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Vol. 16, Issue 8, 3800-3809, August 2005

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Cytosolic Phospholipase A₂- Mediates Endothelial Cell Proliferation and Is Inactivated by Association with the Golgi Apparatus

School of Biochemistry and Microbiology, University of Leeds, Leeds LS2 9JT, United Kingdom

Submitted February 25, 2005; Revised April 27, 2005; Accepted May 24, 2005
Monitoring Editor: Vivek Malhotra

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGMENTS REFERENCES

 
Arachidonic acid and its metabolites are implicated in regulating endothelial cell proliferation. Cytosolic phospholipase A₂- (cPLA₂) is responsible for receptor-mediated arachidonic acid evolution. We tested the hypothesis that cPLA₂ activity is linked to endothelial cell proliferation. The specific cPLA₂ inhibitor, pyrrolidine-1, inhibited umbilical vein endothelial cell (HUVEC) proliferation in a dose-dependent manner. Exogenous arachidonic acid addition reversed this inhibitory effect. Inhibition of sPLA₂ did not affect HUVEC proliferation. The levels of cPLA₂ did not differ between subconfluent and confluent cultures of cells. However, using fluorescence microscopy we observed a novel, confluence-dependent redistribution of cPLA₂ to the distal Golgi apparatus in HUVECs. Association of cPLA₂ with the Golgi was linked to the proliferative status of HUVECs. When associated with the Golgi apparatus, cPLA₂ activity was seen to be 87% inhibited. Relocation of cPLA₂ to the cytoplasm and nucleus, and cPLA₂ enzyme activity were required for cell cycle entry upon mechanical wounding of confluent monolayers. Thus, cPLA₂ activity and function in controlling endothelial cell proliferation is regulated by reversible association with the Golgi apparatus.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGMENTS REFERENCES

 
Endothelial cells line the luminal surface of all blood vessels and are essential for vascular processes including blood clotting, blood pressure, and blood flow (Vane et al., 1990). New blood vessel formation mediated by endothelial cells (angiogenesis) is also a crucial step in wound healing and the establishment of tumors (Folkman and Shing, 1992; Carmeliet, 2000). Antiangiogenic therapy is seen as one of the most promising strategies to restrict tumor growth and prevent metastases (Brem et al., 1993). During the process of angiogenesis endothelial cells migrate, proliferate, and differentiate to form new capillaries (Carmeliet, 2000). Numerous reports have implicated arachidonic acid (AA) in the control of proliferation in a variety of cell types. Actively proliferating endothelial cells in culture liberate elevated levels of AA and its metabolites (Evans et al., 1984; Whatley et al., 1994) and stimulation of endothelial cells with the potent mitogen, basic fibroblast growth factor, induces AA release (Fafeur et al., 1991; Sa et al., 1995). Eicosanoids, the bioactive metabolites of AA, also play a major role in the proliferation and migration of endothelial cells (Dethlefsen et al., 1994; Nie et al., 2000; Dormond et al., 2001; Antoniotti et al., 2003).

Cytosolic phospholipase A₂ (cPLA₂) is an 85-kDa, Ca²⁺-sensitive member of the phospholipase A₂ family of enzymes (Clark et al., 1995) that includes the calcium-independent (iPLA₂) and secretory (sPLA₂) phospholipases A₂. These are responsible for hydrolysis of the sn-2 fatty-acyl bond of phospholipids to simultaneously generate free fatty acid and lysophospholipid (Dennis, 1997). There are at least four cPLA₂ paralogs (cPLA₂, cPLA₂, cPLA₂, and cPLA₂), of which cPLA₂ is most characterized. On cell stimulation and intracellular Ca²⁺ elevation, cPLA₂ translocates from the cytosol to intracellular membrane substrates utilizing an N-terminal Ca²⁺-dependent lipid binding (CalB) domain (Channon and Leslie, 1990; Nalefski et al., 1994; Schievella et al., 1995; Bittova et al., 1999; Evans et al., 2001). cPLA₂ preferentially cleaves phospholipids containing AA at the sn-2 position (Dennis, 1997) and is thus considered the key enzyme responsible for receptor-mediated AA liberation and subsequent eicosanoid synthesis (Kramer and Sharp, 1997). As such, it is a pivotal enzyme in AA-mediated cell proliferation and migration. At confluence endothelial cells undergo contact inhibition of growth, cease proliferating, and enter the G₀ phase of quiescence. Correspondingly there is a significant decline in AA output and eicosanoid synthesis (Evans et al., 1984; Whatley et al., 1994), which has been attributed to the reduced cPLA₂ activity (Whatley et al., 1994). Because of the importance of endothelial cell proliferation in human retinopathies and cancers, there is great emphasis on developing therapies that modulate endothelial cell growth. Despite this, the exact mechanisms by which cPLA₂ activity is differentially regulated in nonconfluent and confluent endothelial cells and the role of cPLA₂ in endothelial cell proliferation remain unclear.

