a significant rise in systemic inflammatory cytokine levels that could, in turn, stimulate cardiomyocyte responses, TNF-α, IL-1β, IL-6, IL-10, and NO were measured in the sham burn and burn serum used for myocyte challenge; cytokine levels were negligible in myocyte supernatants prepared from sham-burned animals; burn trauma over 40% TBSA promoted a small but significant rise (P < 0.05) in serum TNF-α, IL-1β, IL-6, NO, and IL-10 levels 24 h after injury (Table 1). As shown in Table 1, there was no measurable LPS in any serum sample examined. The limit of detection for LPS was 1 pg/mL.
Primary myocyte cultures used in this study had a viability greater than 90%; myocytes maintained in medium under the normal experimental conditions (CO2 incubator at 37°C) retained the rod shaped morphology, and the cellular borders and cell striations remained well defined. There was no evidence of cellular blebbing, necrosis, or apoptosis in untreated primary myocyte cultures. The addition of burn serum (10% by volume) to myocyte supernatant produced a time-dependent decrease in cell viability while CK levels in the supernatant rose over the incubation period (Table 2). Longer periods of cardiomyocyte exposure to medium containing 10% burn serum (48 h) caused a progressive decrease in cell viability and cellular apoptosis (data not shown). Because of these preliminary studies, all subsequent studies were based on cardiomyocyte exposure to burn serum (10% by volume) for 3 h.
Myocyte inflammatory response to burn serum challenge
To determine the effects of burn serum on cardiomyocyte inflammatory response, burn serum (10% by volume) was added to cardiomyocyte suspensions (5 × 104 cells/mL) for 3 hours. This exposure to burn serum produced a significant rise in TNF-α levels (from 17 ± 2 to 1109 ± 91 pg/mL, P < 0.05), a rise in cardiomyocyte secretion of IL-1β (from 4 ± 1.8 to 32 ± 3 pg/mL, P < 0.05), an increase in IL-6 secretion (from 238 ± 13 to 1569 ± 139 pg/mL, P < 0.05), an increase in supernatant NO levels (from 3.1 ± 1.1 to 28 ± 2 pg/mL, P < 0.05), and a rise in myocyte IL-10 secretion (from 4.5 ± 0.7 to 28 ± 1 pg/mL, P < 0.05). The addition of serum prepared from sham-burned animals had a modest effect on either cardiomyocyte secretion of TNF-α (Fig. 1, upper panel), IL-1β (Fig. 1, lower panel), IL-6 (Fig. 2, upper panel), NO (Fig. 2, lower panel), and IL-10 (Fig. 3).
Effects of BPI on burn serum-related inflammatory responses
Despite no measurable LPS in the systemic circulation after burn trauma, we considered that minute levels of either gastrointestinally derived (23) or burn wound–derived (24) LPS could serve as an upstream mediator of cardiomyocyte inflammatory responses, triggering TNF-α secretion by this cell population; TNF-α could, in turn, trigger myocyte secretion of other mediators. To address this hypothesis, cardiomyocytes were pretreated with either recombinant bacterial permeability-increasing protein (rBPI, 15 μg/mL/5 × 104 myocytes) or a TNF-α antibody (rat anti-TNF, 10 μm per 5 × 104 cells, Endogen, Woburn, MA) for 30 min before cardiomyocytes are challenged with burn serum as described above. In our study, either rBPI or anti-TNF ablated the cardiomyocyte inflammatory response as indicated by the decreased secretion of TNF-α (Fig. 1, upper panel), IL-1β (Fig. 1, lower panel), IL-6 (Fig. 2, upper panel), NO (Fig. 2, lower panel), and IL-10 (Fig. 3).
Burn serum alters myocyte sodium/calcium homeostasis
To explore another aspect of cardiomyocyte cellular function, additional aliquots of cardiomyocytes were challenged for 3 h with burn serum (or serum prepared from sham-burned animals to provide appropriate controls); cells were then washed, loaded with either Fura-2AM or SBFI fluorescent indicator, and cardiomyocyte calcium and sodium levels were measured. As shown in Figure 4, burn serum promoted a significant rise in cardiomyocyte calcium levels (upper panel) compared with calcium levels measured in cardiomyocytes incubated in medium alone or in medium containing sham-burn serum. Similarly, exposure of cardiomyocytes to burn serum for 3 h produced a brisk rise in myocyte sodium levels (lower panel). Challenge of cardiomyocytes with serum prepared from sham-burned animals did not alter either intracellular calcium or sodium levels. Pretreating cardiomyocytes with either rBPI or antiTNF attenuated the burn-serum-mediated rise in either intracellular calcium or sodium.
