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Published online before print April 23, 2004
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,
* Departments of Surgery/Trauma,
Pharmacology, and
Medicine, University of California San Diego
1Correspondence: Department of Surgery/Trauma, UCSD Medical Center, 200 West Arbor Drive, San Diego, CA 92103. E-mail: wjunger{at}ucsd.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: phagocyte • hyperosmotic stress • oxidative burst • inflammation • purinergic receptors
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Infusion of hypertonic saline (HS) is a novel therapeutic approach that can be used to reduce post-traumatic complications [2 ]. HS, with or without the addition of colloids such as dextran or hetastarch, can be used to resuscitate patients from hemorrhagic shock. As only small volumes (e.g., 250 ml) of such fluids are sufficient to restore circulation, initial interest in HS was primarily based on the logistical benefits of such treatment. Later, however, we found that HS resuscitation can prevent neutrophil activation in a mouse model of hemorrhagic shock [3 ]. Parallel results were obtained in a clinical trial in which HS resuscitation improved the outcome in patients, apparently by reducing neutrophil-mediated, post-traumatic complications [4 ]. The notion that HS infusion can prevent PMN activation has been confirmed by additional studies in which HS resuscitation protected host tissues from PMN-mediated damage [5 ].
Infusion of HS in patients increases plasma tonicity to levels that can suppress or enhance different PMN functions, including superoxide formation [6 7 8 9 ]. These levels of HS activate signaling pathways that are analogous to those involved in the response of yeast to hypertonic stress [10 ]. Although yeast cells have at least two known osmoreceptors that regulate responses to osmotic changes in their environment [10 ], the molecular mechanisms by which PMN and other mammalian cells detect osmotic stress are poorly understood [11 ].
We have previously reported that HS can enhance or suppress PMN functions depending on the timing of hypertonic stimulation relative to other stimuli. Enhancement is mediated by activation of the p38 mitogen-activated protein kinase (p38 MAPK), and suppression occurs via increases in cyclic adenosine monophosphate (cAMP) and activation of protein kinase A (PKA), which blocks N-formyl-methionyl-leucyl-phenylalanine (fMLP) receptor signals that promote functions such as the oxidative burst [8 , 9 ]. However, the mechanisms by which HS exerts these opposing effects are unclear. We show here that HS-induced cell shrinkage releases adenosine 5'-triphosphate (ATP) from PMN, released ATP augments PMN functions through P2 receptors and p38 MAPK activation, or ATP is converted to adenosine, which suppresses PMN functions via A2 receptors that activate cAMP/PKA signaling. This bidirectional control by released ATP thus allows PMN to register and differentially respond to osmotic changes in their extracellular environment.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Cell isolation
PMN were isolated from peripheral blood of healthy human volunteers and were stimulated exactly as described previously [8
]. Briefly, heparinized blood was mixed at a ratio of 3:1 with a dextran solution (5% w/v) in pyrogen-free, normal saline and was allowed to sediment for 30 min at room temperature. Cells in the supernatant were centrifuged, resuspended in Hanks’ buffered saline solution (HBSS), and separated with Percoll gradient centrifugation according to the manufacturer’s recommendations. The purified PMN preparation was washed twice with HBSS. Viability was >95%, as assessed with trypan blue dye exclusion, and purity exceeded 98%. This isolation procedure was chosen to avoid osmotic shock lysis that is often used to remove residual red blood cells. Cell isolation and all subsequent experiments were performed under sterile and pyrogen-free conditions.
Cell stimulation
Cells were preincubated at 37°C for 1 h before stimulation as described in the experiments below. Cells were stimulated with hypertonic saline by the addition to the culture medium of appropriate volumes of HBSS that contained an additional 1 M NaCl.
