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Originally published In Press as doi:10.1074/jbc.M409592200 on November 19, 2004

J. Biol. Chem., Vol. 280, Issue 4, 2888-2895, January 28, 2005
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Activation of CD38 by Interleukin-8 Signaling Regulates Intracellular Ca2+ Level and Motility of Lymphokine-activated Killer Cells*

So-Young Rah{ddagger}, Kwang-Hyun Park{ddagger}, Myung-Kwan Han{ddagger}§, Mie-Jae Im{ddagger}, and Uh-Hyun Kim{ddagger}||

From the {ddagger}Department of Biochemistry and the Institute of Cardiovascular Research, Chonbuk National University Medical School, Jeonju, 561-182, Republic of Korea

Received for publication, August 20, 2004 , and in revised form, November 16, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
CD38 is an ADP-ribosyl cyclase, producing a potent Ca2+ mobilizer cyclic ADP-ribose (cADPR). In this study, we have investigated a role of CD38 and its regulation through interleukin-8 (IL8) signaling in lymphokine-activated killer (LAK) cells. Incubation of LAK cells with IL8 resulted in an increase of cellular cADPR level and a rapid rise of intracellular Ca2+ concentration ([Ca2+]i), which was sustained for a long period of time (>10 min). Preincubation of an antagonistic cADPR analog, 8-Br-cADPR (8-bromo-cyclic adenosine diphosphate ribose), abolished the sustained Ca2+ signal only but not the initial Ca2+ rise. An inositol 1,4,5-trisphosphate (IP3) receptor antagonist blocked both Ca2+ signals. Interestingly, the sustained Ca2+ rise was not observed in the absence of extracellular Ca2+. Functional CD38-null (CD38-) LAK cells showed the initial rapid increase of [Ca2+]i but not the sustained Ca2+ rise in response to IL8 treatment. An increase of cellular cADPR level by cGMP analog, 8-pCPT-cGMP (8-(4-chlorophenylthio)-guanosine-3',5'-cyclic monophosphate), but not cAMP analog or phorbol 12-myristate 13-acetate was observed. IL8 treatment resulted in the increase of cGMP level that was inhibited by the IP3 receptor blocker but not a protein kinase C inhibitor. cGMP-mediated Ca2+ rise was blocked by 8-Br-cADPR. In addition, IL8-mediated LAK cell migration was inhibited by 8-Br-cADPR and a protein kinase G inhibitor. Consistent with these observations, IL8-induced migration of CD38- LAK cells was not observed. However, direct application of cADPR or 8-pCPT-cGMP stimulated migration of CD38- cells. These results demonstrate that CD38 is stimulated by sequential activation of IL8 receptor, IP3-mediated Ca2+ rise, and cGMP/protein kinase G and that CD38 plays an essential role in IL8-induced migration of LAK cells.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
A type II transmembrane protein CD38, originally known as an activation antigen, displays ADP-ribosyl cyclase (ADPR-cyclase)1 and cyclic ADP-ribose hydrolase (cADPR-hydrolase) activities (1, 2). These two enzyme activities are involved in the conversion of {beta}-nicotinamide adenine dinucleotide ({beta}-NAD+) first to cyclic ADP-ribose (cADPR) and then to ADP-ribose (ADPR) (3–5). CD38 is also ADP-ribosylated by ecto-ADP-ribosyltransferase in the presence of exogenous {beta}-NAD+ (6). This modification results in inactivation of the enzyme activity. The metabolite cADPR is known to induce Ca2+ release from intracellular stores by acting on ryanodine receptor and/or Ca2+ influx through plasma membrane Ca2+ channels in a variety of cells (7–11). Several studies have indicated that cADPR synthesis by CD38 is stimulated through cell surface heterotrimeric G-protein-coupled receptor signaling. The receptors include {beta}-adrenergic receptor in cardiac myocytes (12) and artery smooth muscle cells (13), angiotensin II receptor in cardiac myocytes (14), muscarinic receptor in neuroblastoma NG-108 (15), and pancreatic acinar cells (16). The activation of ADPR-cyclase by cGMP in Aplysia califonica has been reported (17), and cAMP-dependent activation of the enzyme is also observed in artery smooth muscle cells (13). However, the molecular basis of the activation of CD38 and/or ADPR-cyclases has not been clearly defined.

A previous report has indicated that a peptide cytokine interleukin (IL) 8 signaling may utilize cADPR to mobilize [Ca2+]i in IL2-activated natural killer cells (18). IL8, which belongs to the CXC superfamily of chemokines, plays an important role in the motility of various cells such as neutrophils and T cells (19, 20) and also induces angiogenesis and other effects associated with proinflammatory responses (21–23). The IL8 receptor (IL8R) is made of seven transmembrane proteins and couples with Gi and stimulates the production of inositol 1,4,5-trisphosphate (IP3) through the activation of phospholipase C (PLC)-{beta}2 (24, 25). There is a report that the IL8Rs present in natural killer cells and lymphokine-activated killer (LAK) cells may induce cADPR synthesis through the Gs-involved signaling pathway (26). CD38 expression in natural killer and LAK cells is also observed (27, 28). However, the role of CD38 including cADPR in LAK cell functions and the activation pathways of CD38 remain elusive.

In this study, we have investigated IL8-mediated regulation of CD38 by determining intracellular Ca2+ changes and motility of LAK cells. The results indicate that CD38 is activated via cGMP/protein kinase G (PKG) that is activated by IL8 and that CD38 plays a critical role in IL8-mediated Ca2+ signal and migration of LAK cells.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Reagents and Antibodies—Antibodies were obtained as follows: anti-human CD38 monoclonal antibody from BD Biosciences; anti-IL8R monoclonal antibody from Santa Cruz Biotechnology (Santa Cruz, CA); anti-PLC-{beta}2 monoclonal antibody from P. G. Suh at POSTECH (Pohang, Korea); anti-IP3 receptor (IP3R) generously provided by S. H. Kim (Inha University, Incheon, Korea); anti-ryanodine receptor (RyR)common mouse monoclonal antibody from Calbiochem. Ficoll-Hypaque and Percoll were obtained from Amersham Biosciences. Horseradish peroxidase-conjugated anti-mouse or anti-rabbit IgG was purchased from Advanced Biochemicals Inc. (Jeonju, Korea). Human recombinant IL2 was purchased from Chiron BV (Amsterdam, Netherlands). Human recombinant IL8, human AB serum, and all other reagents were obtained from Sigma. RPM1 1640 was from Invitrogen. 125I-cGMP radioimmunoassay kit was from PerkinElmer Life Sciences.