Burn serum depresses function of isolated perfused hearts
To determine the myocardial functional consequences of burn serum or sham-burn serum challenge, several groups were included. First, hearts isolated from naive control rats were perfused with serum-free Krebs-Heinseleit bicarbonate buffer for 55–60 min at a constant preload (0.07 mL LV volume) and constant coronary flow rate (8 mL/min); all measures of left ventricular function (LVP) remained stable during this perfusion, and data collected after 60 min perfusion are summarized in Table 3. Hearts perfused for an identical time period with perfusate containing 10% sham-burn serum had higher LVP values than LVP measured in hearts perfused with Krebs buffer alone (P < 0.05) despite identical preload (0.07 mL LV volume) and heart rate in all hearts during this stabilization period. Addition of burn serum (10% by volume) to cardiac perfusate significantly impaired left ventricular performance, producing measures of LVP and ±dP/dtmax that were significantly lower than values measured in the absence of burn serum. In addition, coronary perfusion pressure measured in burn-serum-challenged hearts was significantly higher than that measured in hearts perfused with buffer in the absence of burn serum (Table 3).
To further assess the effects of burn serum (or sham-burn serum) on myocardial performance, left ventricular function curves were constructed by measuring left ventricular developed pressure and ±dP/dtmax responses to increases in either left ventricular volume (preload) (Fig. 5) or incremental increases in coronary flow rate (Fig. 6). As shown in Figure 5, all measures of left ventricular contraction and relaxation were significantly lower in hearts perfused for 45–60 min with burn serum containing buffer despite identical levels of coronary flow rate and identical heart rates in all perfused hearts. Similarly, examination of left ventricular performance–coronary flow relationships (Fig. 6) showed that hearts perfused with burn-serum-containing buffer generated significantly lower left ventricular pressure and ±dP/dtmax responses at each level of coronary flow compared with hearts perfused with buffer alone or hearts perfused with buffer containing sham-burn serum.
Additional experiments included perfusing control hearts with Krebs-Henseleit buffer for 10–12 min. rBPI (15 μg/mL perfusate) was then added to the perfusate, and perfusion was continued for 10 min before the addition of burn serum (10% by perfusate volume). Perfusion with burn-serum-containing buffer was continued for an additional 40–45 min, and function was measured. As shown in Figures 5 and 6, pretreating hearts with rBPI prevented burn-serum-related myocardial contraction and relaxation defects.
In our study, LPS was not detected in serum collected 24 h after cutaneous burn injury over 40% total body surface area despite previous reports by our laboratory and others that major burn injury alters gastrointestinal barrier function and promotes bacterial translocation to mesenteric lymph nodes, liver, and spleen (23, 25, 26). Exposure of cardiomyocytes to medium containing 10% burn serum by volume evoked significant inflammatory responses as evidenced by the increased myocyte secretion of TNF-α, IL-1β, IL-6, NO, and IL-10. This cardiomyocyte cytokine response was similar to the cardiomyocyte pro- and antiinflammatory cytokine responses that we have shown to occur in vivo after either burn trauma or sepsis (16, 18, 20). Pretreating cardiomyocytes with rBPI (a therapeutic strategy clearly recognized to neutralize LPS) prevented the myocyte inflammatory responses to burn serum challenge.
Burn trauma in the intact subject initiates an inflammatory cascade that includes the synthesis of numerous mediators by diverse cell populations. In our study, the direct application of burn serum to primary cardiomyocyte cultures allowed us to eliminate the contribution of mediators secreted by several cell populations within the myocardium (for example, endothelial cells, emigrated leukocytes or macrophages, as well as fibroblasts). Furthermore, our finding that burn-serum challenge in control naive hearts produced myocyte inflammatory responses and myocardial depression confirmed that circulating mediators, in the absence of neurohumoral and endocrine function, produced downstream organ dysfunction.
The idea of circulating myocardial depressant factors after major burn trauma is not a new concept; in 1966, Baxter and colleagues reported that serum collected from patients with burn trauma over greater than 40% TBSA produced significant contractile deficits when added to the perfusate of control guinea pig hearts (1). Concerns with this early study were twofold; first there were species-related differences involved in the application of human serum to rodent hearts; second, early studies failed to provide information regarding the mechanisms by which burn serum altered cardiac function. More recently, Ferrara and colleagues extended Baxter’s previous work and reported that collection of lymph from a burned limb, and addition of this burned lymph to isolated perfused rat hearts, produced profound contractile defects (2). Mesenteric lymph diversion has been shown to ablate burn-related cardiac contractile dysfunction (3). More recently we reported that interrupting Toll/IL-1 signaling, a pathway clearly mediated by LPS, abrogated burn-related myocardial contractile defects; and in vivo administration of recombinant BPI (rBPI) after burn injury provided considerable cardioprotection (27, 28).
In this present study, blood collected from Sprague-Dawley rats 24 h after scald burn over 40% TBSA was used in ventricular muscle preparations or in primary cardiomyocyte cultures prepared from rats of the same strain, eliminating concerns about species-related differences. In addition, our data suggest that one mechanism by which burn serum promoted myocardial contractile depression was enhanced myocyte secretion of inflammatory cytokines TNF-α, IL-1β, and IL-6.
