Platelets play important roles in the late phase of the immediate hypersensitivity reaction

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Platelets play important roles in the late phase of the immediate hypersensitivity reaction Risa Tamagawa-Mineoka, MD, PhD, Norito Katoh, MD, PhD, and Saburo Kishimoto, MD, PhD Background: Recent studies have shown that platelets have a role in most inflammatory reactions, but involvement of platelets in the immediate hypersensitivity reaction (IHR) in skin has not been examined. Objective: To investigate the role of platelets in a mouse model of IgE-mediated IHR. Methods: Mice were sensitized by injecting ovalbumin intraperitoneally and challenged by injecting ovalbumin intradermally into ears, with or without platelet depletion. Results: Sensitized mice developed biphasic responses characterized by early-phase and late-phase reactions (LPRs). Degranulation of mast cells in skin did not differ between platelet-depleted mice and controls. The early phase reaction was not suppressed at 1 hour, but platelet depletion significantly reduced the LPR at 24 hours (P < .01). Flow cytometry showed that P-selectin expression on platelets and the number of platelet-leukocyte aggregates were both higher in the blood of ovalbumin-challenged mice compared with sham-sensitized mice at 24 hours (P < .05). In platelet-depleted mice, the LPR was restored by infusing platelets from normal mice (P < .01). This effect did not occur by infusing platelets from P-selectin– deficient mice or by pretreating platelets with anti–P-selectin antibody. Injection of activated platelet supernatant into ears led to increased leukocyte infiltration at 24 hours, and this effect was blocked by pretreating the supernatant with several antichemokine antibodies. Systemic administration of antiplatelet compounds also suppressed the LPR significantly. Conclusion: These results show that platelets play important roles in the LPR of the IHR in skin by forming plateletleukocyte complexes via P-selectin in blood and secreting several chemokines that attract leukocytes to skin. (J Allergy Clin Immunol 2009;123:581-7.)

From the Department of Dermatology, Kyoto Prefectural University of Medicine Graduate School of Medical Science. Supported by a grant from the Japanese Ministry of Education, Science, Sports, and Culture (to R.T.-M. and N.K.), by a grant for Basic Dermatological Research from Shiseido Co Ltd (to R.T.-M.), and by a grant from the Cosmetology Research Foundation (to N.K.). Disclosure of potential conflict of interest: The authors have declared that they have no conflict of interest. Received for publication July 14, 2008; revised November 22, 2008; accepted for publication December 24, 2008. Reprint requests: Risa Tamagawa-Mineoka, MD, PhD, or Norito Katoh, MD, PhD, Department of Dermatology, Kyoto Prefectural University of Medicine Graduate School of Medical Science 465, Kajii-cho, Kawaramachi-Hirokoji, Kamigyo-ku, Kyoto 602-8566, Japan. E-mail: [email protected] or [email protected]. 0091-6749/$36.00 Ó 2009 American Academy of Allergy, Asthma & Immunology doi:10.1016/j.jaci.2008.12.1114

Kyoto, Japan

Key words: Atopic dermatitis, chemokine, immediate hypersensitivity reaction, late-phase reaction, platelet, P-selectin

Platelets are well known to facilitate clotting of blood to produce hemostasis and thrombosis, but lately several reports have suggested that platelets also affect induction and maintenance of inflammatory reactions.1-8 On activation, platelets release secretory products and express immune receptors on their membrane, with resultant induction of biological activities including cell adhesion, chemotaxis, cell survival, and proliferation, all of which increase inflammatory reactions.1 Involvement of platelets has also been shown in inflammatory disorders such as asthma, arthritis, or inflammatory bowel disease.2-8 For example, platelets are strongly activated in circulating blood and accumulate at sites of inflammation in patients with asthma or inflammatory bowel disease,2,7,8 and have been shown to contribute to bronchial hyperresponsiveness and leukocyte infiltration in inflamed lesions in animal models of asthma.2-4 Atopic dermatitis (AD) is usually associated with high serum IgE levels, a positive immediate hypersensitivity reaction (IHR) to food and/or environmental antigens, and eosinophilia.9 When patients with AD are challenged with an antigen against which they are sensitized, they may develop biphasic responses characterized by early-phase and late-phase reactions (LPRs). Mast cells are activated by cross-linking of FceRI-bound IgE with antigens and release mediators such as histamine and leukotrienes that induce the early-phase reaction within minutes via a rapid increase in capillary permeability. The LPR develops within hours after this response and is produced by accumulation of activated inflammatory cells, including eosinophils and neutrophils.10,11 Inflammation associated with the LPR is of clinical importance, because it accounts for the morbidity and severity of allergic disorders such as AD. We have recently shown that platelets are necessary for leukocyte recruitment in chronic skin inflammation in a murine model of chronic contact dermatitis, through formation of platelet-leukocyte aggregates via P-selectin in blood and secretion of chemokines at the inflamed lesion.12 In addition, platelets are important for cutaneous inflammation through direct activation of local vascular capillary endothelial cells and attraction of effector T cells into the tissue.13-17 Human and murine platelets have constitutive functional expression of the high-affinity and low-affinity receptors for IgE (FceRI and FceRII/CD23, respectively),3,18,19 suggesting that platelets may release mediators after antigen-specific activation via IgE antibody and increase the IHR. However, the in vivo function of platelets in the IHR in skin has not been examined. In the current study, we investigated the effects of platelet depletion and restoration of the IHR in skin using a mouse model of the IgE-mediated IHR. We then evaluated the effects of antigen exposure on P-selectin expressed on platelets and formation of 581

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Abbreviations used AD: Atopic dermatitis APAS: Antiplatelet antiserum CS: Control serum IHR: Immediate hypersensitivity reaction LPR: Late-phase reaction MIP-1a: Macrophage inflammatory protein 1a PAF: Platelet-activating factor RANTES: Regulated upon activation, normal T-cell expressed and secreted TARC: Thymus and activation-regulated chemokine TRAP: Thrombin receptor activating peptide

platelet-leukocyte complexes in blood, and the effects of deleting and blocking platelet P-selectin on leukocyte recruitment into skin. The effects of mediators released from activated platelets on leukocyte recruitment into skin were also determined in the mouse model, and we examined whether systemic administration of antiplatelet compounds to mice suppresses leukocyte recruitment into skin.