Assay of neutrophil function
O2– formation was determined with a slightly modified version of the cytochrome C reduction assay described previously [9
]. Briefly, PMN (2x106/ml) were incubated at 37°C and treated with the different reagents as described in the experiments below, then 100 µl cytochrome C (2 mM) was added, and the cells were stimulated with 100 nM fMLP for 10 min, placed on ice, and centrifuged in the cold. The optical density at 550 nm of the supernatants was used to calculate O2– formation. As a control, superoxide dismutase (50 µg/ml) was added to verify that cytochrome C reduction was the result of O2– formation. Depending on the different cell preparations and blood donors, superoxide formation could range from 36 to 98 nmol/min/2 x 108 cells in response to 100 nM fMLP and from 0 to 6 nmol/min/2 x 108 cells in unstimulated cells. Unless otherwise indicated, changes of superoxide formation in response to HS and other modulatory agents were expressed as percentage of the full response to 100 mM fMLP.
High-performance liquid chromatography (HPLC)
Freshly isolated PMN (250 µl at 107/ml) were stimulated with HS as described below for the different experiments and placed on ice, and PMN suspensions were centrifuged in the cold (325 g, 1 min). The supernatants (200 µl) were processed for analysis by addition of 1/20 vol ice-cold 8 M perchloric acid, centrifugation at 10,000 g for 5 min, adjustment of pH to 6 with 4 M K2HPO4,and another centrifugation. Then, 25 µl of these samples was subjected to HPLC analysis using a Waters system (Waters Associates, Milford, MA) consisting of two 510 pumps, a U6K injector, a tunable Waters 484 UV absorbance detector set at 254 nm, and a NEC computer system with Baseline 810 chromatography software. Ion-pair separation of nucleotides was achieved with a Supelcosil LC-18-T reverse-phase column (C18, 15 cmx4.6 mm, 3 µm particles) and Supelguard LC-18-T guard column (Supelco, Sigma-Aldrich Co.). Chromatographic conditions were as follows: buffers A (0.1 M KH2PO4, 4 mM tetrabutylammonium hydrogen sulfate, pH 6) and B (buffer A:methanol, 70:30); gradient: 0–2.5 min, 100% A; 2.5–5 min, 0–20% B; 5–10 min, 20–40% B; 10–13 min, 40–100% B; 13–18 min, 100% B; 18–18.1 min, 100–0% B; 18–22 min, 100% A; flow rate: 1.5 ml/min.
Purinergic receptor mRNA levels
Transcription of mRNA encoding the four adenosine receptors and the different P2 receptors was examined by real-time reverse transcriptase-polymerase chain reaction (RT-PCR). Total RNA was isolated from human neutrophils using Qiagen RNeasy RNA kit (Qiagen, Studio City, CA). The real-time RT-PCR was performed using QuantiTect SYBR Green RT-PCR kit (Qiagen) following the manufacturer’s protocol. Total RNA (5 ng) was used as template, and primer concentration was 0.5 µM. An initial activation step of 15 min (95°C) was performed before 49 cycles of the following: 30 s (94°C), 30 s (58°C), and 40 s (72°C). For the adenosine receptors, we used the following primer pairs: A1 (305 bp, 5'-GCCACAGACCTACTTCCACA-3' and 5'-CCTTCTCGAACTCGCACTTG-3'); A2a (216 bp, 5'-CGAATTCAACCTGCAGAACGTCACC-3' and 5'-TCGAATTCGCGGTCAATGGCGATG-3'); A2b (513 bp, 5'-CAGACGCCCACCAACTACTT-3' and 5'-GCCACCAGGAAAATCTTAATG-3'); A3 (328 bp, 5'-ACCACTCACAGAAGAATATG-3' and 5'-ACTTAGCCGTCTTGAACTCC-3'). These and the P2 receptor primer sequences and PCR conditions were based on previously published work [12
13
14
].
