iPLA₂β: front and center in human monocyte chemotaxis to MCP-1

The antisense oligodeoxyribonucleotides (AS-ODN) used in our previous study to identify the involvement of iPLA₂ was directed against the form of iPLA₂ that was eventually classified as iPLA₂β (14). iPLA₂ isoforms (β/γ) display different sensitivities to R or S enantiomers of the pharmacological inhibitor bromoenol lactone (BEL), with iPLA₂β being 10 times more sensitive to (S)-BEL than to (R)-BEL (15). As predicted, (S)-BEL caused stronger reduction of MCP-1–induced monocyte chemotaxis compared with (R)-BEL at all concentrations tested, providing additional evidence that monocyte chemotaxis to MCP-1 is indeed regulated by iPLA₂β (Fig. 1 A).

Unstimulated monocytes showed different morphologies that varied from spherical to polar with diffuse, cytoplasmic iPLA₂β staining (Fig. 1, C, E, and G, left). Upon MCP-1 stimulation, migrating cells displayed polar morphology, and iPLA₂β showed increased distribution at the cell periphery, particularly in the membrane-enriched pseudopod (Fig. 1, C, E, G, right). Quantification of mean fluorescent intensity of iPLA₂β in the tail, midbody, and pseudopod of migrating monocytes suggests that, in the absence of MCP-1, iPLA₂β was localized predominantly in the tail of monocytes; however, in response to MCP-1, the distribution was reversed and the majority of iPLA₂β was detected in the pseudopod of moving cells (Fig. 1 B). These observations demonstrate that MCP-1 induces preferential recruitment of iPLA₂β from the tail to the pseudopod of migrating monocytes.

Because MCP-1–induced chemotaxis of iPLA₂β-deficient cells could be rescued with LPA (14), which is known to induce focal adhesion assembly (16), it is likely that iPLA₂β affects monocyte migration by interacting with an essential component of the pseudopod. Therefore, we evaluated the colocalization of iPLA₂β with F-actin and Cdc42, which are two known markers of the pseudopod (Fig. 1, C and E). In the presence of MCP-1, F-actin was enriched in the pseudopod of migrating monocytes and showed increased colocalization with iPLA₂β (Fig. 1 C, right, merged image). Quantitative evaluation showed that the MCP-1–induced increased colocalization of F-actin with iPLA₂β was not dependent on the polar morphology of monocytes (Fig. 1 D, filled versus black-hatched bar), and that it correlated with significantly enhanced colocalization of iPLA₂β with Cdc42 (Fig. 1 E and 1F). Together, these results show that MCP-1 induces redistribution of iPLA₂β in nonpolar monocytes that eventually culminates in the increased presence of iPLA₂β in the pseudopod of polar monocytes.

Association of iPLA₂β with the cell membrane was evaluated by visualizing the colocalization of iPLA₂β with Na-K-ATPase (Fig. 1 G). Quantitative evaluation suggested that MCP-1–treated polar monocytes displayed significantly higher colocalization of iPLA₂β with Na-K-ATPase compared with untreated monocytes (Fig. 1 H). To rule out the possibility that this was an artifact caused by the small size of the monocytes, the percentage distribution of iPLA₂β associated with the cell membrane was also determined by quantifying pixel intensity of iPLA₂β on the peripheral cell membrane and in the whole cell (17). As shown in Fig. S1 , MCP-1 induced approximately twofold induction of iPLA₂β recruitment to the cell membrane in both polar or nonpolar monocytes, suggesting that recruitment of iPLA₂β to the cell membrane is specifically induced by MCP-1, regardless of cell polarity.

Because of the stimulus and cell type–dependent variability in intracellular locations of cPLA₂α, we initially explored the recruitment of cPLA₂α to early endosomes, Golgi, and endoplasmic reticulum by costaining with Rab 5, lectin GS-II, and protein disulfide isomerase (PDI), respectively. cPLA₂α showed minimal or no localization with the Golgi or early endosomes (unpublished data). In the absence of MCP-1, cPLA₂α was distributed throughout the cytoplasm (Fig. 2 A, left) and did not reveal specific colocalization with PDI (Fig. 2 A, arrowhead). In MCP-1–stimulated cells, cPLA₂α was concentrated in the posterior region of the cell and colocalized with PDI (Fig. 2 A, right). MCP-1 induced a significant increase in recruitment of cPLA₂α to the endoplasmic reticulum, irrespective of cell polarity (Fig. 2 B, filled versus black-hatched bar).