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some experiments, the activated platelet supernatant was treated with 10 mg/ mL antibody against macrophage inflammatory protein 1a (MIP-1a; clone 39624, rat IgG; R&D Systems, Minneapolis, Minn), regulated upon activation, normal T-cell expressed and secreted (RANTES; polyclonal rabbit IgG; ProSci, Poway, Calif), and/or thymus and activation-regulated chemokine (TARC; polyclonal goat IgG; R&D Systems) for 30 minutes at 378C to neutralize the chemokines. Isotype-matched IgG served as a control. After centrifugation, 50 mL activated platelet supernatant was injected into the ears of APAS-treated or untreated mice just before challenge or without challenge. A supernatant of platelets fixed with 2% paraformaldehyde without stimulation with TRAP served as a control.

Administration of antiplatelet compounds Aspirin (100 mg/kg; Sigma-Aldrich) or vehicle was administered intravenously to mice 3 hours before the experiments. In separate experiments, mice were dosed orally with clopidogrel (30 mg/kg/day; Sigma-Aldrich) or vehicle for 2 days. Responses to ovalbumin were elicited at 1 hour after the final dose.

Statistical analysis Data from all studies are expressed as means 6 SDs. Statistical significance was assessed by using 1-way ANOVA, followed by a Tukey multiple comparison test. A P value of less than .05 was considered significant.

METHODS For additional material, see the Online Repository for this article at www.jacionline.org.

Immunization The biphasic allergic murine skin model was produced as described previously.20 Mice were sensitized by intraperitoneal injection of 3 mg ovalbumin (Sigma-Aldrich, St Louis, Mo) and 4 mg aluminum hydroxide gel. Reactions to ovalbumin were elicited by intracutaneous injection of 10 mg ovalbumin into the right ear of mice 2 weeks after sensitization, with an identical amount of vehicle injected into the left ear as a control. The ear swelling response was measured 1 hour and every 6 hours after challenge and was defined as the difference in ear thickness before and after challenge. At 1 hour and 24 hours after challenge, a 5-mm-diameter biopsy of the ear was taken and weighed as an index of tissue swelling.

Platelet depletion Mice were injected intramuscularly with 0.1 mL antiplatelet antiserum (APAS) or control serum (CS), or were injected intraperitoneally with 0.05 U neuraminidase21 (Type VI; Sigma-Aldrich) or vehicle 24 hours before challenge. Alternatively, mice received busulfan22 (25 mg/kg; Sigma-Aldrich) or vehicle intraperitoneally on days –12 and –9 before challenge.

Injection of platelet suspension into the blood of mice Platelet suspension or vehicle was injected intravenously into APAStreated or untreated mice 20 minutes before challenge. The injection volume of 0.1 mL contained 7.5 to 8.0 3 108 platelets. Addition of platelets resulted in an increase of the circulating platelet population to 60% to 70% of the control level in APAS-treated mice and 150% to 160% that in APAS-untreated mice 20 minutes after injection of the platelet suspension. In some experiments, the suspension was treated with 100 mg rat anti–P-selectin blocking antibody (clone RB40.34; Becton Dickinson, San Diego, Calif). Isotype-matched IgG served as a control.

Injection of platelet supernatant into the skin Washed platelets (1 3 109/mL) were stimulated with 20 mmol/L thrombin receptor activating peptide (TRAP; Sigma-Aldrich) for 15 minutes at 378C. In

RESULTS Platelet depletion Mice were depleted of about 90% of circulating platelets by administration of APAS, busulfan, or neuraminidase (see this article’s Table E1 in the Online Repository at www.jacionli ne.org). The number of circulating leukocytes was statistically unaffected in all APAS-treated, busulfan-treated and neuraminidase-treated mice. Administration of these agents did not affect proliferation and chemotaxis of mouse T cells or intercellular adhesion molecule 1 and vascular cell adhesion molecule 1 expression on murine skin endothelial cells and human umbilical vein endothelial cells (data not shown). IgE production Administration of APAS, busulfan, or neuraminidase did not affect the serum levels of total IgE antibody and antiovalbumin IgE antibody at 14 days after sensitization (see this article’s Table E2 in the Online Repository at www.jacionline.org). These data show that the platelet-depletion treatment had no effect on the process of sensitization. Reduction of the LPR in platelet-depleted mice The time course of ear swelling responses was assessed by measuring ear thickness and weight. The increase of ear thickness was significant 1 hour and 24 hours after challenge. These reactions did not occur in mast cell-deficient WBB6F1-KitW/ KitW-v (W/Wv) mice (see this article’s Fig E1, A, in the Online Repository at www.jacionline.org). Platelet depletion by APAS (Fig 1, A), busulfan (see Fig 1, B), or neuraminidase (see Fig E1, B) suppressed the LPR, although the early-phase reaction was unaffected. The ear weight increased in all APAS-treated, busulfan-treated, and neuraminidase-treated mice and in controls in the early phase (see Fig 1, C), but platelet depletion suppressed the ear weight increase in the LPR (see Fig 1, D). A previous report23 suggested that a single dose of APAS might form sufficient

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FIG 1. Changes in ear swelling responses after ovalbumin (OVA) challenge. Ear thickness was measured in the ears of mice (6 per group) after administration of APAS or CS (A), or busulfan or vehicle (B). The ear weight was examined 1 hour (C) and 24 hours (D) after challenge. **P < .01 vs CS or vehicle administration in OVA-challenged mice. NS, Not significant.

platelet-antiplatelet IgG antibody complexes to disturb the IHR. To avoid this effect, daily injection of APAS was performed for 3 days until ovalbumin challenge, which also caused suppression of the LPR without inhibition of the early-phase reaction (see Fig E1, C). Histologic examination of the ear skin revealed edema in the dermis in the early phase, which did not differ between APAStreated mice (Fig 2, D) and controls (see Fig 2, B). In contrast, in the LPR leukocyte infiltration markedly decreased in APAS-treated mice (see Fig 2, H) compared with controls (see Fig 2, F). To quantify leukocyte infiltration into skin in the LPR, the myeloperoxidase and eosinophil peroxidase activities in the ear skin were determined. The myeloperoxidase and eosinophil peroxidase activities in APAS-treated mice were significantly lower than those in the controls (see Fig E1, D). Repeated ovalbumin challenge 3 times every 48 hours was performed to investigate the effect of platelet depletion on priming for the IHR, and this gave results

FIG 2. Histologic examination of ear skin. Skin tissue was excised 1 hour (A-D) and 24 hours (E-H) after ovalbumin challenge. A and E, Injection of vehicle alone with CS administration. B and F, Injection of ovalbumin with CS administration. C and G, Injection of vehicle alone with APAS administration. D and H, Injection of ovalbumin with APAS administration. Hematoxylin and eosin. Original magnification 3200. Bar 5 50 mm.

similar to the increase of ear weight in mice challenged once with ovalbumin (see Fig E1, E). The effect of platelet depletion on mast cell activation in the IHR was also examined, but the distribution and degranulation of mast cells in skin did not differ significantly between APAS-treated mice and controls 1 hour and 24 hours after challenge (see this article’s Table E3 in the Online Repository at www.jacionline.org). These results show that platelet depletion significantly suppresses leukocyte infiltration into skin in the LPR, and that platelet depletion has no effect on mast cell activation in skin.