Our interest in LPS in burn serum as a potential mediator of downstream organ dysfunction arose from our previous finding that burn trauma alters splanchnic perfusion, producing transient intestinal ischemia (23). This loss of gut barrier integrity and translocation of GI-derived bacteria/endotoxin would be expected to initiate an inflammatory cascade that contributed to downstream organ injury and dysfunction. Although LPS was not detectable in any serum sample collected from burned rats, these data do not preclude the presence of minute LPS levels in the systemic circulation that are below the detection limits of our assay system. LPS may be present in the serum but bound to lipoproteins, making LPS unmeasurable by current assay methodology. Alternatively, LPS may be cleared rapidly from the portal venous blood by the liver, secondarily activating second messengers such as TNF-α, which then mediate myocardial depression. It is also possible that LPS in the portal circulation stimulates hepatic synthesis of some as yet unrecognized mediator that, in turn, serves as a proximal signal to initiate downstream organ inflammatory responses. It should be noted that although burn injury promotes splanchnic hypoperfusion and a loss of gut barrier integrity, debridement of injured skin is required during the early postburn period; and burn wound debridement frequently stimulates a transient bacteremia (29–32). This debridement-related bacteremia may also contribute to the overall inflammatory responses described after major trauma.
It is clearly recognized that LPS, a component of the outer membrane of gram-negative bacteria, binds to LBP, a hepatic-derived acute-phase glycoprotein present in the systemic circulation. The LPS/LBP complex bind to the CD14–TLR4 receptor complex, which has been shown to be present not only on the cell membrane of monocytes and mononuclear cells but also on cardiomyocytes (33–35). Binding of LPS to the CD14–TLR4 receptor complex initiates an intracellular signaling cascade that is detected by MyD88 and transferred by IRAK activation of the p38 MAPK pathway (35). This interaction produces phosphorylation of the inhibitory component IκB, culminating in nuclear translocation of NF-κB and subsequent transcription and translation of TNF-α protein (15, 28). The role of LPS in mediating myocardial inflammatory responses to burn serum challenge was supported by our finding that pretreating cardiomyocytes or isolated hearts with recombinant bacterial permeability-increasing protein (rBPI) provided significant cardioprotection, decreasing myocyte inflammatory cytokine secretion, preventing myocyte Na+ and Ca2+ loading, and improving ventricular function. These data are consistent with our finding that in vivo administration of rBPI attenuated postburn contractile depression (28).
The mechanisms by which gut- or wound-derived endotoxin initiates a cardiomyocyte inflammatory cascade that culminates in myocardial contractile dysfunction remains unknown. LPS may directly stimulate cardiomyocytes by binding to the cell surface CD14–TLR4 receptor complex, initiating p38 MAPK activation, promoting NF-κB nuclear translocation and transcription of pro- and antiinflammatory cytokines. Alternatively, LPS or an endotoxin-like mediator may stimulate TNF-α secretion by liver and lung macrophages, promoting a modest rise in circulating TNF-α levels, which, in turn, could target downstream organs such as the heart. In this present study, TNF-α levels in burn serum were significantly higher than values measured in sham burn serum, supporting the hypothesis that noncardiac-derived TNF-α may promote myocyte cytokine synthesis. Therefore cardiomyocytes may serve as both a target and a source of inflammatory cytokines. In addition, TNF-α secreted locally by cardiomyocytes may act in a paracrine fashion, stimulating adjacent cardiomyocytes and exacerbating myocyte secretion of TNF-α. Alternatively, TNF-α secreted within the myocardium may serve as a proximal mediator of the myocyte inflammatory response, promoting secretion of other inflammatory mediators such as IL-1β, IL-6, and NO. Our finding that anti-TNF-α pretreatment of cardiomyocytes ablated cytokine responses triggered by burn serum challenge provided support for TNF-α as another myocardial depressant factor in burn trauma.
In summary, the present study expanded on previous findings from this Burn Center that factors present in burn serum contribute to myocardial abnormalities that are characteristic of burn trauma. The data described herein strongly implicate LPS as one proximal mediator of cardiomyocyte inflammatory cytokine secretion, myocyte calcium and sodium accumulation, and myocardial contractile depression. An endotoxin-like mediator may bind the cardiomyocyte cell surface CD14–TLR4 receptor complex; alternatively, minute amounts of endotoxin in the systemic circulation may stimulate hepatic and lung macrophage synthesis of TNF-α, which, in turn, could produce downstream organ inflammation and injury by binding TNF-α receptors shown to exist on cardiomyocytes. However, the availability of an in vitro burn-serum challenge model will allow us to further explore the mechanisms of cardiomyocyte cytokine synthesis and the myocardial contractile consequences of cardiac-derived cytokines.
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