To further substantiate that MCP-1 induces localization of iPLA₂β and cPLA₂α to different intracellular sites, both enzymes were visualized together, and the percentage of colocalization of the phospholipases was determined. In MCP-1–stimulated monocytes, cPLA₂α was concentrated in the posterior main body of the cell; in contrast, iPLA₂β was concentrated in the pseudopod (Fig. 2 C, right, merged image). Furthermore, the cPLA₂α-rich posterior region was relatively devoid of iPLA₂β staining, whereas the iPLA₂β-enriched pseudopod showed minimal cPLA₂α staining (Fig. 2 C, right, merged image, arrow versus arrowhead). The colocalization of iPLA₂β with cPLA₂α (28 ± 6%), or vice versa (32 ± 6%), remained statistically unaltered because of MCP-1 treatment (Fig. 2 D).

In summary, microscopic observations reveal that MCP-1 induces redistribution of both enzymes; cPLA₂α is recruited to the endoplasmic reticulum, whereas iPLA₂β is recruited to the membrane-enriched pseudopod of monocytes, and these phospholipases do not colocalize.

To gather insight as to how cPLA₂α and iPLA₂β modify chemotaxis of monocytes to MCP-1, we evaluated their effects on the net distance traveled by monocytes in a gradient of MCP-1 using an under-agarose assay. For these studies, we used previously characterized antisense ODNs to assess the effect of iPLA₂β and cPLA₂α deficiency on MCP-1–induced chemotaxis of monocytes. It is important to note that these antisense ODNs have been shown to specifically target these enzymes and do not modulate the expression of CCR2 (14, 18, 19). Monocytes were allowed to respond to MCP-1 for 1 h, fixed, and analyzed for net distance. Net distance was determined by measuring the distance of all migrated cells from the periphery of the well to their final position, and it reflects the cumulative effect of change in speed and direction. As shown (Fig. 3), MCP-1 increased the chemotaxis of monocytes and the migrated cells traveled to 1.82-fold greater net distance (3,082.2 μm) than unstimulated cells (1,722 μm). When treated with iPLA₂β antisense ODN, 98.7% reduction was observed in MCP-1–induced chemotaxis (Fig. 3 A). These monocytes displayed reduced net distance (88%) in response to MCP-1 (1,631.4 μm) compared with MCP-1–stimulated cells that were not treated with ODN (Fig. 3 B). Similarly, inhibition of cPLA₂α expression with antisense ODN caused 94% reduction in MCP-1–induced chemotaxis (Fig. 3 A) and 84% inhibition of net distance traveled compared with ODN-untreated, MCP-1–unstimulated cells (Fig. 3 B). These observations suggest that, upon MCP-1 stimulation, more monocytes migrate toward MCP-1, and that migrating monocytes travel twice the distance compared with unstimulated cells.

Reduced net distance of migration of monocytes rendered deficient in phospholipase A₂ by antisense ODN, as described in the previous section, in response to MCP-1 (Fig. 3 B) can be caused by a change in the total distance, speed, or net migration of monocytes toward MCP-1 (net-X distance). Therefore, the relative contribution of iPLA₂β and cPLA₂α to these parameters was delineated by quantitative analysis of time-lapse photomicrographs.

