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Erythropoietin Requires Endothelial Nitric Oxide Synthase to Counteract TNF-α-Induced Microcirculatory Dysfunction in Murine Striated Muscle

Contaldo, Claudio*; Lindenblatt, Nicole*; Elsherbiny, Ahmed*; Högger, Dominik C.*; Borozadi, Meisam Khorrami*; Vetter, Sebastian T.*; Lang, Karl S.; Handschin, Alexander E.*; Giovanoli, Pietro*

doi: 10.1097/SHK.0b013e3181fd0700
Basic Science Aspects
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In the present study, we aimed to evaluate whether erythropoietin (EPO) treatment may exert nonhematopoietic endothelial protection against TNF-α-induced microvascular inflammation and to determine the involvement of the nitric oxide (NO)-producing enzyme isoforms endothelial NO synthase (eNOS) and inducible NO synthase (iNOS). Murine dorsal skinfold chambers of wild-type (WT) animals were topically stimulated with TNF-α after pretreatment with epoetin beta (1,000 IU/kg body weight i.p.) or physiological saline. Leukocyte behavior, microvascular perfusion, and apoptosis were assessed by in vivo fluorescence microscopy. To study the involvement of NO, we compared eNOS-deficient (eNOS−/−) and iNOS-deficient (iNOS−/−) mice with WT animals. TNF-α-associated leukocyte activation, perfusion failure, and apoptosis were substantially attenuated in EPO-pretreated WT mice, which was accompanied by marked reduction of perivascular infiltration with F4/80-stained macrophages. The anti-inflammatory protective effects of EPO were abolished in eNOS−/−, but not in iNOS−/− mice, both with unaffected intercellular adhesion molecule 1 expression. However, the antiapoptotic effect of EPO was maintained in both eNOS−/− and iNOS−/− mice, indicating that this mechanism might rather be independent of NO. We conclude that EPO treatment elicits protection against TNF-α-induced microcirculatory dysfunction, depending on NO derived from endothelial cells, but not on the inducible isoform.

*Division of Plastic and Reconstructive Surgery, and Institute of Experimental Immunology, Department of Pathology, University Hospital Zurich, Zurich, Switzerland

Received 3 Mar 2010; first review completed 14 Mar 2010; accepted in final form 14 Sep 2010

Address reprint requests to Claudio Contaldo, MD, Division of Plastic and Reconstructive Surgery, Department of Surgery, University Hospital Zurich, CH-8091 Zurich, Switzerland. E-mail: claudio.contaldo@usz.ch.

This research was financially supported by the Helmut Horten Foundation and the Elite-Med Foundation.

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INTRODUCTION

Erythropoietin (EPO) is a hypoxia-inducible renal glycoprotein hormone that is required for normal erythropoiesis (1). It is increasingly recognized that, beyond erythropoiesis, EPO fulfills many nonhematopoietic biological tasks that might counteract proinflammatory cytokine actions in a variety of tissue injuries (2). During vascular inflammation, TNF-α may act as a direct neutrophil chemoattractant (3, 4). Upon TNF-α stimulation, selectins initiate leukocyte rolling along the inflamed endothelium effecting integrin activation, which in turn interacts with intercellular adhesion molecule 1 (ICAM-1), mediating the development of leukocyte firm adherence (5, 6). In previous studies, exogenous EPO application has been shown to inhibit inflammatory processes entailed by injury and to downregulate proinflammatory cytokine production (7). Moreover, EPO has been observed to both directly and indirectly stimulate vascular nitric oxide (NO) and therefore reduce tissue injury, but the exact mechanism remains elusive (8, 9). Both the constitutively expressed endothelial NO synthase (eNOS) isoform, as well as the inducible NO synthase (iNOS) isoform, which is expressed only after transcriptional induction, have been shown to mediate nonhematopoietic actions of EPO (10, 11).

Our aim was to determine whether EPO treatment may exert nonhematopoietic protection of striated muscle microcirculation after TNF-α stimulation using an in vivo mouse model that allows for repetitive evaluation of the microcirculation. Furthermore, this study was designed to determine the involvement of the NO-producing enzyme isoforms eNOS and iNOS during EPO-mediated microvascular protection.