Preparation of LAK Cells—LAK cells were prepared as described previously (29, 30). Briefly, blood obtained from healthy volunteers was layered over Ficoll-Hypaque and centrifuged at 700 x g for 30 min to remove red blood cells. The red blood cells removed cell preparations were incubated on a nylon-wool column at 37 °C for 1 h in a 5% CO2 incubator to remove B lymphocytes and macrophages. Nylon-wool nonadherent cells were collected and further separated by a Percoll density gradient centrifugation. Four layers of Percoll were used: 37, 44, 52, and 60%. After centrifugation at 700 x g for 20 min, cells of the 52% Percoll layer were collected, washed with serum-free RPMI 1640, and incubated at 2 x 106 cells/ml density in a culture media containing 3000 IU/ml IL2 in a 5% CO2 incubator at 37 °C. The culture medium used is RPMI 1640 supplemented with 10% human AB serum, 0.25 µg/ml amphotericin B, 50 µg/ml gentamycin, 10 units/ml penicillin G, 100 µg/ml streptomycin, 1 mM L-glutamine, 1% nonessential amino acids, and 50 µM 2-mercaptoehanol. After incubation for 24 h, the floating cells were removed, and the adherent cells were cultured in the same culture media containing 1500 IU/ml IL2. LAK cells induced by IL2 for 10 days were used throughout the study, otherwise indicated in the figure legends. Induction of LAK cells was ensured by determining ADPR-cyclase activity as detailed below.

Western Blotting—LAK cells were washed with phosphate-buffered saline and then lysed with an ice-cold lysis buffer (20 mM HEPES, pH 7.2, 1% Triton X-100, 10% glycerol, 100 mM NaCl, 1 mM EDTA, 1 mM phenylmethylsulfonyl fluoride, 50 mM NaF, 1 mM Na3VO4, 10 µg/ml leupeptin, 10 µg/ml pepstatin, and 10 µg/ml aprotinin). After centrifugation at 20,000 x g for 10 min, supernatants were taken. The extracts (20 µg/lane) were subjected to SDS-PAGE in a 5% gel to identify RyR or IP3R, and in a 10% gel to identify PLC-{beta}2, IL8R, or CD38. After transfer to nitrocellulose membranes, blots were incubated in a blocking buffer (10 mM Tris-HCl, pH 7.6, 150 mM NaCl, 0.05% Tween 20) containing 5% nonfat dry milk for 2 h at room temperature and then with primary antibodies (dilution factors were: CD38, 1:500; RyR, 1:100; IP3R, 1:2000; PLC-{beta}2, 1:2000; IL8R, 1:200) in the blocking buffer overnight at 4 °C. The blots were rinsed four times with the blocking buffer and incubated with horseradish peroxidase-conjugated anti-mouse IgG (1: 5000 dilution) or anti-rabbit IgG (1:5000 dilution) in the blocking buffer for 1 h at room temperature. The immunoreactive proteins with respective secondary antibodies were determined using an enhanced chemiluminescence kit (Amersham Biosciences) and exposed to BIOMAX films (Eastman Kodak Co.). Protein concentrations were determined using a Bio-Rad protein assay kit, and known concentrations of bovine serum albumin (BSA) were used as the standard.

Synthesis of cADPR—cADPR was synthesized using ADPR-cyclase purified from A. califonica ovotestes as described previously (31). The nucleotide was purified by high performance liquid chromatography (HPLC) using AG MP-1 resin (Bio-Rad). The nucleotide was eluted with a nonlinear gradient of 150 mM trifluoroacetic acid and water. Purified cADPR was dried using a SpeedVac concentrator. The purity of cADPR used in the study was ~97% as determined by HPLC.

Determination of ADPR-cyclase Activity—ADPR-cyclase activity was determined by fluorometrically using nicotinamide guanine dinucleotide (NGD+) as a substrate (32). Cells were incubated with 200 µM NGD+ in 0.1 M sodium phosphate buffer (pH 7.2) at 37 °C for 10 min. Fluorescence of cGDPR produced was determined at excitation/emission wavelengths of 297/410 nm (Hitachi F-2000).

{beta}-NAD+-dependent Inactivation of CD38—LAK cells were incubated with 100 µM {beta}-NAD+ for 6 h according to the method described previously (6), and after incubation, the cells were washed with phosphate-buffered saline and used for the study.

Measurement of cADPRi—The level of [cADPR]i was measured using a cyclic enzymatic assay as described previously (33). Briefly, LAK cells were treated with 0.5 ml of 0.6 M perchloric acid under sonication. Precipitates were removed by centrifugation at 20,000 x g for 10 min. Perchloric acid was removed by mixing the aqueous sample with a solution containing 3 volumes of 1,1,2-trichlorotrifluoroethane to 1 volume of tri-n-octlyamine. After centrifugation for 10 min at 1500 x g, the aqueous layer was collected and neutralized with 20 mM sodium phosphate (pH 8). To remove all contaminating nucleotides, the samples were incubated with the following hydrolytic enzymes overnight at 37 °C: 0.44 unit/ml nucleotide pyrophosphatase, 12.5 units/ml alkaline phosphatase, 0.0625 unit/ml NADase, and 2.5 mM MgCl2 in 20 mM sodium phosphate buffer (pH 8.0). Enzymes were removed by filtration using Centricon-3 filters. To convert cADPR to NAD+, the samples (0.1 ml/tube) were incubated with 50 µl of a cycling reagent containing 0.3 µg/ml Aplysia ADPR-cyclase, 30 mM nicotinamide, and 100 mM sodium phosphate (pH 8) at room temperature for 30 min. The samples were further incubated with the cycling reagent (0.1 ml) containing 2% ethanol, 100 µg/ml alcohol dehydrogenase, 20 µM resazurin, 10 µg/ml diaphorase, 10 µM riboflavin 5'-phosphate, 10 mM nicotinamide, 0.1 mg/ml BSA, and 100 mM sodium phosphate (pH 8.0) for 2 h at room temperature. An increase in the resorufin fluorescence was measured at 544 nm excitation and 590 nm emission using a fluorescence plate reader (Molecular Devices Corp., Spectra-Max GEMINI). Various known concentrations of cADPR were also included in the cycling reaction to generate a standard curve.