Effect of platelet restoration on ear swelling and leukocyte recruitment into skin To confirm that platelets are involved in induction of the LPR, the effects of platelet restoration were examined in plateletdepleted mice. A platelet suspension obtained from ovalbuminsensitized or sham-sensitized mice was injected intravenously into APAS-treated mice before challenge. The ear swelling

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FIG 3. Effect of platelet restoration on ear swelling responses and leukocyte infiltration. Mice (6 per group) were injected intravenously with platelet suspension or vehicle. Ear swelling responses (A) and the number of infiltrated cells (B) were examined. Data are shown as percentages of the basal number of cells in the dermis 24 hours after challenge **P < .01 vs mice injected with vehicle. OVA, Ovalbumin.

response increased significantly in the LPR after platelet restoration compared with that under thrombocytopenic conditions (Fig 3, A). In addition, the levels of infiltrated cells in skin increased markedly in the LPR after platelet restoration (see Fig 3, B). Injection of platelet suspension into mice that were not platelet-depleted caused a greater increase of the LPR (see this article’s Fig E2, A, in the Online Repository at www.jacion line.org). Use of ovalbumin-sensitized or sham-sensitized mice as donors did not change the results. Therefore, to investigate the in vitro effect of platelet sensitization, platelets obtained from ovalbumin-sensitized or sham-sensitized mice were incubated in the serum of ovalbumin-sensitized mice, followed by ovalbumin-challenge. Chemokine release increased in platelets of ovalbumin-sensitized mice, compared with those of sham-sensitized mice (see Fig E2, B). These findings indicate that platelets are involved in the LPR in skin, and that platelet sensitization is not related to the in vivo IHR in skin.

Requirement for platelet P-selectin in leukocyte recruitment into skin Leukocyte recruitment from circulating blood to sites of inflammation is regulated by adhesion molecules, including Pselectin.1 Therefore, we examined the expression of P-selectin on platelets and the presence of platelet-leukocyte complexes in the blood of mice 24 hours after ovalbumin challenge. Expression of P-selectin on blood platelets markedly increased in ovalbuminchallenged mice compared with sham-sensitized mice (Fig 4, A). In addition, a significant increase in the percentage of leukocytes positive for the platelet-specific marker CD41 was found in blood obtained from ovalbumin-challenged mice compared with sham-sensitized mice (see Fig 4, B). To examine the role of platelet P-selectin on leukocyte recruitment into skin, P-selectin–

FIG 4. Necessity of platelet P-selectin for cutaneous leukocyte recruitment. P-selectin expression on platelets (A) and the number of platelet-leukocyte complexes (B) were examined. Platelets isolated from P-selectin knockout (P-selectin–/–) mice (C) or treated with anti–P-selectin antibody (D) were injected. Data are shown as percentages of basal values in sham-sensitized mice or in mice injected with vehicle (5 mice per group). *P < .05; **P < .01. OVA, Ovalbumin; WT, wild-type.

deficient platelets were injected intravenously into APAS-treated mice before challenge. Injection of platelets obtained from Pselectin knockout mice (see Fig 4, C) or platelets incubated with anti–P-selectin antibody (see Fig 4, D) significantly suppressed leukocyte infiltration into skin, whereas injection of platelets obtained from wild-type mice or platelets incubated with a control IgG significantly increased leukocyte infiltration into skin. Injection of platelet suspension into APAS-treated mice significantly increased the percentage of platelet-leukocyte complexes, whereas injection of P-selectin–deficient platelets did not increase this percentage (see this article’s Fig E3, A, in the Online Repository at www.jacionline.org). Chemokine levels released from P-selectin–deficient platelets decreased slightly but insignificantly compared with those from normal platelets (see Fig E3, B). These results indicate that antigen challenge to skin induces P-selectin expression on platelets and formation of leukocyte-platelet complexes in blood. Moreover, the results show that platelet P-selectin plays an important role in leukocyte recruitment into skin.

Effect of platelet-derived soluble factors on leukocyte recruitment into skin We assessed the ability of platelet-derived soluble factors to attract leukocytes into skin by injecting activated platelet supernatant into the ear skin of APAS-treated mice before challenge and without challenge, or APAS-untreated mice without challenge. Injection of the supernatant significantly increased leukocyte infiltration in the dermis of all mice (Fig 5; see this article’s Fig E4, A and B, in the Online Repository at www.jacionline.org). Pretreatment of the supernatant with neutralizing antibodies against MIP-1a, RANTES, and/or TARC significantly suppressed dermal infiltration, whereas injection of vehicle treated with antichemokine antibodies did not suppress the infiltration (see Fig E4, C). This implies that these chemokines are released from platelets in cutaneous tissue and contribute to leukocyte recruitment in the LPR.

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FIG 5. Effect of platelet-derived soluble factors on leukocyte recruitment into skin. Washed platelets were stimulated with TRAP. The activated platelet supernatant was injected into ear skin of APAS-treated mice (5 per group) just before challenge, after adding anti–MIP-1a, anti-RANTES, antiTARC, or isotype control antibody to the supernatant. The supernatant of platelets fixed without stimulation with TRAP served as a control. **P < .01.

Effect of antiplatelet drugs on ear swelling and leukocyte recruitment into skin To investigate whether systemic administration of antiplatelet drugs inhibits the IHR in skin, aspirin or clopidogrel was administered to mice before challenge. Neither aspirin nor clopidogrel suppressed the ear weight increase in the early phase (Fig 6, A), but the LPR was significantly decreased (see Fig 6, B). In addition, the levels of infiltrated cells in skin decreased in the LPR after administration of aspirin or clopidogrel (see Fig 6, C). P-selectin expression on platelets and the number of platelet-leukocytes complexes both decreased in the blood of mice treated with antiplatelet agents (see this article’s Fig E5, A, in the Online Repository at www.jacionline.org). Release of MIP1a, RANTES, and TARC was significantly inhibited in platelets exposed to antiplatelet agents (see Fig E5, B), but administration of these drugs did not affect proliferation and chemotaxis of mouse T cells or intercellular adhesion molecule 1 and vascular cell adhesion molecule 1 expression on murine skin endothelial cells and human umbilical vein endothelial cells (data not shown). These findings suggest that leukocyte recruitment into skin in the LPR can be suppressed by controlling platelet activity.