Purinergic receptor expression
Adenosine receptors were visualized by fluorescence microscopy as described before [15
] except that PMN were treated with 0.25 mM diisopropyl fluorophosphate (Sigma-Aldrich Co.) at 0°C for 30 min and fixed with 4% paraformaldehyde in phosphate-buffered saline at room temperature for 15 min. To allow antibodies to bind to transmembrane and intracellular receptor epitopes, cells were permeabilized with 0.5% Triton X-100 for 1 min on ice before staining with 20 µg/ml antibodies to A1, A2a, A2b, A3, P2X7 (Santa Cruz Biotechnology, Santa Cruz, CA), or P2Y2 (Zymed Laboratories, South San Francisco, CA) at room temperature. Control cells were treated identically to the antibody-stained cells, except that both antibodies were omitted, or the primary antibody was replaced by unrelated immunoglobulin G.
Intracellular cAMP measurements
Neutrophil cAMP levels were determined with an immunoassay kit (Amersham Pharmacia Biotech, Piscataway, NJ) as described before [9
]. Briefly, neutrophils (106 cells) were plated in 96-well tissue-culture plates and incubated at 37°C for 1 h. Cells were treated with receptor antagonists for 30 min, 3-isobutyl-1-methylxanthine (IBMX) was added at a final concentration of 80 µM, and cells were incubated with 20 mM HS for 10 min at 37°C. Then 20 µl of the lysis reagent supplied with the enzyme immunoassay kit was added, and cAMP levels were determined according to the manufacturer’s instructions.
Measurement of MAPK activation
To measure p38 and extracellular-regulated kinase (ERK) MAPK activation, PMN (106 cells/sample), stimulated as described below, were placed on ice, centrifuged, resuspended in 100 µl ice-cold sodium dodecyl sulfate sample buffer containing 100 mM dithiothreitol, and lysed by boiling for 5 min. The cell lysates were separated by polyacrylamide gel electrophoresis using 8–16% Tris/glycine polyacrylamide gradient gels (Novex, San Diego, CA). Proteins were transferred to polyvinylidene difluoride membranes (Immobilon-P; Millipore Corp., Bedford, MA), and the membranes were subjected to immunoblotting with phospho-specific antibodies that recognize the phosphorylated and thereby activated forms of p38 and ERK MAPKs, respectively (New England Biolabs, Beverly, MA). Antibodies recognizing the active and inactive forms of these MAPKs were used to control that equal amounts of protein were present in all samples. The appropriate antibodies were obtained from Santa Cruz Biotechnology. Secondary horseradish antibody conjugate and the enhanced chemiluminescence assay kit were from Pierce (Rockford, IL).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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S. Exposure to 100 µM ATP or ATPS rapidly increased fMLP-induced O2– formation, but although the stimulation by ATP converted to a suppression over an incubation period of >15 min, the stimulation by ATPS diminished only slightly (Fig. 2D)
. These results are consistent with the idea that extracellular ATP enhances, and metabolism of this ATP to adenosine suppresses PMN function. In Figure 2D
the conversion of the 100 µM ATP to effective concentrations of its suppressive products seemed to require 
20 min of expose to PMN. However, significant levels of suppressive ATP products appear to be produced within 10 min of exposure to HS (Fig. 2B)
. The apparent paradox between the time requirements in Figure 2B
and 2D
suggests that the local concentrations of ATP, the dynamics of ATP release, and the diffusion of ATP and its products between the cell surface and the bulk media contribute to the functional response of PMN to ATP.