Measurement of Intracellular cGMP Level—Levels of cGMP were determined by a 125I-cGMP radioimmunoassay kit according to the manufacturer's protocol. LAK cells were preincubated with 0.5 mM isobutylmethylxanthine and phosphodiesterase inhibitor and then challenged with various reagents as detailed in the figure legends. After incubation for 30 min, cells were treated with equal volume of 12% trichloroacetic acid. To determine cGMP extraction efficacy, [3H]cGMP (1500 cpm) was added. After centrifugation at 20,000 x g at 4 °C for 10 min, supernatants were collected and extracted four times with a 5-ml portion of water-saturated diethyl ether. The water part was collected, dried using SpeedVac, and dissolved in 200 µl of 50 mM sodium acetate buffer (pH 6.2). Prior to performing radioimmunoassay, the sample (100 µl) was acetylated using acetic anhydride in the presence of triethylamine. A standard curve of acetylated cGMP was also prepared as described in the manufacturer's protocol.

Measurement of [Ca2+]i—LAK cells were washed with Hanks' balanced salt solution (2 mM CaCl2, 145 mM NaCl, 5 mM KCl, 1 mM MgCl2, 5 mM D-glucose, 20 mM HEPES, pH 7.3) containing 1% BSA and incubated in the same solution containing 1% BSA for 6 h. Starved LAK cells were incubated with 5 µM Fluo3 AM (Molecular Probe, Eugene, OR) in Hanks' balanced salt solution containing 1% BSA at 37 °C for 40 min. The cells were washed three times with Hanks' balanced salt solution. Changes in [Ca2+]i in LAK cells were determined at 488 nm excitation/530 nm emission by air-cooled argon laser system (34). The emitted fluorescence at 530 nm was collected using a photomultiplier. One image every 6 s for 10 min was scanned using confocal microscope (Nikon, Japan). For the calculation of [Ca2+]i, the method of Tsien et al. (35) was used with the following equation: [Ca2+]i = Kd(F - Fmin)/(Fmax - F), where Kd is 450 nM for Fluo-3 and F is the observed fluorescence levels. Each tracing was calibrated for the maximal intensity (Fmax) by the addition of ionomycin (8 µM) and for the minimal intensity (Fmin) by the addition of EGTA 50 mM at the end of each measurement.

Determination of Cell Migration—Cell migration was determined as described previously (28). In brief, cells were washed with RPMI 1640, scrapped using a policeman, and washed with RPM1 1640. Transwells (Costar Corning, Corning, NY) with 8-µm pore size polycarbonate filters 6.5 mm in diameter were used. Lower chambers contained 500 µl of RPMI 1640 containing 1% BSA. LAK cells (4 x 105) in 100 µl RPM1 1640 containing various agents (specified in the figure legends) were placed in the upper chamber. The chambers were incubated in a 5% CO2 incubator at 37 °C for 2 h. The filters were removed, fixed with ice-cold 100% methanol, and stained with 15% Wright-Giemsa stain for 7 min. The cells were counted under a phase-contrast microscope.

Statistical Analysis—Data represent means ± S.E. of the mean (S.E.) of at least three separate experiments. Statistical analysis was performed using Student's t test. A value of p < 0.05 was considered significant.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Induction of CD38 Expression in LAK Cells—Firstly, we assessed expression levels of CD38 along with the other related signaling molecules during induction of LAK cells by IL2 treatment. Expression of CD38 was gradually increased in a time-dependent manner and highly induced at 7–10 days (Fig. 1A). Expression of RyR, a putative receptor for cADPR, was also highly induced at 7–10 days. On the other hand, expression of IL8R, IP3R, and PLC-{beta}2 was observed in the freshly isolated T cells and was not influenced by IL2 treatment (data not shown). Expression of CD38 was ascertained by measuring ADPR-cyclase activity using NGD+, which is converted to cGDPR only by the enzyme (Fig. 1B). Consistent with the above observations, the production of cGDPR in cell lysates was increased in a time-dependent manner. Next, we examined whether IL8 stimulates CD38 in LAK cells induced by treatment with IL2 for 10 days. As shown in Fig. 1C, [cADPR]i in LAK cells was increased significantly by the treatment with IL8. These results show that expression of CD38 and RyR along with transformation of T cells to LAK cells are induced by IL2 treatment and that IL8 signaling may activate CD38.



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  		FIG. 1.
Expression of CD38 and RyR during induction of LAK cells. Isolated T lymphocytes were treated with IL2 for 10 days. During induction of LAK cells, expression of CD38 and RyR was determined. A, CD38 and RyR are expressed during induction of LAK cell by IL2. A typical Western blot of CD38 and RyR expression during induction of LAK cells is shown. Cell lysate (20 µg) was subjected to immunoblotting using CD38 and RyR antibodies. B, increase of ADPR activity during LAK cell induction. ADPR cyclase activity in cell lysates (10 µg) prepared from IL2 treated cells was determined using 200 µM NGD+ as a substrate. NA, no activity found. C, [cADPR]i is increased in response to IL8 treatment. LAK cells were incubated with IL8 (10 pM) for 1.5 min. Formation of cADPR was determined as described under "Experimental Procedures." The data ± S.E. from three independent experiments are shown. **, p < 0.005.