As shown in wind-rose plots (Fig. 4 A), in the absence of MCP-1, monocytes showed random migration (top left). Upon MCP-1 stimulation, cells migrated toward the source of MCP-1 and traveled longer distances (bottom left). Cells treated with sense ODN to iPLA₂β migrated similarly to untreated cells (Fig. 4 A, middle). Interestingly, monocytes treated with iPLA₂β antisense ODN showed a more clustered migratory pattern both in the absence or presence of MCP-1 (Fig. 4 A, right top and bottom). Quantitative analysis of all cells that could be tracked for 60 min (n) revealed that in response to MCP-1, monocytes migrated significantly faster than unstimulated monocytes and equally to sense ODN–treated cells (Fig. 4 A). In contrast, treatment of monocytes with antisense ODN to iPLA₂β significantly decreased the speed of migrating monocytes from 7.58 ± 0.62 to 5.93 ± 0.49 m/min (Fig. 4 A). Furthermore, iPLA₂β antisense ODN–treated cells showed a significant reduction in net distance, whereas the net distance traveled by sense ODN-treated monocytes remained unaltered (Fig. 4 C). These findings confirm the results presented in Fig. 3.

We reasoned that speed or net distance merely reflect migration of monocytes, irrespective of their direction. In contrast, the ability of monocytes to reach inflamed sites depends on their ability to navigate toward the source of MCP-1. Hence, the contribution of iPLA₂β in directed migration of monocytes toward MCP-1 was determined by quantification of net-X distance that denotes the net value of the X coordinate toward MCP-1. As shown in Fig. 4 D, MCP-1 induced a fivefold induction in net-X. Treatment of monocytes with sense ODN to iPLA₂β had no effect; in contrast, monocytes treated with antisense ODN to iPLA₂β displayed significant reduction in net-X.

To further probe whether the loss of directionality contributed to the reduced net-X distance (Fig. 4 D) and clustered migratory pattern of iPLA₂β antisense ODN-treated monocytes (Fig. 4 A, bottom right), turn analysis was performed. As shown in Fig. 4 E, MCP-1–treated monocytes make fewer wide turns (≥30 degrees, 9 ± 0.81), which is similar to monocytes treated with sense ODN to iPLA₂β (9.66 ± 0.76). In contrast, iPLA₂β antisense ODN–treated monocytes make more frequent wide turns (>30 degrees, 14.2 ± 1.39). The MCP-1–stimulated monocytes without the ODN treatment showed a significant reduction in average angle per turn from 71.08 ± 7.37 to 35.54 ± 4.18 degrees compared with monocytes without MCP-1 (Fig. 4 F). This effect was totally abrogated in iPLA₂β antisense ODN–treated monocytes (Fig. 4 F), suggesting that iPLA₂β is a contributor to the cellular compass.

As shown in Fig. 5 A, wind-rose plots of untreated or monocytes treated with sense ODN to cPLA₂α showed random migration in the absence of MCP-1 and directed migration in the presence of MCP-1. Migratory patterns of monocytes treated with antisense ODN to cPLA₂α were more clustered than the other groups. Quantitative analysis of the migratory patterns of the maximum number of monocytes (n) that could be tracked for 60 min revealed that antisense ODN to cPLA₂α significantly reduced the speed of monocytes (3.96 ± 0.28 μm/min) compared with ODN-untreated, MCP-1–stimulated cells (7.58 ± 0.62 μm/min; Fig. 5 B). cPLA₂α antisense ODN–treated cells also displayed significantly reduced net distance (Fig. 5 C) and net migration toward MCP-1 (net–X distance; Fig. 5 D) compared with either no ODN or sense ODN–treated monocytes. Notably, cPLA₂α antisense ODN-treated cells displayed no significant reduction in the number of wide (≥30 degrees) turns or the average angle per turn compared with controls (Fig. 5, E and F). These observations suggest that cPLA₂α regulates speed, but not directionality, of monocyte chemotaxis to MCP-1.

MCP-1–induced recruitment of iPLA₂β to the F-actin–rich pseudopod and the significant loss of directionality of iPLA₂β antisense ODN–treated monocytes suggested that iPLA₂β might be involved in actin polymerization. We tested this possibility by evaluating the effect of iPLA₂β inhibition on MCP-1–induced actin polymerization. MCP-1 induced a significant increase in the level of F-actin. Interestingly, inhibition of iPLA₂β by BEL caused significant reduction in F-actin levels (Fig. 6 A). Furthermore, iPLA₂β antisense ODN–treated monocytes displayed a significant reduction in actin polymerization compared with ODN-untreated or iPLA₂β sense ODN–treated monocytes (Fig. 6 B). These observations indicate that iPLA₂β regulates MCP-1–induced actin polymerization.