DISCUSSION The time course of LPR in this mouse model peaks 24 hours after antigen challenge, which is somewhat different from the peak of the human LPR, which occurs much sooner. Our experiments using mast cell–deficient W/Wv mice suggest that the biphasic cutaneous reactions are dependent on activation of mast cells, and it has been shown that antigen-specific IgE20 and FceRI24 are essential for development of the biphasic reactions as seen in this model, which indicates that findings in the W/Wv mouse are relevant to the human IHR. Depletion of about 90% of circulating platelets in the mice significantly reduced the LPR, suggesting that platelets are important for leukocyte recruitment into skin. The mechanism underlying stimulation of the LPR by platelets may involve mediation of leukocyte adhesion to the endothelium via platelet P-selectin in blood and secretion of chemokines at sites of inflammation in skin. Platelet P-selectin is an important adhesion molecule that mediates adhesion of activated platelets to monocytes, neutrophils, eosinophils, basophils, and T cells, resulting in formation of

FIG 6. Effect of antiplatelet drugs on ear swelling responses and leukocyte infiltration. The ear weight was examined 1 hour (A) and 24 hours (B) after challenge, and skin tissue was examined 24 hours after challenge (C; 5 mice per group). Data are shown as the percentage of the basal number of cells in the dermis after vehicle administration. **P < .01. NS, Not significant.

platelet-leukocyte complexes and permitting rolling of leukocytes along the endothelial cells.1,25 This phenomenon enables leukocytes to transmigrate into subendothelial tissue. Indeed, platelets have been reported to increase leukocyte rolling in murine skin, and P-selectin expression on platelets increases in the blood of psoriatic patients in parallel with disease severity.13 In this study, depletion or blockade of P-selectin expression markedly decreased leukocyte recruitment into skin, which suggests that platelet P-selectin is necessary for leukocyte recruitment into skin in the LPR. In platelets, chemokines synthesized by megakaryocytes are stored in a-granules, and chemokine receptors are expressed on membranes.1 The platelet-derived chemokines, which not only stimulate and attract leukocytes but also further activate other platelets, are crucial for the pathogenesis of an allergic dermatitis such as AD.15,18 In patients with AD, high levels of TARC are found in platelets15 and expressed in the lesional skin,26 and moreover, TARC induces extravasation of TH2 cells and skin-homing memory T cells expressing CCR4.27 RANTES and MIP-1a, which are secreted from stimulated platelets, are strong chemoattractants for lymphocytes, monocytes, and eosinophils,28,29 and are involved in the IHR, cell adhesion, chemotaxis, and fibrosis in allergic disorders.26,30,31 Our results demonstrate that injection of activated platelet supernatant into the skin tissue of platelet-depleted mice can restore cutaneous leukocyte recruitment in the LPR. Moreover, this phenomenon was almost completely blocked by antibody neutralization of MIP-1a, RANTES, and TARC in the supernatant. Therefore, our data suggest that platelets that reach the dermis by forming platelet-leukocyte complexes may be stimulated by collagen in the dermis and further activate other platelets by release of adenosine diphosphate, ATP, and other molecules. Chemokines released from activated platelets in the dermis may then induce leukocyte recruitment into the inflamed skin. In in vitro experiments, stimulation of human platelets via FceRI induces the release of proinflammatory mediators such as serotonin and RANTES.18 It has also been shown that platelets migrate extravascularly in response to antigen via a mechanism

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dependent on interactions among the antigen, antigen-specific IgE, and FceRI.3 These results imply that platelets may affect the response of the IHR. In our in vivo study, platelet depletion significantly reduced the LPR, but the early-phase reaction was not suppressed. Therefore, platelets in blood may not be stimulated directly by antigen exposure to skin and might not be associated with the early-phase reaction. However, it has been shown that platelet depletion suppresses the early phase of an IHR induced by injecting antigen intravenously in an animal model.32 Taken together, these results indicate that platelets play important roles in the early and late phases of the IHR. The exact mechanisms through which blood platelets are activated in the IHR in skin have not been elucidated. Plateletactivating factor (PAF), which is a potent phospholipid mediator released from leukocytes and endothelial cells stimulated by proinflammatory cytokines, contributes to the IHR at various sites, including the skin.23,33,34 Moreover, the PAF-induced IHR is dependent on platelet activation.5 It has also been reported that PAF induces P-selectin expression on platelets in blood35 and stimulates release of mediators from the platelets.36 We observed no difference between ovalbumin-sensitized and shamsensitized mice as donors in experiments with platelet restoration, suggesting that leukocyte recruitment into skin depends simply on the presence of platelets as circulating blood elements, regardless of sensitization of the platelets. Therefore, PAF, rather than antigen-specific IgE, may induce platelet activation in the IHR in skin. Once blood platelets are activated by PAF released during an inflammatory reaction, platelets may induce concomitant activation of other platelets and contribute to leukocyte recruitment into skin by binding to leukocytes via P-selectin in blood and through secretion of chemokines at the site of inflammation in the skin. Our results also suggest that control of platelet activity may lead to suppression of cutaneous allergic inflammation. Aspirin impairs platelet activation by inhibiting COX activity in platelets, which metabolizes arachidonic acid to a variety of prostanoids including thromboxane A2.37 Clopidogrel also has a significant antiplatelet effect by inhibiting adenosine diphosphate–mediated platelet activation,38 and has been reported to be activated by liver microsomal cytochrome P450 to give a metabolite that acts as an antagonist of the irreversible P2Y12 receptor.39 In the current study, systemic administration of aspirin or clopidogrel to mice significantly suppressed leukocyte recruitment into the cutaneous inflammatory lesion, and previous reports have shown that aspirin and clopidogrel inhibit P-selectin expression on platelets40,41 and block secretion of mediators from platelets.42 Thus, these drugs may inhibit leukocyte infiltration into skin by inhibiting formation of platelet-leukocyte complexes via P-selectin and reducing secretion of chemokines from platelets. Our data suggest that systemic administration of antiplatelet drugs may be beneficial for treatment of allergic dermatitis. However, suppression of the activity of circulating platelets can induce spontaneous or excessive bleeding in various organs. In this study, we observed a severe bleeding tendency in some mice after administration of large doses of antiplatelet drugs compared with the standard dose used in treatment of human thrombotic disorders. Therefore, topical application of antiplatelet drugs to skin may be useful for suppression of platelet activity at a cutaneous inflammatory site with lesser effects on other organs, and we are currently studying the effect of epicutaneous application of antiplatelet drugs on allergic skin inflammation.