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MATERIALS AND METHODS

Animals and drugs

Twenty-eight wild-type (WT) C57BL/6J, 22 eNOS-deficient (eNOS−/−, B6.129P2-Nos3tm1Unc/J), and 22 iNOS-deficient (iNOS−/−, B6.129P2-Nos2tm1Lau-chtl/J) male mice were included in this study (a total of 72 mice, 12-18 weeks of age, 22-25 g body weight [BW]; Jackson Laboratories, Bar Harbor, Maine). The animals received humane care according to the guidelines of the University Hospital of Zurich. The study protocol was approved by the Federal Veterinary Office of the Canton of Zurich. To induce an inflammatory response, 200 μL of recombinant mouse TNF-α (Sigma-Aldrich, Buchs, Switzerland) was topically applied on the chambers at a concentration of 100 ng/mL. The animals were randomly assigned and equally distributed to three control groups and three pretreatment groups receiving EPO (Recormon; Roche, Basel, Switzerland) at a dose of 1,000 IU/kg BW i.p. Control animals received the same amount of physiological saline 0.9%.

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Experimental model

The dorsal skinfold chamber in mice was used for intravital microscopy as previously described in detail (12). Briefly, mice were anesthetized i.p. with a mixture of 90 mg/kg BW ketamine hydrochloride (Ketavet; Parke Davis, Freiburg, Germany) and 25 mg/kg BW xylazine hydrochloride (Rompun; Bayer, Leverkusen, Germany), and two symmetric titanium frames were implanted to sandwich the extended double layer of the skin. One layer was removed in a 15-mm-diameter circular area. The remaining layer consisting of epidermis, subcutaneous tissue, and striated skin muscle was covered with a glass coverslip, incorporated in one of the titanium frames. Animals tolerated the chamber well and showed no signs of discomfort or changes in sleeping and feeding habits. A recovery period of 3 days was allowed before the experiment was started.

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Intravital fluorescence microscopy

Microhemodynamic measurements were performed using an epi-illumination intravital microscope (Leica DM/LM; Leica Microsystems, Wetzlar, Germany) attached to a blue (excitation/emission wavelength, 450-490/>520 nm), a green (excitation/emission wavelength, 530-560/>580 nm), and an ultraviolet (excitation/emission wavelength, 330-390/>430 nm) filter system. Microscopic images were captured by a television camera (intensified charge-coupled device camera; Kappa Messtechnik, Gleichen, Germany), displayed on a television screen (Trinitron PVM-20N5E; Sony, Slough, UK), and recorded on video (50 Hz; Panasonic AG-7350-SVHS, Tokyo, Japan) for subsequent off-line analysis. The preparation was observed visually with a water-immersion objective ×20 with a numerical aperture of 0.50, which resulted in a total optical magnification of ×800 on the video monitor. Animals received a tail vein injection of 0.05 mL fluorescein isothiocyanate-dextran (50 mg/mL saline) (Sigma-Aldrich) for vascular contrast enhancement and 0.05 mL rhodamine 6G (0.1 mg/mL saline) (Sigma-Aldrich) for leukocyte staining in vivo. Nuclei of tissue cells were visualized in vivo by topical application of 0.1 mL bisbenzimide Hoechst 33342 (1 mg/mL saline) (Sigma-Aldrich).

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Quantitative microcirculation analysis

The chamber was at first observation scanned for random selection of distinct observation areas, which included four to six second- and third-order arterioles, nine nutritive capillary fields, and four to six draining postcapillary venules. Video printouts were made during videography and initially marked to indicate the exact localization for measurements of vessel diameter and red blood cell (RBC) velocity. Using a computer-assisted image analysis system (CapImage; Zeintl Software, Heidelberg, Germany), functional capillary density (FCD) was assessed as the length of RBC-perfused capillaries per observation area (in centimeters per centimeter squared) (12). Diameters were measured in micrometers perpendicularly to the vessel path. Centerline RBC velocity was analyzed by computer assistance using the line-shift method (CapImage). Volumetric blood flow was calculated from diameter (d) and RBC velocity (v) by Q = π * (d / 2)2 * v / 1.6 (pl/s), where 1.6 represents the Baker-Wayland factor to correct for the parabolic velocity profile in microvessels with diameters of greater than 20 μm (13). The number of permanent adherent leukocytes (defined as cells that adhered to the venular vessel wall over a period of 30 s) was evaluated as number of cells per millimeter squared of endothelial surface (calculated from diameter and length of the vessel segment studied, assuming cylindrical geometry). Rolling leukocytes were defined as cells moving with a velocity less than two fifths of the centerline velocity and are given as the number of cells per minute passing a reference point within the microvessel (14). Topical application of the vital dye bisbenzimide Hoechst 33342 allowed us to assess in vivo cell nuclear morphology with apoptosis-associated condensation, fragmentation, and crescent-shaped formation of chromatin (15). Using the ×40 objective for recording (magnification ×1,500), cells exhibiting these apoptotic characteristics were counted and are given as number per observation field.