We developed a novel adoptive transfer mouse model to validate the regulatory functions of iPLA₂β and cPLA₂α in monocyte chemotaxis to MCP-1 in vivo. It is based on the following: (a) migration of monocytes to the peritoneum in thioglycolate-induced peritonitis in wild-type mice is dependent on MCP-1(7); (b) mouse peripheral blood mononuclear cell preparations do not contain neutrophils, and lymphocytes do not respond to MCP-1; (c) monocytes, by virtue of being fluorescently labeled with a stable PKH26, can be distinguished from unlabeled endogenous cells of the recipient mice; (d) we hypothesize that treatment of cells with phospholipase A₂–specific inhibitors or antisense ODNs before adoptive transfer will cause migration of fewer adoptively transferred monocytes; and (e) because adoptively transferred cells constitute only a small fraction of total blood cells (11–14%), migration of endogenous leukocytes of the recipient mouse to the peritoneum will not be affected (Fig. 7 A).

Earlier, we reported that the iPLA₂ inhibitor racemic-BEL and the cPLA₂/iPLA₂ inhibitor arachidonyl trifluoromethyl ketone (AACOCF₃) both suppressed MCP-1–induced chemotaxis of primary monocytes (14). As predicted from these in vitro data, significantly fewer AACOCF₃-treated monocytes migrated to the peritoneum in the adoptive transfer model (Fig. 7 B). Surprisingly, no significant difference was observed in the migration of monocytes treated with racemic-BEL compared with untreated monocytes. The inefficiency of racemic-BEL to inhibit monocyte migration in vivo was found to be caused by time-dependent recovery of BEL-treated monocytes at 37°C in vitro (Fig. S2). To determine whether the apparent efficacy of AACOCF₃ in vivo was caused by toxicity, we examined monocyte survival and chemotaxis to MCP-1 24 h after treatment with AACOCF₃ in vitro. As shown in Fig. S3, AACOCF₃ treatment did not affect monocyte survival, and the monocytes still displayed significant reduction in chemotaxis to MCP-1 compared with untreated monocytes. These observations suggest that effectiveness of AACOCF₃ was caused by the stable inhibitory effect of AACOCF₃ on cPLA₂α and iPLA₂β. As expected, no change was observed in either total monocytes or total leukocytes in the peritoneal lavage (Fig. 7 B).

Because AACOCF₃ inhibits both iPLA₂ and cPLA₂ and racemic-BEL proved ineffective in vivo, we tested the effect of specific antisense ODNs. iPLA₂β antisense ODN used in this study has previously been demonstrated to reduce iPLA₂β protein expression in both murine and human monocytes/macrophages (14, 18, 19). It significantly reduced the expression of iPLA₂β without affecting cPLA₂α protein expression or enzyme activity (14) and was effective in reducing monocyte chemotaxis to MCP-1 in vitro even 24 h later (unpublished data). Similarly, cPLA₂α antisense ODN reduced monocyte chemotaxis in vitro without affecting iPLA₂β protein expression or enzyme activity (14). It reduced the expression of cPLA₂α both in human and murine monocytes (Fig. S4) (14).

As shown (Fig. 8 A), adoptively transferred monocytes treated with iPLA₂β antisense ODN displayed statistically significant reduction in migration to the peritoneum compared with untreated or sense ODN–treated monocytes (top). Migration of endogenous monocytes and total leukocytes remained statistically similar. These data suggest that unlike racemic-BEL, iPLA₂β antisense ODN–treated monocytes displayed sustained reduced migration in vivo.