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HS suppresses PMN by cAMP accumulation through A2a and A2b adenosine receptors
We have previously shown that HS rapidly increases intracellular cAMP levels by up to sevenfold [10
]. The concentration of cAMP correlated with the applied level of hypertonicity and the degree of suppression of PMN function. Based on these findings and the results shown above, we hypothesized that HS causes cAMP accumulation through the activation of A2a and A2b receptors, as both are Gs-coupled receptors. To explore this possibility, we tested if A2 receptor antagonists could prevent the activation of cAMP by HS. We pretreated PMN with the A2a receptor antagonist CSC (1 µM) or the A2b receptor antagonist enprofylline (10 nM) for 30 min, stimulated the cells with 20 mM HS in the presence of 80 µM of the phosphodiesterase inhibitor IBMX for 10 min, and measured cAMP concentration. Unstimulated cells and cells that were stimulated with the cAMP-elevating drug forskolin served as negative and positive controls, respectively. HS (20 mM) augmented cAMP accumulation to levels similar to those we have previously reported [9
]. These cAMP levels were significantly reduced by the A2a and A2b receptor antagonists CSC and enprofylline (P<0.005) and by the A2-specific antagonist DMPX (P<0.005; Fig. 6
). These data lead us to conclude that HS exerts its suppressive effect on PMN function at least in part by the accumulation of intracellular cAMP through the activation of the two A2 receptors. However, it is possible that other Gs protein-coupled receptors might be involved in the generation of cAMP. For example, very recently, a new P2Y receptor member (P2Y15) has been identified, which binds to adenosine and cAMP and results in the elevation of intracellular cAMP [24
].
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15 nucleotide receptors known to date, and these are classified as members of the G protein-coupled P2Y or ion-channel P2X groups [14
15
16
]. As HS can enhance PMN function under certain circumstances and as ATP can significantly increase fMLP-induced O2– formation (Fig. 2A)
we investigated the expression of P2 receptors in PMN. Real-time RT-PCR analysis revealed that human PMN express high levels of mRNA copies that encode the following P2 receptor subtypes (in order of relative abundance): P2Y2, P2Y4, and P2X7 and to a lesser extent, P2Y11 and P2Y6 (Fig. 7A
). We verified expression of P2X7 and P2Y2 receptors on PMN using immunofluorescence (Fig. 4)
; antibodies for the other receptors identified by PCR were not available. To test the role of P2 receptors in the enhancement of oxidative burst by ATP, we treated PMN with increasing concentrations of the P2 antagonists, suramin, o-ATP, and KN-62 for 30 min. Next we added 100 µM ATP for 10 min, followed by stimulation with 100 nM fMLP, and then we measured superoxide formation (Fig. 7B)
. All three P2 antagonists inhibited the enhancement of fMLP-stimulated O2– formation by ATP, suggesting that activation of P2 receptors is responsible for the enhancing effects of HS on PMN function. At concentrations that reversed the ATP response, these P2 antagonists had no discernible effect on superoxide formation of cells that was stimulated with fMLP in the absence of exogenous ATP (data not shown).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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A number of studies have focused on the signaling mechanisms through which osmotic stress regulates the functions of immune cells and other mammalian cells. However, the nature of the receptors that mediate recognition and response of these cells to osmotic stress has remained poorly defined. Here, we show that physiologically relevant osmotic stress causes the regulated release of ATP from PMN and that ATP and its products can increase or suppress neutrophil responses through autocrine-feedback regulation via P2 and P1 receptors. The findings complement and extend previous work that has evaluated the impact of ATP and adenosine on PMN responses [20 21 22 23 , 25 ]. It is interesting that it has been shown that the A2a adenosine receptor has a nonredundant role in the attenuation of inflammation and tissue damage in a mouse model of septic shock [26 ]. Together with these data, our findings offer an intriguing model to explain how HS can enhance and suppress PMN functions (Fig. 8 ). In this model, hypertonic stress shrinks PMN and causes the release of intracellular ATP. Adenosine, derived by conversion of the released ATP, activates A2 receptors that stimulate adenylyl cyclase activity and increase intracellular cAMP accumulation; the increase in cAMP then blocks PMN activation, presumably via PKA and PKA-promoted phosphorylation of key target proteins. Under conditions where extracellular ATP concentration is favored relative to that of adenosine, PMN functions are enhanced by P2 receptor activation and in turn, the activation of p38 and ERK MAPKs.