 
IL8-mediated Elevation of [Ca2+]i Involves Activation of CD38—On the basis of the above observation that IL8 treatment increases [cADPR]i, the molecular basis of CD38 activation by IL8 in LAK cells was examined by determining IL8-mediated changes in [Ca2+]i under various conditions. The addition of IL8 to the cells resulted in a rapid increase in [Ca2+]i, and the increased [Ca2+]i levels were sustained for more than 10 min (Fig. 2A). The pretreatment of an antagonistic cADPR analog, 8-Br-cADPR, abolished the IL8-mediated sustained Ca2+ signal but not the initial rapid increase (Fig. 2B). Interestingly, xestospongin C, an IP3 receptor antagonist, completely abolished both Ca2+ signals (Fig. 2C). In contrast, calphostin C, a protein kinase C inhibitor, had no effect on the IL8-mediated increase of [Ca2+]i (Fig. 2D). To examine whether the sustained Ca2+ increase is due to Ca2+ release from the intracellular store or Ca2+ influx, the IL8-mediated increase of [Ca2+]i was determined in the presence of EGTA. The sustained Ca2+ rise was not observed while the initial Ca2+ increase was present (Fig. 2E). The effects of various agents on the initial and sustained Ca2+ increases were summarized in Fig. 2, F and G, respectively. These results indicate that cADPR is responsible for IL8-mediated sustained Ca2+ influx that is dependent on the IP3-mediated initial Ca2+ rise in the LAK cells.



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  		FIG. 2.
cADPR is involved in IL8-mediated increase in [Ca2+]i. A, the treatment of LAK cells with IL8 (10 pM) induces long-lasting intracellular Ca2+ signal. B, antagonistic cADPR analog abolishes the IL8-mediated sustained Ca2+ increase, whereas the initial Ca2+ increase remains. LAK cells were preincubated with 8-Br-cADPR (100 µM) for 30 min, and IL8 (10 pM) was then added to the cells. C, IP3R blocker completely eliminates IL8-induced Ca2+ signals. LAK cells were preincubated with xestospongin C (2 µM) for 30 min. D, protein kinase C inhibitor does not block IL8-mediated increase of [Ca2+]i. Prior to treatment with IL8, calphostin C (100 µM) was preincubated at 37 °C for 30 min. E, IL8-mediated sustained Ca2+ rise is due to Ca2+ influx. IL8-mediated Ca2+ signals were determined in the presence of 3 mM EGTA (Ca2+-free). Three representative Ca2+ traces are shown. Arrows indicate the time point of the addition of IL8. F, a direct comparison of mean [Ca2+]i during initial increases of [Ca2+]i. The data shown are analyzed at 54 s. #, buffer versus IL8 or IL8 plus agents indicated, p < 0.001; ***, IL8 versus xestospongin C plus IL8, p < 0.001. G, a direct comparison of mean [Ca2+]i during sustained increases of [Ca2+]i. The data shown are analyzed at 300 s. #, buffer versus IL8 or calphostin C plus IL8, p < 0.001; *** and **, IL8 versus IL8 plus agents indicated, p < 0.001 and p < 0.005, respectively. Cell numbers are presented in the parentheses. Data are mean ± S.E.

 
Inactivation of CD38 by ADP-ribosylation Abolishes the Sustained Rise of [Ca2+]i Induced by IL8—We have previously demonstrated that CD38 in the activated T cells is ADP-ribosylated in the presence of exogenous {beta}-NAD+, resulting in a loss of enzyme activity (6). To further elucidate the role of CD38/cADPR in IL8-induced Ca2+ signaling, functional CD38-null (CD38-) LAK cells were prepared by treatment with {beta}-NAD+. As shown in Fig. 3A, ADPR-cyclase activities, which were observed in the lysates and CD38 immunoprecipitates prepared from the control LAK cells, were completely eradicated by the incubation with {beta}-NAD+. These results indicated that CD38 represents the only ADPR-cyclase in LAK cells but also indicate that CD38 is inactivated in the presence of {beta}-NAD+, consistent with previous observations (6). When IL8-mediated Ca2+ signal in CD38- LAK cells was compared with the control LAK cells (Fig. 3B), CD38- LAK cells showed only the initial rise of [Ca2+]i but not the sustained rise of [Ca2+]i by the treatment with IL8 (Fig. 3C). To ensure that the sustained Ca2+ signal is mediated by cADPR, changes of [Ca2+]i in the CD38- LAK cells were determined in the presence of cADPR. Indeed, cADPR was able to induce the sustained Ca2+ signal (Fig. 3D). As summarized in Fig. 3, E and F, the sustained Ca2+ increase is due to the activation of CD38/cADPR through IL8 signaling.



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  		FIG. 3.
Functional inactivation of CD38 results in an absence of sustained increase of [Ca2+]i induced by IL8. A, the treatment of LAK cells with {beta}-NAD+ abolishes completely ADPR-cyclase activity. LAK cells were incubated with and without 100 µM {beta}-NAD+ for 6 h. ADPR-cyclase activity was determined in cell lysates or CD38 immunoprecipitates (IP) using NGD+ as a substrate. The mean ± S.E. of three independent experiments is shown. NA, no activity found. B, IL8 increases a sustained intracellular Ca2+ level in LAK cells. C, CD38- LAK cells induces the immediate burst increase of [Ca2+]i but not the sustained Ca2+ increase in response to IL8 treatment. D, cADPR induces the sustained increase of [Ca2+]i. Three representative Ca2+ traces are shown. Arrows indicate the time points of additions of 10 pM IL8 or 250 µM cADPR. E, a direct comparison of [Ca2+]i during initial increases of [Ca2+]i. The data shown are analyzed at 54 s. #, buffer versus IL8 or NAD plus IL8, p < 0.001. F, a direct comparison of mean [Ca2+]i during sustained increases of [Ca2+]i. The data shown are analyzed at 300 s. #, buffer versus IL8 or NAD plus cADPR, p < 0.001; ***, IL8 versus NAD plus IL8, p < 0.001. Cell numbers are presented in the parentheses. Data are mean ± S.E.