To evaluate the in vivo requirement for cPLA₂α in MCP-1 chemotaxis, adoptively transferred cells were pretreated with antisense ODN to cPLA₂α and monitored for their response to thioglycolate. As shown in Fig. 8 B (top), significantly fewer cPLA₂α antisense ODN–treated cells migrated to the peritoneum compared with untreated or cPLA₂α sense ODN–treated cells. As expected, infiltration of monocytes (Fig. 8 B, middle) or total leukocytes (Fig. 8 B, bottom) remained unaffected in animals that received either untreated or cPLA₂α sense ODN–treated cells. These results demonstrate that iPLA₂β and cPLA₂α are required for MCP-1–induced chemotaxis of monocytes in vivo, thus validating our previous in vitro findings.

Our studies characterize the important relative contributions of cPLA₂α and iPLA₂β for regulating monocyte chemotaxis to MCP-1. Reduced migration of adoptively transferred, phospholipase-antisense ODN–treated mouse monocytes to the peritoneum in response to thioglycolate indicates the important signaling roles of iPLA₂β and cPLA₂α in monocyte chemotaxis to MCP-1 in vivo.

The intracellular locations of these phospholipases vary with the type of cell and nature of stimulus. For example, in INS-1 cells, iPLA₂β translocates to the perinuclear region during endoplasmic reticulum stress or stimulation with glucose (20, 21), whereas in smooth muscles cells, iPLA₂β is associated with the plasma membrane (22). Similarly, cPLA₂α has been reported to be associated with the nuclear membrane, Golgi complex, endoplasmic reticulum, plasma membrane, and an undefined intracellular compartment (23–25). Our observations of the significant increase in colocalization of iPLA₂β with membrane and pseudopod markers and of cPLA₂α with the ER marker in nonpolar but MCP-1–stimulated monocytes suggest that their recruitment to distinct intracellular locations is induced by MCP-1. This conclusion is further supported by the insignificant colocalization of iPLA₂β with cPLA₂α, and vice versa.

Earlier, we showed the restoration of chemotaxis of iPLA₂β-deficient monocytes to MCP-1 with LPA (14). LPA has been reported to induce phospholipase D (PLD) activity, cytoskeleton rearrangements, and assembly of focal adhesions (26). PLD can generate phosphatidic acid (PA) from membrane phospholipids (27). Interestingly, PA is a preferred substrate of iPLA₂β, yielding LPA (28).We have observed that PLD activity is induced by MCP-1 and required for monocyte chemotaxis (unpublished data). Furthermore, PLD is required for LPA-induced cytoskeletal rearrangements, likely through LPA formation (27). As schematically represented in Fig. 9, these observations tempt us to speculate that in response to MCP-1, PLD acts on membrane phospholipids to generate PA, and then iPLA₂β acts on PA to liberate LPA. LPA possibly interacts with components of pseudopod assembly and regulates cytoskeletal reorganization; hence, speed and directionality of monocytes. This conjecture is reinforced by observations that actin polymerization is spatially localized to the membrane-enriched pseudopod, where iPLA₂β is recruited in response to MCP-1 (Fig. 1, B and C), MCP-1 induces association of iPLA₂β with F-actin, irrespective of cell polarity (Fig. 1 D), and MCP-1–induced actin polymerization is significantly reduced in response to iPLA₂β inhibition or reduced expression (Fig. 6).

MCP-1 chemotaxis can be restored in cPLA₂α-deficient monocytes by AA (14). Thus, cPLA₂α may contribute to monocyte migration by generating AA in the endoplasmic reticulum (Fig. 9). AA can be converted into various metabolites, such as prostaglandins, lipoxins, LTB4, and H(P)ETEs via COX-2, cytochrome P450, or lipoxygenase pathways (29). Lipoxins induce RhoA- and Rac-mediated actin reorganization in monocytes/macrophages (30), whereas prostaglandin 2 increases migration of dendritic cells (31). Our observations, in view of these reports, call for identification of key metabolites of AA that regulate monocyte migration to MCP-1.