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Immunohistochemistry

Histological analyses were performed on snap-frozen tissue. Sections were stained with rat monoclonal antibodies against F4/80 (1:800; BMA Biomedicals, Augst, Switzerland) and against murine ICAM-1 (1:600; AbD Serotec, Oxford, UK). Bound F4/80 antibody was detected using a secondary goat anti-rat antibody (1:150; Caltag Laboratories, Carlsbad, Calif), and bound ICAM-1 was detected using a secondary goat anti-hamster antibody (1:200; Jackson ImmunoResearch Laboratories), both with a tertiary alkaline phosphatase-coupled donkey anti-goat antibody (1:80; Jackson ImmunoResearch Laboratories) and new fuchsine as a substrate. The sections were counterstained with hemalaun. The intensity of the staining reactions of ICAM-1 in endothelial cells was evaluated by a Zeiss Axioplan 2 imaging system at a magnification of ×100 (Carl Zeiss, Oberkochen, Germany) using a semiquantitative score (graded as 0 = no, 1 = weak, 2 = moderate, and 3 = strong staining). F4/80-positive cells were counted in 10 randomly selected visual fields at a magnification of ×40. Photographs were taken with a digital imaging system (AxioCam; Carl Zeiss; Jena, Germany).

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Immunoblotting

Dorsal skinfold chamber tissues were surgically separated in skin and panniculus carnosus containing striated muscle. Aliquots of cell lysates from skin and muscle samples (150 mg of protein) were loaded on an 8% sodium dodecyl sulfate-polyacrylamide gel electrophoresis gel and transferred to a polyvinylidene difluoride membrane (Millipore; Bedford, Mass). Membranes were blocked with 5% milk in Tris-buffered saline-Tween buffer (20 mM Tris-HCl, pH 7.4; 137 mM NaCl; 0.1% Tween). Blocked membranes were incubated with primary antibody overnight at 4°C followed by secondary antibody (horseradish peroxidase-linked whole antibody; 1:1,000; GE Healthcare; Buckinghamshire, UK). Membranes were then incubated with enhanced chemiluminescence reagent (Millipore; Billerica, Mass) and exposed to x-ray films (Fuji Super RX Medical x-ray films; Fuji Photo Film, Düsseldorf, Germany) for the recommended optimal time, depending on the signal strength. As primary antibodies we used anti-phosphorylated-eNOS (Ser-1177) (1:300; Cell Signaling Technology Inc, Beverly, Mass); anti-phospho-eNOS (Thr-495) (1:500; BD Transduction Laboratories; San Jose, Calif), and anti-eNOS/NOS type III (1:500; BD Transduction Laboratories). Primary human aortic endothelial cells (Clonetics, Allschwil, Switzerland) from passages 4 to 6 were used. The cells were cultured and passaged in EBM-2 medium supplied with EGM-2 BulletKit (Clonetics, Walkersville, Md). Blots of total eNOS were analyzed using an appropriate software (Advanced Image Data Analyser AIDA, Raytest, Berlin, Germany).

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Experimental protocol

Systemic EPO pretreatment was performed 1 h before TNF-α exposure. After baseline (BL) microscopy, chambers were exposed to TNF-α to induce a sustained and marked inflammatory response. Microscopic recordings with microcirculatory analysis were repeated 15, 30, 60, and 120 min, as well as 24 h after TNF-α exposure. Animals were killed with an i.p. overdose of the anesthetic mixture. The study comprised six different groups (n = 6 per group). Wild-type, eNOS−/−, and iNOS−/− mice were exposed to TNF-α after pretreatment with either EPO or saline. Five additional animals of each group were killed 30 min after TNF-α exposure to study ICAM-1 expression and eNOS expression in small arterioles and to assess perivascular accumulation of F4/80-positive cells using immunohistochemistry. Three additional animals of each pretreated and control WT group were killed 30 min after TNF-α exposure to study phosphorylation status of eNOS in cell lysates using immunoblotting.