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In conclusion, our results indicate that platelets are important for development of the LPR in skin. We showed that platelets induce leukocyte recruitment into skin by forming plateletleukocyte complexes via P-selectin in circulating blood and secreting several chemokines that attract leukocytes to skin. Systemic administration of antiplatelet drugs produced significant suppression of leukocyte infiltration, suggesting that treatment of allergic skin inflammation such as that in AD may be improved by control of platelet activity. Key messages d

Platelets are important in the late phase of the IHR in skin.

d

Platelets induce leukocyte recruitment into skin by forming platelet-leukocyte complexes via P-selectin in circulating blood and secreting several chemokines that attract leukocytes to skin.

d

Systemic administration of antiplatelet drugs to mice significantly suppresses leukocyte infiltration into skin.

REFERENCES 1. von Hundelshausen P, Weber C. Platelets as immune cells: bridging inflammation and cardiovascular disease. Circ Res 2007;100:27-40. 2. Pitchford SC, Yano H, Lever R, Riffo-Vasquez Y, Ciferri S, Rose MJ, et al. Platelets are essential for leukocyte recruitment in allergic inflammation. J Allergy Clin Immunol 2003;112:109-18. 3. Pitchford SC, Momi S, Baglioni S, Casali L, Giannini S, Rossi R. Allergen induces the migration of platelets to lung tissue in allergic asthma. Am J Respir Crit Care Med 2008;177:604-12. 4. Pitchford SC, Momi S, Giannini S, Casali L, Spina D, Page C, et al. Platelets P-selectin is required for pulmonary eosinophil and lymphocyte recruitment in a murine model of allergic inflammation. Blood 2005;105:2074-81. 5. Coyle AJ, Spina D, Page CP. PAF-induced bronchial hyperresponsiveness in the rabbit: contribution of platelets and airway smooth muscle. Br J Pharmacol 1990;101:31-8. 6. Schmitt-Sody M, Klose A, Gottschalk O, Metz P, Gebhard H, Zysk S, et al. Platelet-endothelial cell interactions in murine antigen-induced arthritis. Rheumatology 2005;44:885-9. 7. Danese S, de la Motte C, Sturm A, Vogel JD, West GA, String SA, et al. Platelets trigger a CD40-dependent inflammatory response in the microvasculature of imflammatory bowel disease patients. Gastroenterology 2003;124:1249-64. 8. Andoh A, Tsujikawa T, Hata K, Araki Y, Kitoh K, Sasaki M, et al. Elevated circulating platelet-derived microparticles in patients with active inflammatory bowel disease. Am J Gastroenterol 2005;100:2042-8. 9. Leung DY, Boguniewicz M, Howell MD, Nomura I, Hamid QA. New insights into atopic dermatitis. J Clin Invest 2004;113:651-7. 10. Galli SJ, Nakae S, Tsai M. Mast cells in the development of adaptive immune response. Nat Immunol 2005;6:135-42. 11. Gould HJ, Sutton BJ, Beavil AJ, Beavil RLMcCloskey N, Coker HA, et al. The biology of IGE and the basis of allergic disease. Annu Rev Immunol 2003;21:579-628. 12. Tamagawa-Mineoka R, Katoh N, Ueda E, Takenaka H, Kita M, Kishimoto S. The role of platelets in leukocyte recruitment in chronic contact hypersensitivity induced by repeated elicitation. Am J Pathol 2007;170:2019-29. 13. Ludwig RJ, Schultz JE, Boehncke WH, Podda M, Tandi C, Krombach F, et al. Activated, not resting, platelets increase leukocyte rolling in murine skin utilizing a distinct set of adhesion molecules. J Invest Dermatol 2004;122:830-6. 14. Mitsuhashi M, Tanaka A, Fujisawa C, Kawamoto K, Itakura A, Takaku M, et al. Necessity of thromboxane A2 for initiation of platelet-mediated contact sensitivity: dual activation of platelets and vascular endothelial cells. J Immunol 2001;166:617-23. 15. Fujisawa T, Fujisawa R, Kato Y, Nakayama T, Morita A, Katsumata H, et al. Presence of high contents of thymus and activation-regulated chemokine in platelets and elevated plasma levels of thymus and activation-regulated chemokine and macrophage-derived chemokine in patients with atopic dermatitis. J Allergy Clin Immunol 2002;110:139-46. 16. Tamagawa-Mineoka R, Katoh N, Ueda E, Masuda K, Kishimoto S. Elevated platelet activation in patients with atopic dermatitis and psoriasis: increased plasma levels of b-thromboglobulin and platelet factor 4. Allegol Int 2008;57:391-6.