Directed cell migration is initiated by sensing of chemotactic stimuli, and culminates in the directed migration of cells. During this process, an external shallow gradient of chemokine is perceived, translated, and amplified into a stable intracellular gradient of signaling molecules that results in the formation of pseudopod at the front and uropod in the rear of a migrating cell. The pseudopod is formed by the activation of the heterotrimeric GTP binding protein Gi, whereas the uropod is formed by the activation of heterotrimeric GTP-binding protein G_12/13. Phosphatidylinositol (3,4,5) triphosphate (PtdIns[3,4,5]P₃), a key lipid messenger, is synthesized in the leading edge by PI(3)Ks and is degraded by PTEN in the rear edge of migrating cells (32). This vectorial metabolism gives rise to an intracellular gradient of PtdIns(3,4,5)P₃. Pseudopod is organized by Rac/PtdIns(3,4,5)P₃/actin and is stabilized by the scaffold Hem-1 (33, 34). In contrast, the uropod is organized by Rho-mediated, myosin-based contractions (35). This distinct intracellular hierarchy is thought to commence polarity and ensure directed migration of cells in a chemokine gradient. Multiple signaling molecules are believed to orchestrate these distinct events at the opposing edges of a migrating cell. For example, Cdc42 regulates stability and location of the leading edge, whereas Rac, which transforms random migration of fibroblasts and epithelial cells to directed migration (36), only controls morphological and invasion characteristics of macrophages (37). Vav1 is supposed to control the speed of migrating macrophages (38). In neutrophils, deficiency of Rho GEF Lsc causes faster migration and loss of directionality (39), whereas deficiency of PI3Kγ causes only loss of directionality (40). In Dictyostelium discoideum, PTEN regulates levels of PtdIns(3,4,5)P₃; in contrast, in neutrophils, it is metabolized by the PtdIns(3,4,5)P₃ phosphatase SHIP1 rather than PTEN. These observations not only point toward evolution of cell-based variability in key signaling molecules, they also provide an impetus to search for unknown members of these cascades (41, 42).

Once considered a probable cellular compass, recent reports suggest that PtdIns(3,4,5)P₃ is primarily a component of positive-feedback loop controlling persistent actin dynamics (43). Emerging concepts argue for the existence of parallel signaling pathways for chemotaxis and indicate that an unknown cellular compass exists upstream of PI3Kγ (42, 44). Interestingly, a calcium-independent phospholipase (PLA₂A) that liberates AA from phosphatidylcholine and acts in parallel with PI3K/PTEN has recently been implicated in chemotaxis of D. discoideum (45). Because iPLA₂ does not show a preference for the fatty acyl moiety at the sn-2 position of phospholipids, and D. discoideum does not contain cPLA₂α, it is possible that PLA₂A merely performs twin functions of cPLA₂α and iPLA₂β by liberating LPA and AA. Our observations on loss of directionality of iPLA₂β-deficient monocytes raise the possibility that iPLA₂β is either a cellular compass or a component of a cellular compass. Our observations, together with recent reports (44, 45), suggest that phospholipases are critical components of parallel signaling pathways regulating monocyte chemotaxis as schematically represented in Fig. 9. How iPLA₂β directs monocytes toward MCP-1, as well as whether it is a universal regulator of chemotaxis, remains to be deciphered. One key function ascribed to iPLA₂β is membrane remodeling and membrane trafficking that brings about changes in shape and fluidity of the plasma membrane (46). It will be interesting to explore whether iPLA₂β regulates monocyte chemotaxis by changing lipid composition of membrane rafts, thereby modifying assembly of signaling molecules in the pseudopod of migrating cells (47).

The impaired in vivo migration of adoptively transferred mouse monocytes that contain either inactive PLA₂ (by AACOCF₃) or were rendered deficient in PLA₂ by AS-ODN validates our in vitro observations (14) in a dynamic in vivo milieu. We now have adapted this in vivo assay to monitor human monocyte chemotaxis (48). Our reports show the importance of in vitro studies for identification of potential signaling components. Our in vivo MCP-1 chemotaxis assay provides a straightforward approach to verify the relevance of signaling pathways that appear promising for regulating MCP-1–dependent chemotaxis in vitro.