 
CD38 Is Activated by cGMP Produced by IL8/Ca2+ Signaling—It has been demonstrated that ADPR-cyclases including CD38 are probably activated by cAMP or cGMP (13, 17). Stimulation of IL8R produces two second messengers, IP3 and diacylglycerol. There is a report that IL8R may couple with Gs (26). To elucidate activation mechanism of CD38 by IL8 signaling, we determined [cADPR]i in LAK cells treated with various agonistic agents. As presented in Fig. 4A, a membrane-permeable cGMP analog, 8-pCPT-cGMP, significantly increased the level of [cADPR]i that was comparable with that induced by IL8. Moreover, a membrane-permeable PKG inhibitor, Rp-8-pCPT-cGMPS, blocked IL8-induced cADPR formation. On the other hand, phorbol 12-myristate 13-acetate or the cAMP analog, 8-CPT-cAMP, or Ca2+ ionophore, ionomycin, was not able to increase [cADPR]i. We further examined whether cGMP is produced by stimulation of IL8R. The treatment of IL8 indeed increased the intracellular cGMP level, and the IL8-induced cGMP increase was completely blocked by xestospongin C but not by calphostin C (Fig. 4B). Supporting the result that ionomycin had no effect on cADPR formation, cGMP level was also not increased by the ionomycin treatment. The observations that initial Ca2+ rise proceeded to increase the sustained Ca2+ signal (Fig. 2, A and B) indicated that cGMP increase induced by IL8 may proceed to form cADPR. The time course of cGMP and cADPR formation induced by IL8 was evaluated. The results showed that levels of cGMP and cADPR were increased in a time-dependent manner, reaching maximal levels at ~60 and 90 s, respectively (Fig. 4C). The formation of cGMP was rapidly reduced, whereas the formation of cADPR was slowly decreased. Supporting these results, a cGMP analog, 8-pCPT-cGMP, generated the sustained increase of [Ca2+]i (Fig. 4D). The cGMP analog-mediated increase of [Ca2+]i was completely blocked by pretreatment with 8-Br-cADPR (Fig. 4E) but not by xestospongin C (Fig. 4F). Moreover, Rp-8-pCPT-cGMPS blocked IL8-induced sustained Ca2+ rise but not the initial Ca2+ rise (Fig. 4G). Differences in [Ca2+]i during initial and sustained increases of [Ca2+]i were summarized in Fig. 4, H and I, respectively. These findings indicate that cGMP is involved in the activation of CD38 in LAK cells and that IL8-induced cGMP formation is due to IP3-mediated Ca2+ rise. In addition, we also assessed whether phosphorylation of CD38 via cGMP/PKG during IL8 treatment occurred. Phosphorylation of CD38 was not observed. The increased production of cGDPR by CD38 isolated from LAK cells by immunoprecipitation with CD38 antibody was not observed in the presence of cGMP, PKG, or both together or Ca2+ (data not shown).



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  		FIG. 4.
IL8-induced cADPR formation is mediated by cGMP/PKG. A, cGMP increases the level of [cADPR]i. Levels of cADPR were determined after the treatment of LAK cells with 10 pM IL8, 100 nM phorbol 12-myristate 13-acetate (PMA), 1 mM 8-CPT-cAMP, 1 mM 8-pCPT-cGMP, or 200 nM ionomycin for 90 s. Prior to incubating with IL8 for 90 s, Rp-8-pCPT-cGMPS (20 µM) was preincubated for 30 min. The reactions were terminated by the addition of 0.5 ml of 0.6 M perchloric acid at time point indicated. The level of cADPR was determined as detailed under "Experimental Procedures." The mean ± S.E. of three independent experiments is shown. *, p < 0.05; **, p < 0.005. B, IL8-mediated cGMP formation is blocked by xestospongin C but not calphostin C or ionomycin. Cells were preincubated with 2 µM xestospongin C or 100 nM calphostin C for 30 min prior to incubating with IL8 (10 pM) for 60 s. Ionomycin (200 nM) was incubated for 60 s. The reactions were stopped by the addition of equal volume of 12% trichloroacetic acid. The level of cADPR was determined as detailed under "Experimental Procedures." The mean ± S.E. of three independent experiments is shown. *, p < 0.05; **, p < 0.005. C, increase of cGMP levels induced by IL8 precedes cADPR formation. D, cell-permeable cGMP analog induces a sustained increase of [Ca2+]i. The concentration of 8-pCPT-cGMP was 1 mM. E, antagonistic cADPR blocks cGMP-induced sustained Ca2+ increase. The concentration of 8-Br-cADPR was 100 µM. F, cGMP-induced sustained Ca2+ increase is not blocked by pretreatment with xestospongin C (2 µM). G, a sustained Ca2+ rise induced by IL8 is blocked by a PKG inhibitor. Rp-8-pCPT-cGMPS was 20 µM. Arrows indicate the time point of the addition of agents. H, a direct comparison of mean [Ca2+]i during initial increases of [Ca2+]i. The data shown are analyzed at 54 s. #, buffer versus 8-pCPT-cGMP or Rp-8-pCPT-cGMPS plus IL8, p < 0.001; ***, 8-pCPT-cGMP versus 8-Br-cADPR plus 8-pCPT-cGMP or xestospongin C plus 8-pCPT-cGMP, p < 0.001. I, a direct comparison of mean [Ca2+]i during sustained increases of [Ca2+]i. #, buffer versus 8-pCPT-cGMP or Rp-8-pCPT-cGMPS plus IL8, p < 0.001; ***, 8-pCPT-cGMP versus 8-Br-cADPR plus 8-pCPT-cGMP or xestospongin C plus 8-pCPT-cGMP, p < 0.001. The data shown are analyzed at 300 s. Cell numbers are presented in the parentheses. Data are mean ± S.E.

 
cGMP/PKG/cADPR Signaling System Is Essential for IL8-induced Migratory Activity of LAK Cells—It is well known that the activation of IL8R induces migration of various cell types (19, 20). To examine whether CD38/cADPR plays any role in IL8-mediated cell migration, we evaluated the effect of cADPR on the migratory activity of LAK cells. As presented in Fig. 5A, LAK cell migration was significantly induced by IL8. The IL8-mediated cell migration was blocked by pretreatment with 8-Br-cADPR. In addition, we also examined the effects of Ca2+ on LAK cell migration (Fig. 5A). IL8-induced cell migration was not observed in the absence of extracellular Ca2+, and the increase of [Ca2+]i by ionomycin did not induce cell migration.