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17. Katoh N. Platelets as versatile regulators of cutaneous inflammation. J Dermatol Sci 2009;53:89-95. 18. Hasegawa S, Pawankar R, Suzuki K, Nakahata T, Furukawa S, Okumura K, et al. Functional expression of the high affinity receptor for IgE (FceRI) in human platelets and its intracellular expression in human megakaryocytes. Blood 1999;93:2543-51. 19. Joseph M, Gounni AS, Kusnierz JP, Vorng H, Sarfati M, Kinet JP, et al. Expression and functions of the high-affinity IgE receptor on human platelets and megakaryocyte precursors. Eur J Immunol 1997;27:2212-8. 20. Sawada K, Nagai H, Basaki Y, Yamaha H, Ikizawa K Watanabe M, et al. The expression of murine cutaneous late phase reaction requires both IgE antibodies and CD4 T cells. Clin Exp Allergy 1996;27:225-31. 21. Stenberg PE, Levin J, Baker G, Mok Y, Corash L. Neuraminidase-induced thrombocytopenia in mice: effects on thrombopoiesis. J Cell Physiol 1991;147:7-16. 22. Kuter DJ, Rosenberg RD. The reciprocal relationship of thrombopoietin (c-Mpl ligand) to changes in the platelet mass during busulfan-induced thrombocytopaenia in the rabbit. Blood 1995;85:2720-30. 23. Strait RT, Morris SC, Yang M, Qu XW, Finkelman FD. Pathways of anaphylaxis in the mouse. J Allergy Clin Immunol 2002;109:658-68. 24. Miyahara S, Miyahara N, Takeda K, Joetham A, Gelfand EW. Physiologic assessment of allergic rhinitis in mice: role of the high-affinity IgE receptor (FceRI). J Allergy Clin Immunol 2005;116:1020-7. 25. de Brujine-Admirral LG, Modderman PW, von dem Borne AEGK, Sonnenberg A. P-selectin mediates Ca21-dependent adhesion of activated platelets to many different forms of leukocytes: detection by flow cytometry. Blood 1992;80:134-42. 26. Uchida T, Suto H, Ra C, Ogawa H, Kobata T, Okumura K. Preferential expression of Th2-type chemokine and its receptor in atopic dermatitis. Int Immunol 2002;14:1431-8. 27. Yoshie O, Imai T, Nomiyama H. Chemokines in immunity. Adv Immunol 2001;78: 57-110. 28. Kameyoshi Y, Dorschner A, Mallet AI, Christophers E, Schroder JM. Cytokine RANTES released by thrombin-stimulated platelets is a potent attractant for human eosinophils. J Exp Med 1992;176:587-92. 29. Klinger MH, Wilhelm D, Bubel S, Sticherling M, Schroder JM, Kuhnel W. Immunocytochemical localization of the chemokines RANTES and MIP-1a within human platelets and their release during storage. Int Arch Allergy Immunol 1995;107:541-6. 30. Tonnel AB, Gosset P, Molet S, Tillie-Leblond I, Jeannin P, Joseph M. Interactions between endothelial cells and effector cells in allergic inflammation. Ann N Y Acad Sci 1996;796:9-20.

TAMAGAWA-MINEOKA, KATOH, AND KISHIMOTO 587

31. Gruber BL, Kew RR, Jelaska A, Marchese MJ, Garlick J, Ren S, et al. Human mast cells activate fibroblasts: tryptase is a fibrogenic factor stimulating collagen messenger ribonucleic acid synthesis and fibroblast chemotaxis. J Immunol 1997; 158:2310-7. 32. Pinckard RN, Halonen M, Palmer JD, Butler C, Shaw JO, Henson PM. Intravascular aggregation and pulmonary sequestration of platelets during IgE-induced systemic anaphylaxis in the rabbit: abrogation of lethal anaphylactic shock by platelet depletion. J Immunol 1977;119:2185-93. 33. Vadas P, Gold M, Perelman B, Liss GM, Lack G, Blyth T, et al. Platelet-activating factor, PAF acetylhydrolase, and severe anaphylaxis. N Engl J Med 2008;358: 28-35. 34. Shalit M, Valone FH, Atkins PC, Ratnoff WD, Goetzl EJ, Zweiman B. Late appearance of phospholipid platelet-activating factor and leukotriene B4 in human skin after repeated antigen challenge. J Allergy Clin Immunol 1989;83:691-6. 35. Zeng S, Yi FX, Guo ZG. Platelet activating factor-induced P-selectin expression in platelets and its related signal transduction. Zhongguo Yao Li Xue Bao 1999;20:948-50. 36. Lapetina EG. Platelet-activating factor stimulates the phosphatidylinositol cycle. Appearance of phosphatidic acid is associated with the release of serotonin in horse platelets. J Biol Chem 1982;257:7314-7. 37. Roth GJ, Calverley DC. Aspirin, platelets, and thrombosis: theory and practice. Blood 1994;83:885-98. 38. Foster CJ, Prosser DM, Agans JM, Zhai Y, Smith MD, Lachowicz JE, et al. Molecular identification and characterization of the platelet ADP receptor targeted by thienopyridine antithrombotic drugs. J Clin Invest 2001;107:1591-8. 39. Savi P, Combalbert J, Gaich C, Rouchon MC, Maffrand JP, Berger Y, et al. The antiaggregating activity of clopidogrel is due to a metabolic activation by the hepatic cytochrome P450-1A. Thromb Haemost 1994;72:313-7. 40. McKenzie ME, Malinin AI, Bell CR, Dzhanashvili A, Horowitz ED, Oshrine BR, et al. Aspirin inhibits surface glycoprotein IIb/IIIa, P-selectin, CD63, and CD107a receptor expression on human platelets. Blood Coagul Fibrinolysis 2003;14: 249-53. 41. Angiolillo DJ, Ferna´ndez-Ortiz A, Bernardo E, Ramı´rez C, Sabate´ M, Ban˜uelos C, et al. High clopidogrel loading dose during coronary stenting: effects on drug response and interindividual variability. Eur Heart J 2004;25:1903-10. 42. Nannizzi-Alaimo L, Alves VL, Phillips DR. Inhibitory effects of glycoprotein IIb/ IIIa antagonists and aspirin on the release of soluble CD40 ligand during platelet stimulation. Circulation 2003;107:1123-8.

587.e1 TAMAGAWA-MINEOKA, KATOH, AND KISHIMOTO

METHODS Mice Male BALB/c, WBB6F1-1/1, and WBB6F1-W/Wv mice were obtained from Shimizu Laboratory Supplies (Kyoto, Japan). Male P-selectin knockout mice (C57BL/6J background)E1 and control wild-type C57/BL6J mice were obtained from The Jackson Laboratory (Bar Harbor, Me). All mice were 6 to 8 weeks old at the time of use.

Isolation of murine platelets Blood was mixed with acid citrate dextrose at a 9:1 ratio and centrifuged at 150g for 10 minutes. The platelet-rich plasma obtained was washed using the method of Mustard et al.E2 Briefly, washed platelets were suspended in minimum essential medium containing 300 ng/mL prostaglandin I2 and 10% FCS. The cellular content of the suspension was >99% platelets, as demonstrated by flow cytometry (data not shown).

Preparation of APAS Antiplatelet antiserum was prepared as previously described.E3 Briefly, washed platelets (3 3 109) were isolated from normal BALB/c mice, homogenized in complete Freund adjuvant, and injected subcutaneously in rabbits. A second immunization was performed 10 days later by injecting platelets in incomplete Freund adjuvant intramuscularly. Thirty days after the second immunization, the rabbits were bled, and antiserum was inactivated at 568C for 60 minutes. Serum obtained from nonimmunized rabbits served as CS.