In summary, our studies provide insight into how two seemingly similar phospholipases regulate directed migration of monocytes from different intracellular sites. We report that MCP-1 induces chemotaxis of monocytes by increasing speed, as well as introducing directionality. By translocating to the pseudopod and regulating directionality and speed, iPLA₂β acts front and center, whereas in contrast, cPLA₂α is recruited to the endoplasmic reticulum and regulates only speed. The profound defects in migration of iPLA₂β- or cPLA₂α-deficient mouse monocytes in vivo validate the important roles for these enzymes in regulating extravasation of blood monocytes to sites of inflammation.

BEL and AACOCF₃ were obtained from Biomol. Bel enantiomers (R and S) were provided by R. Gross (Washington University School of Medicine, St. Louis, MO). Human recombinant MCP-1 (BD Biosciences) was used at a concentration of 5.75 nM. Anti-iPLA₂β (1:50; goat polyclonal), anti–human Cdc-42 (1:50; rabbit polyclonal), and anti–human PDI (1:100; rabbit polyclonal) were purchased from Santa Cruz Biotechnology. Anti–human cPLA₂α (rabbit polyclonal) and a mouse monoclonal antibody against human Na-K ATPase (1:200) were obtained from Cell Signaling Technology. Rhodamine phalloidin (1:40), Rab5, lectin GSII-Alexa Fluor 594, and Alexa Fluor 594– or 488–tagged species-specific secondary antibody raised in donkey (1:200) were obtained from Invitrogen.

Human monocytes were isolated from freshly drawn human whole blood, as described by McNally et al. (49), and maintained in DME (10% BCS) in polypropylene tubes at 2 × 10⁶ cells/ml. Studies on human monocytes were approved by the Institutional Review Board of the Cleveland Clinic.

This assay was performed as previously described (14) and was used to quantify monocyte migration to MCP-1.

The under-agarose cell migration assay was performed as described by Heit and Kubes (50) using 10% BCS containing DME. Net distance in this assay denotes the distance between the edge of the well and the final position of migrating cells. It was determined from the composite photomicrograph of the entire field of migration.

Monocytes were pretreated with Racemic-BEL or its R or S enantiomers (0.1–5 μM) or AACOCF₃ (50 μM) at 37°C in 10% CO₂ for 1 h, and washed in serum-free DME before being used for chemotaxis. The sense and antisense ODN sequences for phospholipases used in this study were phosphorothioate modified and purified by HPLC (Sigma-Aldrich). These ODNs were of proven efficacy and specificity, and were delivered under standardized experimental conditions, as described (14). These specific antisense ODNs were effective in reducing the expression of iPLA₂β or cPLA₂α, as described in our previous study (14).

Monocytes were allowed to migrate using the under-agarose cell migration assay (1 h), fixed, and permeabilized with Triton X-100 (0.1% in PBS for 4 min). Nonspecific sites were blocked with 3% BSA in PBS (37°C for 1 h). Cells were incubated with primary antibody (for 1 h at 37°C), washed, and visualized with fluorescently labeled secondary antibody (for 1 h at 37°C). Stained cells were mounted in Vectashield (Vector Laboratories) and observed using a laser-scanning confocal microscope (DMR XE attached to TCS SP2; Leica). Horizontal sections were imaged using a 63× objective (magnification 5×, pinhole 1, and format of 1,024 × 1,024). Images were analyzed by Image-Pro Plus (Media Cybernetics) and processed using Photoshop CS (Adobe).

For this purpose, under-agarose cell migration was performed in serum-coated polystyrene six-well culture dishes, essentially as described in the previous section. The plate was placed on a preheated stage at 37°C under 10% CO₂. Images were acquired with an inverted microscope (DMIRB; Leica) using a 10× phase objective. Images were acquired at 3-min intervals using 30-ms exposure times and recorded with a cooled charge-coupled device camera (Cool SNAP HQ; Roper Scientific) using Metamorph software (Universal Image).

Image analysis was conducted with Image-Pro Plus. In the absence of MCP-1, randomly migrating cells often return to the well where they originated. Because of the large number of cells in the well itself, returning cells could not be followed after reentering the well. Therefore, cells that can be tracked for the 60-min duration were used for migration analysis. Total distance depicts total length of a migratory path, speed denotes total distance/60 min, and net distance depicts the linear distance between the first and the last position of a migrating cell. Net-X is the net value of the X coordinate after 60 min. The turn angle is the absolute value of the angle between successive positions of migrating cell measured from the direction of the preceding position. Average angle/turn depicts the average angles of the turns taken by a migrating cell during 60 min of migration at 3-min intervals.