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Statistical analysis

All values are expressed as means ± SEM. For comparison between individual time points within each group, ANOVA was performed followed by the paired Student t test, including correction of the α error according to Bonferroni probabilities for repeated measurements. Differences between groups were evaluated using the unpaired ANOVA with Tukey post hoc test (GraphPad Prism 4; GraphPad Software, La Jolla, Calif). A P value of less than at least 0.05 was considered to be statistically significant.

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RESULTS

Baseline microcirculation data are summarized in Table 1. Note that there were no significant differences between the different animal groups and between control animals and EPO-pretreated animals before TNF-α exposure.

TABLE 1

TABLE 1

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Leukocyte-endothelial cell interaction after TNF-α stimulation

In control WT animals, TNF-α exposure rapidly increased leukocyte rolling and adherence to the venular endothelial lining, indicating a pronounced inflammatory response, which progressively normalized during the 24-h observation period (Fig. 1). Upon pretreating WT animals with EPO, we observed a substantially dampened TNF-α-associated venular leukocyte-endothelial cell interaction, indicating successful attenuation of TNF-α-induced microvascular inflammation. To differentiate between the contribution of endothelial-derived NO and inducible NO, additional genetically depleted animals were exposed to TNF-α and pretreated with either vehicle solution or EPO (Figs. 2 and 3). Animals knocked out of eNOS showed a marked inflammatory response upon TNF-α exposure (Fig. 2) comparable to that of WT animals. When EPO-pretreated eNOS−/− animals were exposed to TNF-α, the inflammatory response was completely unaffected. TNF-α-exposed iNOS−/− animals revealed matchable high numbers of rolling leukocytes and even higher counts of leukocyte adherence, when compared with WT animals (Fig. 3). When iNOS−/− animals were pretreated with EPO, we observed a reduced TNF-α-associated inflammatory response comparable to that in WT animals.

Fig. 1

Fig. 1

Fig. 2

Fig. 2

Fig. 3

Fig. 3

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Apoptosis and FCD after TNF-α stimulation

TNF-α exposure caused a decrease in nutritive tissue perfusion in WT animals, as given by the FCD demonstrating an average value of only 68% of BL at the end of the 24-h observation period (Fig. 4). Finally, TNF-α-associated apoptotic tissue injury was indicated by a ∼5-fold increase in detached cells with nuclear fragmentation (anoikis) and greater than 25-fold increase in nuclear condensation (apoptosis). Erythropoietin pretreatment almost completely abolished apoptotic tissue injury, which was paralleled by a marked restoration of nutritive perfusion. In eNOS−/− animals, the reduction of nutritive perfusion to 80% of BL upon TNF-α exposure was less pronounced but corresponded well with that of WT animals. Accordingly, TNF-α-associated apoptotic tissue injury was indicated by a ∼2-fold increase in anoikis and greater than 30-fold increase in apoptosis. In parallel to the completely unaffected inflammatory response, EPO treatment of eNOS−/− TNF-α-exposed animals failed to normalize microcirculatory dysfunction but could limit cell apoptosis. TNF-α-exposed iNOS−/− animals revealed a decrease in nutritive tissue perfusion to a value of 88% of BL at the end of the 24-h observation period. Upon EPO treatment, however, there was a complete restoration of FCD to BL levels observed in control iNOS−/− animals. In contrast to WT animals and eNOS−/− animals, in iNOS−/− control animals, apoptotic cell death upon TNF-α exposure was found to be almost completely prevented.

Fig. 4

Fig. 4

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Immunoblotting

Erythropoietin pretreatment resulted in eNOS phosphorylation at residue Ser-1177 and dephosphorylation at residue Thr-495 (Fig. 5A, c and e) in both muscle tissue and skin of the dorsal skinfold. Endothelial NO synthase expression was similar in EPO-pretreated as well as in untreated control animals (Fig. 5A, a-e).