Blood sampling for platelet, leukocyte, and serum IgE profiles Platelets were counted using the method of Brecher and Cronkite.E4 Briefly, blood was diluted 1:20 with 1% ammonium oxalate to hemolyze erythrocytes. For leukocyte counts, blood was mixed with a hemolysis solution at a 9:1 ratio. Levels of serum total IgE and ovalbumin-specific IgE were analyzed using ELISA kits purchased from Bethyl Laboratories (Montgomery, Tex) and Dainippon Sumitomo Pharma (Osaka, Japan), respectively.

Histologic examination The ears of mice were excised, fixed with 10% formalin, and then embedded in paraffin. Sections (5 mm) were cut using a microtome and stained with hematoxylin and eosin for general histological evaluation or toluidine blue for mast cell staining. For immunohistochemistry, frozen sections (5 mm) were cut in a cryostat, placed on siliconized glass slides, and air-dried. They were then incubated with rat anti–intercellular adhesion molecule 1 (clone KAT-1; R&D Systems, Abingdon, United Kingdom [UK]) or anti–vascular cell adhesion molecule 1 (clone M/K-2; Serotec, Oxford, UK) antibody overnight at 48C. Sections were incubated sequentially for 30 minutes in room air with biotinylated rabbit antirat IgG (DAKO, Santa Barbara, Calif) and then horseradish peroxidase–conjugated avidin-biotin (DAKO). Sections were developed with 3,39-diaminobenzidine tetrahydrochloride and hydrogen peroxidase and then counterstained with hematoxylin.

Myeloperoxidase and eosinophil peroxidase activities Ear tissue was homogenized and myeloperoxidase activity was measured using a myeloperoxidase assay kit (CytoStore, Calgary, Alberta, Canada) according to the manufacturer’s instructions, with absorbance measured at 450 nm. Eosinophil peroxidase activity was measured based on absorbance at 490 nm after incubation of a homogenate of ear tissue (50 mL) with 100 mL 50 mmol/LTRIS-buffered saline containing 0.1% Triton X-100, 1 mmol/L o-phenylenediamine, and 500 mmol/L hydrogen peroxide, as previously reported.E5

Flow cytometry Blood in citrate was incubated with fluorescein isothiocyanate–labeled rat anti-CD41 (clone MWReg30; Serotec) and rabbit polyclonal anti-CD62P (Chemicon Europe, Hampshire, UK) antibodies, followed by secondary

J ALLERGY CLIN IMMUNOL MARCH 2009

phycoerythrin-labeled goat antirabbit IgG. In some experiments, blood was incubated with fluorescein isothiocyanate–labeled anti-CD41. Live gating was performed for leukocyte-sized events, the forward and side scattering characteristics of which exclude the presence of single platelets alone, as described previously.E6 Events in this region that were positive for CD41 were considered to represent platelet-leukocyte aggregates. In separate experiments, human umbilical vein endothelial cells (Health Science Research Resources Bank, Osaka, Japan) were cultured in MCDB 107 medium (Cosmo Bio, Tokyo, Japan) supplemented with heparin (100 mg/mL), 10% FBS, and endothelial cell growth supplement (50 mg/mL). Confluent human umbilical vein endothelial cells were then incubated with 10% APAS, 2 mmol/L busulfan, or 0.05 U/mL neuraminidase, or with 1 mmol/L aspirin or 1 mmol/L clopidogrel for 30 minutes at 378C, and analyzed by using phycoerythrin-labeled mouse anti–intercellular adhesion molecule 1 (eBioscience, San Diego, Calif) and fluorescein isothiocyanate– labeled mouse anti–vascular cell adhesion molecule 1 (Serotec) antibodies.

Measurement of chemokines in platelet supernatant Washed platelets (1 3 109/mL) were stimulated with 20 mmol/L TRAP for 15 minutes at 378C. In some experiments, washed platelets were treated with 1 mmol/L aspirin or vehicle for 30 minutes at 378C before stimulation with TRAP. In separate experiments, mice were dosed orally with clopidogrel (30 mg/kg/day) or vehicle for 2 days, and blood was collected 1 hour after the final dose. Washed platelets were isolated, suspended, and then stimulated with TRAP. The levels of MIP-1a, RANTES, and TARC in the platelet supernatant and lysates were measured by using ELISA kits (R&D Systems, Minneapolis, Minn). A supernatant of platelets fixed with 2% paraformaldehyde without stimulation with TRAP served as a control. Data are shown as the percentage of the maximum number of chemokines in platelet lysates.

Cell proliferation assay Purified T cells were obtained from blood of mice by using magnetic microbeads recognizing CD5 (Miltenyi Biotec, Bergisch Gladbach, Germany) and then stimulated with anti-CD3/anti-CD28–coated beads (Invitrogen, Carlsbad, Calif) in medium supplemented with recombinant IL-2 according to the manufacturer’s instructions. After stimulation, 4-[3-(4-iodophenyl)2-(4-nitrophenyl)-2H-5-tetrazolio]-1,3-benzene disulfonate (Premix WST-1; Takara Bio, Shiga, Japan) was added, and the rate of reduction of the tetrazolium salt was estimated according to the manufacturer’s protocol.

Chemotaxis assay T cells purified as described were added to the upper chamber of chemotaxicells (8-mm pore size; Kurabo, Osaka, Japan), and the supernatant of platelets stimulated with TRAP was added to the lower chamber. After incubation for 2 hours at 378C, the membranes of the chemotaxicells were removed, fixed, and Giemsa-stained. The number of cells that transmigrated toward the lower chambers was counted under a microscope. REFERENCES E1. Bullard DC, Qin L, Lorenzo I, Quinlin WM, Doyle NA, Bosse R, et al. P-selectin/ ICAM-1 double mutant mice: acute emigration of neutrophils into the peritoneum is completely absent but is normal into pulmonary alveoli. J Clin Invest 1995;95: 1782-8. E2. Mustard JF, Perry DW, Ardlie NG, Packam MA. Preparation of suspensions of washed platelets from humans. Br J Haematol 1972;22:193-204. E3. Merhi Y, Provost P, Guidoin R, Latour JG. Importance of platelets in neutrophil adhesion and vasoconstriction after deep carotid arterial injury by angioplasty in pigs. Arterioscler Thromb Vasc Biol 1997;17:1185-91. E4. Brecher G, Cronkite EP. Morphology and enumeration of human blood platelets. J Appl Physiol 1950;3:365-75. E5. Kawase Y, Hoshino T, Yokota K, Kuzuhara A, Kirii Y, Nishiwaki E, et al. Exacerbated and prolonged allergic and non-allergic inflammatory cutaneous reaction in mice with targeted interleukin-18 expression in the skin. J Invest Dermatol 2003;121:502-9. E6. Shattil SJ, Cunningham M, Hoxie JA. Detection of activated platelets in whole blood using activation-dependent monoclonal antibodies and flow cytometry. Blood 1987;70:307-15.