Mean fluorescent intensity of iPLA₂β in tail, midbody, and pseudopod of polar cells was determined by measuring pixel intensity of iPLA₂β and area clearly demarcated based on cellular morphology shown by differential interference contrast (DIC) images using Image-Pro Plus. Colocalization (percentage) of enzymes with marker proteins were determined as previously described (51).

This assay was used to determine the effect of iPLA₂β inhibition on F-actin polymerization, as described by Finkel et al. (52) using 2 million cells.

Animals (BALB/CJ; Jackson ImmunoResearch Laboratory) were used according to the protocol approved by the Cleveland Clinic Foundation Institutional Animal Care and Use Committee. BALB/CJ female mice (7–8 mo, two donors per recipient) were anesthetized with avertin (0.4 ml/animal). Blood was collected by cardiac puncture in a 1-ml syringe containing 0.05 ml of EDTA (0.5 M). Blood (10 ml) was diluted with 10 ml of Tricine-buffered saline (TBS; 10 mM, pH 7.4) and mixed with 5 ml of OptiPrep (Sigma-Aldrich). Blood was transferred to a suitable tube, overlayed with 0.5 ml of TBS and centrifuged at 1,000 g for 30 min at 20°C without the brake. The supernatant, containing mononuclear cells, was collected, diluted with 2 volumes of TBS, and centrifuged at 400 g for 10 min. The pellet was reconstituted in 0.5 ml of TBS, layered on dialyzed calf serum (2 ml), and centrifuged at 200 g for 15 min to remove platelets. Cells were suspended in TBS and counted. The recovery of mononuclear cells by this method was 7.45 × 10⁴/g mouse.

Cells were labeled per the manufacturer's instructions. Cells were suspended in DMEM and treated with inhibitors as previously described (14).

Recipient mice (3–4 per group) were lightly anesthetized with avertin. Peritoneal inflammation was induced by thioglycolate injection (1 ml, 4% in physiological saline) into the peritoneal cavity. Tail veins were dilated with limonene and cleaned with 95% ethanol. The labeled mononuclear cells (1.4–1.8 million per animal) were injected into the tail vein, and the time of injection was recorded. Injected cells represent only 11–14% of total peripheral blood mononuclear cells of similar weight recipient animals. After 24 h, peritoneal cells were harvested, washed, resuspended in PBS (1 ml), and counted. Cells were fixed and used to prepare 10 cytospins for each sample. Half of them were stained with HEMA per the manufacturer's (Thermo Fisher Scientific) instruction, to determine the number of monocytes. The remainder was mounted in Vectashield. PKH26-positive cells were counted on an upright microscope (model DMR; Leica) using a Texas red filter.

Statistical significance of observations was calculated using the Student's t tests. P < 0.05 was considered significant.

Fig. S1 presents data indicating that MCP-1 induces membrane localization of iPLA₂β in both polar and nonpolar monocytes. Fig. S2 demonstrates the transient nature of BEL inhibition of monocyte chemotaxis to MCP-1 in a time-dependent manner. Fig. S3 shows the stable inhibitory effect of AACOCF₃ on monocyte chemotaxis to MCP-1, and that AACOCF₃ does not affect monocyte viability under these conditions. Fig. S4 shows inhibition of cPLA₂α protein expression in murine monocytes/macrophages by treatment with antisense ODN. Supplemental materials and methods are also provided.

The authors thank Dr. Richard Gross for the gift of BEL enantiomers; Amit Vasanji for help with image analysis; and Drs. Josephine Adams and Ashish Bhattacharjee for their helpful comments.

This study is supported by National Institutes of Health grants (HL 510681 and HL 61971) to M.K. Cathcart and by the General Clinical Research Center Grant M01-RR-018390.

The authors have no competing financial interests.