Fig. 5

Fig. 5

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ICAM-1 and F4/80 expression after TNF-α stimulation

Expression of ICAM-1 was distinct in the endoluminal aspects of endothelial cells in sham animals (data not shown). Thirty minutes after TNF-α exposure, however, the entire cytoplasm showed substantial ICAM-1 expression in all groups (Fig. 6A; Fig. 7, A and B). In line with this, F4/80 staining revealed abundant accumulation of activated macrophages in the surrounding perivenular tissue, which was almost completely prevented in EPO-pretreated WT and iNOS−/− animals, but not in eNOS−/− animals (Fig. 6B; Fig. 7, C and D).

Fig. 6

Fig. 6

Fig. 7

Fig. 7

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DISCUSSION

The present study demonstrates that microvascular dysfunction as induced by TNF-α stimulation is considerably mitigated after systemic pretreatment with 1,000 IU/kg BW of EPO i.p. This beneficial effect of EPO was absent in eNOS−/− but not in iNOS−/− mice, which strongly suggests that NO derived from endothelial cells plays a crucial role in EPO-mediated microvascular protection.

Nonhematopoietic vasculoprotective effects of EPO are increasingly recognized. An essential function of EPO for the endothelial system is already evident during embryogenesis (16). Erythropoietin-receptor knockout mice showed severely affected angiogenesis and endothelial cell apoptosis, and most of them died in utero (17). The physiological endothelial cell response to endogenously produced EPO includes proliferation, progenitor cell migration, and production of NO (18-20). Hence, the concept of exogenous EPO administration arose to achieve vasculoprotection in a variety of pathological settings, such as ischemia-reperfusion (I/R), trauma, and inflammation (21). Increased bioavailability of NO therefore seems to represent a key factor for the EPO-mediated vasculoprotective effects (22). Although the exact mechanism for increased NO activity remains elusive, other studies suggest that the rapid increase in eNOS activity after EPO stimulation might be associated with enhanced eNOS phosphorylation and/ or protein kinases activation.

In the present study, inflammatory alterations after TNF-α stimulation included the rapid recruitment of leukocytes, which was paralleled by upregulation of ICAM-1 and perivascular infiltration with F4/80-positive stained macrophages. One single shot of 1,000 IU/kg BW of EPO i.p., administered before TNF-α stimulation, markedly reduced the inflammatory response in WT animals. Leukocyte-endothelial cell adherence and subsequent migration of leukocytes into the perivascular tissue during inflammation are coordinated by the interplay of multiple signal molecules (23). In our study, we observed a significant reduction of TNF-α-induced ICAM-1 expression after EPO treatment, which was paralleled by a substantially decreased number of both rolling and firmly sticking leukocytes. The expression of ICAM-1 is advocated as a marker for leukocyte rolling and adherence to the endothelial lining and is known to be regulated by the production of NO (24, 25). However, there are contradicting data to be found in prior research reports suggesting that EPO's pleiotropic functions might be triggered via multiple mechanisms either dependent or independent of NO. In our own earlier studies in which we investigated critically ischemic tissues, EPO pretreatment reduced leukocyte transmigration by approximately 75% and increased partial tissue oxygen tension in the ischemic tissue from 8.2 to 15.8 mmHg, which was paralleled by a 2.2-fold increase in eNOS protein expression (12). Of interest, l-NAME completely abolished all the beneficial effects of EPO pretreatment. Nevertheless, we could not definitively attribute the beneficial anti-inflammatory effect to eNOS, because l-NAME is an unspecific NO synthase blocker, and the involvement of iNOS could not be ruled out ultimately. To define, therefore, the role of NO during EPO-mediated dampening of microvascular inflammatory response in the setting of TNF-α stimulation, we compared eNOS−/− and iNOS−/− mice with WT animals. The effect of EPO on ICAM-1 expression was absent in both eNOS−/− and iNOS−/− mice, whereas leukocyte rolling and sticking remained merely unaffected in EPO-treated eNOS−/− animals compared with saline treatment. This leads to the conclusion that endothelial NO, but not the inducible form, might be the crucial protein in EPO-mediated anti-inflammatory effects. However, after EPO treatment, iNOS−/− mice in the present study exhibited reduced leukocyte trafficking, comparable to that of WT animals, whereas ICAM-1 was clearly not downregulated. Thus, ICAM-1 modulation may not be the only factor in the counteraction of proinflammatory cytokines. Although not elucidated in our experiments, one possible explanation for the EPO-mediated anti-inflammatory effect of NO might be the suppression of leukocyte recruitment elicited by mast cell activation. Mast cells are located in proximity to the microvasculature and release numerous mediators associated with acute microvascular inflammation (26). Mast cells have been implicated as being regulated by NO. The inhibition of endogenous NO synthesis with l-NAME has been shown to increase mast cell degranulation and subsequently leukocyte infiltration (27). Thus, it can be assumed that, in the present study, EPO might have suppressed leukocyte-endothelial cell interaction as elicited by mast cell activation.