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The precise nature of the adenosine and P2 receptors that are involved in mediating the inhibitory and stimulatory responses observed in the response of PMN to HS will require further study. It is intriguing that PMN appear to express a large number of receptors capable of recognizing adenosine and ATP as well as other nucleotides. In spite of the higher expression of A1 and A3 mRNA, studies with antagonists suggest that A2 receptors are responsible for the suppression of PMN functions by HS. Based on abundance of mRNA and known properties for recognition of nucleotides, the stimulatory P2 receptors of PMN that respond to ATP might be P2Y2, P2X7, or perhaps P2Y11. These receptors, in particular P2X7, are known to be inhibited by the three antagonists (suramin, o-ATP, and KN-62) used in the current study and have been implicated in other immunostimulatory and proinflammatory activities [15 , 27 , 28 ].
We found relatively low concentrations of extracellular ATP with a maximum of 17 nmol/2.5 x 106 cells released in response to osmotic stimulation of PMN (Fig. 1B)
. This value is lower than the value reported by Eltzschig et al. [29
], who detected a maximum level of 215 nmol ATP released by 107 PMN in response to fMLP stimulation. These differences may be a result of the different agents used to evoke ATP release from PMN or differences in the preparation and handling of cells. According to the results shown in Figure 2A
the concentration of ATP that we and Eltzschig et al. [29] detect would modestly suppress superoxide formation. However, in addition to ATP, the cell supernatants of HS-treated cells would also contain concentrations of AMP and adenosine, which would further suppress PMN function. In addition, the concentrations of these molecules are likely to be highest in the vicinity of the cell membrane, where ATP is released and converted, and concentrations assessed in the bulk phase, as we have done here with HPLC, are considerably lower, as previously shown in other cell systems [30
]. The inverse is likely to be true for the experiments wherein we studied the effects of exogenous ATP, AMP, and adenosine on PMN responses, where the concentrations of these substances are likely higher in the supernatant than close to the cell membrane.
The model proposed in Figure 8 suggests an important role for the enzymes that catalyze the conversion of ATP to ADP, AMP, and adenosine. The relative activities and localization of these enzymes, as well as those of the P1 and P2 receptors, in relation to the sites where ATP is released represent critical parameters that may determine whether osmotic stress stimulates or suppresses PMN functions. In addition, the availability and distribution of these molecules may determine how PMN respond to ATP from other sources. Damaged and dying cells and microbes also release ATP, which may increase the recruitment of scavenging PMN to inflammatory or infectious sites and regulate PMN function through ATP and adenosine receptor activation. It is interesting that the ancient method of applying honey, sugar, salt, or other hyperosmolar agents to infected wounds could have its basis in the regulation of such defense mechanisms through the osmoreceptor system we identified in neutrophils, perhaps by augmenting neutrophil responses to the chemical signals obtained from invading microbes.
Several recent studies have shown that A2 receptor activation can be used to control inflammation in vivo [22 , 23 , 26 ]. It is intriguing to speculate that anti-inflammatory actions of hypertonic fluids could be based on the same principle of A2 receptor activation. Additional studies are required to define the exact P1 and P2 receptor types and the mechanism of their differential activation by osmotic stress. This knowledge may make it possible to therapeutically fine-tune neutrophil responses so as to increase or prevent cell activation and thereby assist in the treatment of infectious and inflammatory diseases.
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ACKNOWLEDGEMENTS |
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Received February 3, 2004; revised March 23, 2004; accepted March 25, 2004.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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) or without (•) apyrase (20 U/ml) for 30 min. The cells were then exposed to increasing concentrations of ATP for 10 min and stimulated with 100 nM fMLP, and superoxide formation was measured with a cytochrome C reduction assay. (B and C) Cells were stimulated with increasing concentrations of HS, ADP, AMP, or adenosine, and fMLP-induced superoxide formation was determined as above. (D) Cells were incubated for increasing periods of time with 100 µM ATP or the nonhydrolyzable form, ATP