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  		FIG. 5.
CD38/cADPR mediates IL8-stimulated migratory activity of LAK cells. A, IL8-induced stimulation of LAK migration is blocked by antagonistic cADPR analog 8-Br-cADPR. Cells treated with 100 µM 8-Br-cADPR for 30 min were incubated with IL8 (10 pM). Hanks' balanced salt solution containing 3 mM EGTA was used to deplete extracellular Ca2+, and ionomycin was 200 nM. The cell migration assay was performed as detailed under "Experimental Procedures." *, p < 0.05; **, p < 0.005. B, IL8-induced migratory activity of CD38- LAK cells is absent. Incubation of CD38- LAK cells with cADPR (250 µM) stimulates cell migration. *, p < 0.05; ** p < 0.005. C, cGMP/PKG stimulates migration of LAK cells. Cells pretreated with Rp-8-pCPT-cGMPS (20 µM) for 30 min were incubated with IL8 (10 pM). For the determination of cGMP-mediated cell migration, the cell-permeable cGMP analog, 8-pCPT-cGMP (1 mM), was used. **, p < 0.005.

 
The above observations that CD38/cADPR is involved in LAK cell migration were further examined using CD38- LAK cells. IL8-meidated migration of CD38- LAK cells was not observed (Fig. 5B). However, the treatment of CD38- LAK cells with exogenous cADPR stimulated cell migration similar to the control LAK cells induced by IL8. To further support the observation that cGMP is able to activate CD38, thus resulting in an increase of cADPR production, cGMP-mediated LAK cell migration was examined. An agonistic membrane-permeable cGMP analog, 8-pCPT-cGMP, stimulated the migratory activity of LAK cells independent of the activation of IL8R (Fig. 5C). The pretreatment of PKG inhibitor, Rp-8-pCPT-cGMPS, completely blocked IL8-induced migration of LAK cells. These results indicate that CD38/cADPR plays an important role in IL8-mediated migration of LAK cells.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
CD38 is a bifunctional enzyme having ADPR-cyclase and hydrolase activity that produces and hydrolyzes cADPR, which is a powerful and universal Ca2+-mobilizing second messenger. Studies have proposed that CD38, including ADPR-cyclases, is activated by G-protein-coupled receptor. However, the regulation pathway(s) of CD38 by G-protein-coupled receptor remains unclear. In this study, we for the first time demonstrate that CD38 induced in LAK cells is stimulated by cGMP/PKG that is generated upon the activation of IL8R. Our results have also revealed that a CD38 metabolite, cADPR, plays an essential role in regulation of [Ca2+]i and migratory activity of LAK cells.

The regulation pathway of CD38 in IL8 signaling is summarized in Fig. 6. IL8R couples with Gi and stimulates PLC-{beta}2, producing two second messengers, IP3 and diacylglycerol (24, 25). The IL8 treatment of LAK cells exhibits a rapid rise of [Ca2+]i that sustains for a long period of time (>10 min). Our results show that the initial Ca2+ signal is mediated by IP3 and that the IL8-mediated sustained Ca2+ signal is due to the activation of CD38, resulting in an increase of [cADPR]i. Thus, the pretreatment of cells with 8-Br-cADPR, an antagonistic analog of cADPR, displays only the initial rise of [Ca2+]i. An IP3R blocker, xestospongin C, abolishes completely the IL8-mediated elevation of the initial and sustained [Ca2+]i, indicating that the activation of CD38 requires the IP3-mediated increase of [Ca2+]i. However, in contrast to IL8-induced formation of cADPR via IP3-mediated Ca2+ rise, nonphysiological elevation of [Ca2+]i by ionomycin does not increase the level of [cADPR]i. Consistent with our results, a previous report demonstrates that increase of [Ca2+]i by thapsigargin or ionomycin is not able to elevate the level of [cADPR]i, whereas the treatment of human Jurkat T cells with agonistic OKT3 antibody increases the level of cADPR as well as the sustained Ca2+ signal (10). Together, it appears that CD38 in LAK cells, including T cells, is not activated unspecifically by increased [Ca2+]i. Our results also indicate that cADPR induces Ca2+ influx in LAK cells. The depletion of extracellular Ca2+ using EGTA generates the initial Ca2+ rise but not the sustained Ca2+ signal in response to IL8. CD38/cADPR-mediated Ca2+ entry has been observed with neutrophils (11) and intact human T cells (36). A recent study has reported that type 3 RyR plays an essential role in the sustained Ca2+ response in T cells (37). Although the molecular mechanism by which cADPR-mediated sustained Ca2+ rise remains to be clarified, our results strongly suggest that cADPR produced by CD38 is a Ca2+-mobilizing second messenger and mediates the sustained phase of Ca2+ signal by means of Ca2+ influx in LAK cells.



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  		FIG. 6.
Schematic presentation of IL8-mediated CD38 activation in LAK cell. Ligation of IL8R receptor stimulates PLC-{beta}2 via Gi, resulting in the production of IP3 and diacylglycerol (24, 25). IP3 binding to the receptor releases Ca2+ from the intracellular Ca2+ stores. The IP3-mediated increase of the intracellular Ca2+ level results in the elevation of the cGMP level via guanylyl cyclase (GC), thus activating PKG. cGMP/PKG-mediated signaling stimulates intracellular cADPR production via CD38. cADPR-mediated Ca2+ influx is requisite for LAK cell migration. cADPR may act on RyR present on Ca2+ store to regulate the sustained Ca2+ rise in LAK cells since type 3 RyR in T cells regulates the sustained Ca2+ response (37).

 
Studies have indicated that cAMP activates the production of cADPR in artery smooth muscle cell (13) and that cGMP is the activator of A. califonica ADPR-cyclase (17). Our results suggest that the activation of CD38 expressed in LAK cells is mediated by PKG activation via cGMP generated by IL8 signaling. Thus, the treatment of various agents shows that cGMP increases [cADPR]i in LAK cells. Consistent with these results, cGMP induces the sustained rise of [Ca2+]i, and a PKG blocker, Rp-8-pCPT-cGMPS, abolishes the sustained Ca2+ signal but not the initial Ca2+ signal in response to IL8. More importantly, IL8-induced increase of cGMP precedes the production of cADPR. These results demonstrate that the activation of IL8R increases cGMP production in LAK cells and that cGMP/PKG regulates CD38 activity. However, it appears that cGMP/PKG does not directly act on the enzyme; neither PKG-mediated phosphorylation of CD38 nor cGMP-mediated activation of isolated CD38 using the CD38 antibody is observed.