TAMAGAWA-MINEOKA, KATOH, AND KISHIMOTO 587.e2

B Ear weight increase at 24 h (mg)

Ear weight increase at 1 h (mg)

A 8 6 4 2 0

W/Wv

+/+

8 6 4 2 0 +/+

W/Wv

Ear swelling response (μm)

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OVA+Vehicle OVA+Neuraminidase 150

Sham+Vehicle Sham+Neuraminidase

100 50 0 01

6

12

18

24

30

36

42

48

Time after challenge (hours)

4 2 0

E

8 6 4 2 0

APAS

CS

APAS

0.6 0.5

0.3

EPO activity (OD at 490 nm)

6

MPO activity (OD at 450 nm)

Ear weight increase at 24 h (mg)

8

CS

Ear weight increase at 1 h (mg)

D

NS

0.4 0.3 0.2 0.1 0 CS

APAS

0.2 0.1 0

CS

APAS

NS

8 6 4 2 0 CS

APAS

Ear weight increase at 24 h (mg)

Ear weight increase at 1 h (mg)

C

8 6 4 2 0 CS

APAS

FIG E1. Changes in ear swelling responses after ovalbumin (OVA) challenge. The ear weight of W/Wv mice (A) and that of mice injected with APAS (C) or challenged (E) repeatedly and the ear thickness of mice injected with neuraminidase (B) were examined. The myeloperoxidase (MPO) and eosinophil peroxidase activities of the skin tissue were measured (D). Six mice per group. **P < .01 vs vehicle administration in OVA-challenged mice.

587.e3 TAMAGAWA-MINEOKA, KATOH, AND KISHIMOTO

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FIG E2. Effect of platelet addition on ear swelling responses and chemokine release from in vitro ovalbumin-challenged platelets. The ear weight of mice without platelet depletion (6 per group) was examined after injection of platelet suspension (A). The levels of MIP-1a, RANTES, and TARC were measured in triplicate in the supernatant of platelets treated with serum of sensitized mice and ovalbumin (B). *P < .05. NS, Not significant.

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TAMAGAWA-MINEOKA, KATOH, AND KISHIMOTO 587.e4

FIG E3. Effect of platelet restoration on formation of platelet-leukocyte complexes. Mice (5 per group) were injected with platelet suspension obtained from ovalbumin (OVA)-sensitized or sham-sensitized mice, or P-selectin–deficient mice. The number of platelet-leukocyte complexes in blood was examined. Data are shown as percentages of basal values in sham-sensitized mice (A). Chemokine levels were measured in triplicate in the supernatant of P-selectin–deficient platelets (B). *P < .05; **P < .01. WT, Wild-type. NS, Not significant.

587.e5 TAMAGAWA-MINEOKA, KATOH, AND KISHIMOTO

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FIG E4. Effect of platelet-derived soluble factors on leukocyte recruitment into skin. The activated platelet supernatant was injected into ear skin of mice with (A) and without (B) platelet depletion, without challenge. Vehicle treated with antichemokine antibodies was also injected into ear skin (C). Data are shown as percentages of basal numbers in mice injected with the supernatant of fixed platelets. Five mice per group. **P < .01. NS, Not significant.

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TAMAGAWA-MINEOKA, KATOH, AND KISHIMOTO 587.e6

FIG E5. Effect of antiplatelet drugs on formation of platelet-leukocyte complexes and chemokine release from platelets. P-selectin expression on platelets and the number of platelet-leukocyte complexes were examined in the blood of mice (6 per group) treated with antiplatelet agents (A). Chemokine levels were measured in triplicate in the supernatant of platelets treated with antiplatelet agents (B). **P < .01. OVA, Ovalbumin.

587.e7 TAMAGAWA-MINEOKA, KATOH, AND KISHIMOTO

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TABLE E1. Platelet and leukocyte counts before and after APAS, busulfan, or neuraminidase administration Platelet count (/mL)

APAS

Busulfan

Neuraminidase

1 h before sensitization 1 h before challenge 1 h after challenge 48 h after challenge 1 h before sensitization 1 h before challenge 1 h after challenge 48 h after challenge 1 h before sensitization 1 h before challenge 1 h after challenge 48 h after challenge

Data are expressed as the means 6 SDs of 6 mice per group. **P < .01 vs platelet count at 1 hour before sensitization.

9.2 1.0 1.0 1.2 9.6 1.1 1.1 1.2 9.8 1.1 1.0 1.1

6 6 6 6 6 6 6 6 6 6 6 6

0.4 0.2 0.4 0.3 0.3 0.1 0.3 0.2 0.4 0.2 0.3 0.3

3 3 3 3 3 3 3 3 3 3 3 3

8

10 108** 108** 108** 108 108** 108** 108** 108 108** 108** 108**

Leukocyte count (/mL)

6.4 6.9 6.7 7.2 7.5 5.9 6.2 6.1 7.8 6.7 6.9 6.5

6 6 6 6 6 6 6 6 6 6 6 6

0.9 0.3 0.5 0.4 0.7 0.4 0.5 0.3 0.6 0.5 0.3 0.6

3 3 3 3 3 3 3 3 3 3 3 3

106 106 106 106 106 106 106 106 106 106 106 106

TAMAGAWA-MINEOKA, KATOH, AND KISHIMOTO 587.e8

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TABLE E2. Serum levels of total IgE antibody and antiovalbumin IgE antibody after sensitization

Nonsensitization CS APAS Vehicle Busulfan Vehicle Neuraminidase

Total IgE (ng/mL)

Ovalbumin-specific IgE (ng/mL)

6 6 6 6 6 6 6

Not detected 213 6 20 215 6 17 220 6 16 216 6 22 207 6 12 208 6 15

10.3 237 243 241 240 230 233

1.0 23 22 15 18 11 14

Data are expressed as the means 6 SDs of 6 mice per group.

587.e9 TAMAGAWA-MINEOKA, KATOH, AND KISHIMOTO

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TABLE E3. Mast cell numbers and percentage of degranulated mast cells

Mast cell numbers (/mm2) 1h 24 h Percentage of degranulated mast cells (%) 1h 24 h

APAS

CS

70 6 9 72 6 6

73 6 11 74 6 7

46 6 6 10 6 2

44 6 5 11 6 4

Data are expressed as the means 6 SDs of 6 mice per group.