In our study, microvascular alterations after TNF-α stimulation were furthermore characterized by the reduction of nutritive perfusion, as assessed by the FCD. Erythropoietin treatment improved microcirculation by restoring capillary perfusion and normalizing FCD in the presented study. This goes along with observations by other authors, who were able to demonstrate that EPO improves capillary density in a sepsis model of cecal ligation (28). In our study, improved capillary functionality after EPO treatment was abolished in eNOS−/− but not iNOS−/− mice, emphasizing endothelial-derived NO as the main factor.

Erythropoietin has been reported to stimulate NO production from vascular endothelial cells particularly at reduced oxygen levels in vitro (26). Although, we can assume normoxic conditions in the setting of the present study, this is in line with another previous in-house study, where EPO effectively attenuated musculoskeletal I/R injury by preserving nutritive perfusion, which was associated with the upregulation of eNOS and EPO-receptor protein expression (14). The mechanism of capillary perfusion failure is not fully understood. In the setting of I/R, decreased capillary patency is believed to be caused by leukocytic inflammation (29). This view is supported by studies in which abrogation of venular leukocyte adherence by monoclonal antibodies directed against CD11/CD18 showed that capillary perfusion failure could be reversed (30). Taking into consideration the distinctive roles of NO within the microcirculation, we might deduce from the present study that EPO protects capillary perfusion by two NO-mediated ways of action, namely, by regulating the vascular tone and through an anti-inflammatory effect.

In the present study, EPO treatment significantly dampened anoikis and apoptosis during acute inflammation. Interestingly, the antiapoptotic effect was not absent in eNOS depleted mice, indicating that the antiapoptotic effects of EPO might be mediated only in part by NO. TNF-α triggers apoptosis by limiting the spread of a noxious stimulus. Erythropoietin is well known to counteract apoptotic processes, be it during physiological erythropoiesis or after metabolic stress (31). However, other studies report that the antiapoptotic effect of EPO is eNOS mediated (32). On the other hand, NO has been shown to enhance and also to suppress apoptosis (32-34). The proapoptotic effect seems to be linked to pathological conditions, such as inflammation with production of large amounts of NO by iNOS. In contrast to this, the physiological continuous low production and release of NO by eNOS have been shown to prevent apoptosis (31, 32). Therefore, the supposition is valid that, regarding TNF-α-induced inflammation, proapoptotic and antiapoptotic processes may be balanced through the involvement of NO. Thus, apoptosis may be predominantly determined by other mechanisms under these conditions.

Several studies have shown that the phosphorylation status of eNOS has important implications for its enzymatic activity and biological function (35). In line with this, our study demonstrates that the specific phosphorylation at residue Ser-1177 as well as the specific dephosphorylation at residue Thr-495 in EPO-stimulated animals might be associated with the increase in eNOS activity, without significant changes in total eNOS expression.

In conclusion, our data demonstrate that EPO administration before onset of microvascular inflammation offers significant protection against damage within the microcirculation. The main mechanism of this anti-inflammatory effect seems to be linked to eNOS. Furthermore, the findings of the present study might provide new insight regarding basic understanding of EPO's microvascular effects, which may help define biological end points for potential future clinical evaluation.

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ACKNOWLEDGMENTS

The authors thank Mrs Yi Shi and Mr Thomas Lüscher from the Institute of Physiology and Center for Integrative Human Physiology (ZIHP) at the University of Zurich, Switzerland, for assistance during performance of immunoblots.

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Keywords:

Intravital microscopy; apoptosis; NO synthase; functional capillary density; leukocyte-endothelial interaction

© 2011 by the Shock Society