Our results have indicated that CD38/cADPR plays a critical role in the IL8-mediated migration of LAK cells. Thus, IL8-mediated cell migration is blocked by an antagonistic cADPR analog, 8-Br-cADPR, and by inactivation of CD38 with {beta}-NAD+. Moreover, cADPR alone is able to induce LAK cell migration. Consistent with the observations that cGMP/PKG signaling activates CD38, the pretreatment of the PKG inhibitor blocks IL8-mediated migration of LAK cells. We have also found that IL8-induced LAK cell migration is strictly controlled by Ca2+ influx; in the absence of extracellular Ca2+, IL8-induced migration of LAK cells is not observed, and nonphysiological increase of [Ca2+]i with ionomycin does not induce cell migration. A previous study has also indicated that dendritic cell trafficking depends on Ca2+ influx induced by CD38/cADPR (38). Studies have demonstrated that the biological role of CD38/cADPR in LAK cells as well as natural killer cells isolated is to trigger lytic and secretory responses (27). This cytotoxic role of CD38 has been observed by direct ligation of CD38 with an agonist antibody IB4 (39). The main signaling event in the cytotoxicity seems to be an involvement of protein tyrosine kinase(s) signaling via binding of the agonistic antibody IB4 to CD38 (39, 40). Taken together, CD38 plays important roles in migration and cytotoxicity of LAK cells, including natural killer cells. The present study shows the regulation-signaling pathway of CD38 through the sequential coupling of IL8R, Ca2+ and cGMP/PKG and describes the biological role of CD38 in this signaling pathway, which is involvement in IL8-induced cell migration through the generation of cADPR/sustained Ca2+ elevation.


   FOOTNOTES
 
* This work was supported by the Korea Science and Engineering Foundation Grant R01-2000-000-00134-0 (to U.-H. K.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

§ Present address: Dept. of Microbiology, Chonbuk National University Medical School, Jeonju, 561-182, Republic of Korea. Back

|| To whom correspondence should be addressed: Dept. of Biochemistry, Chonbuk National University Medical School, Keumam-dong, Jeonju, 561-182, Republic of Korea. Tel.: 82-63-270-3083; Fax: 82-63-274-9833; E-mail: uhkim{at}chonbuk.ac.kr.

1 The abbreviations used are: ADPR, ADP-ribose; ADP-ribosyl cyclase, ADPR-cyclase; cADPR, cyclic ADPR; cGDPR, cyclic GDP-ribose; 8-CPT-cAMP, 8-(4-chlorophenylthio)-adenosine-3',5'-cyclic monophosphate; 8-Br-cADPR, 8-bromo-cADPR; 8-pCPT-cGMP, 8-(4-chlorophenylthio)-guanosine-3',5'-cyclic monophosphate; Rp-8-pCPT-cGMPS, 8-(4-chlorophenylthio)-guanosine-3',5'-cyclic monophosphorothioate, Rp-isomer; IL, interleukin; IL8R, interleukin 8 receptor; IP3, inositol 1,4,5-trisphosphate; IP3R, IP3 receptor; LAK, lymphokine-activated killer; {beta}-NAD+, {beta}-nicotinamide adenine dinucleotide; NGD, nicotinamide guanine nucleotide; PKG, protein kinase G; RyR, ryanodine receptor; PLC, phospholipase C; BSA, bovine serum albumin; HPLC, high performance liquid chromatography. Back


   ACKNOWLEDGMENTS
 
We appreciate very much the volunteers who donated their blood for the study.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Lund, F., Solvason, N., Grimaldi, J. C., Parkhouse, R. M., and Howard, M. (1995) Immunol. Today 16, 469-473[CrossRef][Medline] [Order article via Infotrieve]
  2. Mehta, K., Shahid, U., and Malavasi, F. (1996) FASEB J. 10, 1408-1417[Abstract]
  3. Kim, H., Jacobson, E. L., and Jacobson, M. K. (1993) Science 261, 1330-1333[Abstract/Free Full Text]
  4. Howard, M., Grimaldi, J. C., Bazan, J. F., Lund, F. E., Santos-Argumedo, L., Parkhouse, R. M., Walseth, T. F., and Lee, H. C. (1993) Science 262, 1056-1059[Abstract/Free Full Text]
  5. Zocchi, E., Franco, L., Guida, L., Benatti, U., Bargellesi, A., Malavasi, F., Lee, H. C., and De Flora, A. (1993) Biochem. Biophys. Res. Commun. 196, 1459-1465[CrossRef][Medline] [Order article via Infotrieve]
  6. Han, M. K., Cho, Y. S., Kim, Y. S., Yim, C. Y., and Kim, U. H. (2000) J. Biol. Chem. 275, 20799-20805[Abstract/Free Full Text]
  7. Lee, H. C. (2001) Annu. Rev. Pharmacol. Toxicol. 41, 317-345[CrossRef][Medline] [Order article via Infotrieve]
  8. Galione, A., Lee, H. C., and Busa, W. B. (1991) Science 253, 1143-1146[Abstract/Free Full Text]
  9. Lee, H. C., Aarhus, R., and Graeff, R. M. (1995) J. Biol. Chem. 270, 9060-9066[Abstract/Free Full Text]
  10. Guse, A. H., da Silva, C. P., Berg, I., Skapenko, A. L., Weber, K., Heyer, P., Hohenegger, M., Ashamu, G. A., Schulze-Koops, H., Potter, B. V. L., and Mayr, G. W. (1999) Nature 398, 70-73[CrossRef][Medline] [Order article via Infotrieve]
  11. Partida-Sanchez, S., Cockayne, D. A., Monard, S., Jacobson, E. L., Oppenheimer, N., Garvy, B., Kusser K., Goodrich, S., Howard M., Harmsen, A., Randall, T. D., and Lund, F. E. (2001) Nat. Med. 7, 1209-1216[CrossRef][Medline] [Order article via Infotrieve]
  12. Higashida, H., Egorova, A., Higashida, C., Zhong, Z. G., Yokoyama, S., Noda, M., and Zhang, J. S. (1999) J. Biol. Chem. 274, 33348-33353[Abstract/Free Full Text]
  13. Boittin, F. X., Dipp, M., Kinnear, N. P., Galione, A., and Evans, A. M. (2003) J. Biol. Chem. 278, 9602-9608[Abstract/Free Full Text]
  14. Higashida, H., Zhang, J. S., Hashii, M., Shintaku, M., Higashida, C., and Takeda, Y. (2000) Biochem. J. 352, 197-202
  15. Higashida, H., Yokoyama, S., Hashii, M., Taketo, M., Higashida, M., Takayasu, T., Ohshima, T., Takasawa, S., Okamoto, H., and Noda, M. (1997) J. Biol. Chem. 272, 31272-31277[Abstract/Free Full Text]
  16. Sternfeld, L., Krause, E., Guse, A. H., and Schulz, I. (2003) J. Biol. Chem. 278, 33629-33636[Abstract/Free Full Text]
  17. Graeff, R. M., Franco, L., De Flora, A., and Lee, H. C. (1998) J. Biol. Chem. 273, 118-125[Abstract/Free Full Text]
  18. Inngjerdingen, M., Al-Aoukaty, A., Damaj, B., and Maghazachi, A. A. (1999) Biochem. Biophys. Res. Commun. 262, 467-472[CrossRef][Medline] [Order article via Infotrieve]
  19. Huber, A. R., Kunkel, S. L., Todd, R. F., and Weiss, S. J. (1991) Science 254, 99-102[Abstract/Free Full Text]
  20. Baggiolini, M., Loetscher, P., and Moser, B. (1995) Int. J. Immunopharmacol. 17, 103-108[CrossRef][Medline] [Order article via Infotrieve]
  21. Oppenheim, J. J., Zachariae, C. O., Mukaida, N., and Matsushima, K. (1991) Annu. Rev. Immunol. 9, 617-648[Medline] [Order article via Infotrieve]
  22. Koch, A. E., Polverini, P. J., Kunkel, S. L., Harlow, L. A., DiPietro, L. A., Elner, V. M., Elner, S. G., and Strieter, R. M. (1992) Science 258, 1798-1801[Abstract/Free Full Text]
  23. Baggiolini, M., Dewald, B., and Moser, B. (1997) Annu. Rev. Immunol. 15, 675-705[CrossRef][Medline] [Order article via Infotrieve]
  24. Wu, D., LaRosa, G. J., and Simon, M. I. (1993) Science 261, 101-103[Abstract/Free Full Text]
  25. Neptune, E. R., Iiri, T., and Bourne, H. R. (1999) J. Biol. Chem. 274, 2824-2828[Abstract/Free Full Text]
  26. Maghazachi, A. A. (2000) Int. J. Biochem. Cell Biol. 32, 931-943[CrossRef][Medline] [Order article via Infotrieve]
  27. Mallone, R., Funaro, A., Zubiaur, M., Baj, G., Ausiello, C. M., Tacchetti, C., Sancho, J., Grocci, C., and Malavasi, F. (2001) Int. Immunol. 13, 397-409[Abstract/Free Full Text]
  28. Sconocchia, G., Titus, J. A., Mazzoni, A., Visintin, A., Pericle, F., Hicks, S. W., Malavasi, F., and Segal, D. M. (1999) Blood 94, 3864-3871[Abstract/Free Full Text]
  29. Melder, R. J., Whiteside, T. L., Vujanovic, N. L., Hiserodt, J. C., and Herberman, R. B. (1988) Cancer Res. 48, 3461-3469[Abstract/Free Full Text]
  30. Sebok, K., Woodside, D., Al-Aoukaty, A., Ho, A. D., Gluck, S., and Maghazachi, A. A. (1993) J. Immunol. 150, 1524-1534[Abstract]
  31. Lee, H. C., and Aarhus, R. (1991) Cell Regul. 2, 203-209[Medline] [Order article via Infotrieve]
  32. Graeff, R. M., Walseth, T. F., Fryxell, K., Branton, W. D., and Lee, H. C. (1994) J. Biol. Chem. 269, 30260-30267[Abstract/Free Full Text]
  33. Graeff, R., and Lee, H. C. (2002) Biochem. J. 361, 379-384[CrossRef][Medline] [Order article via Infotrieve]
  34. Xie, G. H., Rah, S. Y., Yi, K. S., Han, M. K., Chae, S. W., Im, M. J., and Kim, U. H. (2003) Biochem. Biophys. Res. Commun. 307, 713-718[CrossRef][Medline] [Order article via Infotrieve]
  35. Tsien, R. Y., Pozzan, T., and Rink, T. J. (1982) Nature 295, 68-70[CrossRef][Medline] [Order article via Infotrieve]
  36. Guse, A. H., Berg, I., da Silva, C. P., Potter, B. V. L., and Mayr, G. W. (1997) J. Biol. Chem. 272, 8546-8550[Abstract/Free Full Text]
  37. Schwarzmann, N., Kunerth, S., Weber, K., Mayr, G. W., and Guse, A. H. (2002) J. Biol. Chem. 277, 50636-50642[Abstract/Free Full Text]
  38. Partida-Sanchez, S., Goodrich, S., Kusser, K., Oppenheimer, N., Randall, T. D., and Lund, F. E. (2004) Immunity 20, 279-291[CrossRef][Medline] [Order article via Infotrieve]
  39. Cesano, A., Visonneau, S., Deaglio, S., Malavasi, F., and Santoli, D. (1998) J. Immunol. 160, 1106-1115[Abstract/Free Full Text]
  40. Zubiaur, M., Izquierdo, M., Terhorst, C., Malavasi, F., and Sancho, J. (1997) J. Immunol. 159, 193-205[Abstract]

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