In this study we demonstrate that cPLA₂ activity is required for endothelial cell proliferation and cell cycle control. We also describe a novel redistribution of cPLA₂ to the Golgi apparatus of human umbilical vein endothelial cells (HUVECs) in a confluence-dependent manner. The subcellular localization of cPLA₂ is closely linked to the proliferative status of HUVECs. When associated with the Golgi apparatus, cPLA₂ activity is 87% inhibited. After mechanical wounding, cPLA₂ redistribution is linked to the induction of Ki67 expression and entry into the cell cycle. These observations shed light on the mechanisms by which cPLA₂ localization, activity, and function are modulated in quiescent and proliferating endothelial cells.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGMENTS REFERENCES

 
Materials
Alexafluor 594–conjugated concanavalin A (ConA) and secondary anti-goat, anti-mouse, and anti-rabbit Alexafluor 488– or 594–conjugated antibodies were purchased from Molecular Probes (Eugene, OR). Rhodamine-conjugated wheat germ agglutinin (WGA) was purchased from Sigma (Poole, United Kingdom). Affinity-purified goat polyclonal anti-cPLA₂ antibodies were purchased from Santa Cruz Biotechnology (sc-1724, Santa Cruz, CA). Rabbit anti-GM130 antibodies were from M. Lowe (University of Manchester, United Kingdom). Mouse monoclonal anti-ERGIC-53 and -1,4-galactosyltransferase (GalT) antibodies were provided by H. P. Hauri (Basel, Switzerland) and T. Suganuma (Miyazaki, Japan), respectively. Rabbit polyclonal anti-TGN46 was supplied by S. Ponnambalam (University of Leeds, United Kingdom). Rabbit polyclonal anti-mannosidase II (ManII) antibodies were purchased from Serotec Ltd (Oxford, United Kingdom). HRP-conjugated anti-goat secondary antibodies were purchased from Pierce (Tattenhall, United Kingdom). 2-(p-amylcinnamoyl)amino-4-chlorobenzoic acid (ONO-RS-082) was purchased from BioMol (Plymouth Meeting, PA). Pyrrolidine-1 was supplied by M. H. Gelb (University of Washington, Seattle,WA). All other reagents were obtained from Sigma or Life Technologies (Paisley, United Kingdom) unless otherwise stated.

Cell Culture
HUVECs were isolated from human umbilical cords as previously described (Jaffe, 1984; Howell et al., 2004). Cells were cultured in endothelial cell basal medium supplemented with human recombinant epidermal growth factor (EGF; 5 ng/ml), hydrocortisone (0.2 µg/ml), vascular endothelial growth factor (VEGF, 0.5 ng/ml), human recombinant basic fibroblast growth factor (bFGF, 10 ng/ml), recombinant long R3 insulin-like growth factor-1 (IGF-1, 20 ng/ml), ascorbic acid (1 µg/ml), heparin (22.5 µg/ml), amphotericin B (50 ng/ml), gentamicin (50 µg/ml), and 2% (vol/vol) fetal calf serum (PromoCell, Heidelberg, Germany). All HUVEC cultureware was coated with 0.1% (wt/vol) pigskin gelatin unless stated otherwise. HUVECs were never used in excess of three passages and expressed the characteristic endothelial markers von Willebrand factor and platelet-endothelial cell adhesion molecule-1. All cells were grown at 37°C in a humid incubator containing 5% (vol/vol) CO₂.

Cell Proliferation ELISA
HUVEC proliferation rates in the presence or absence of varying concentrations of ONO-RS-082 or pyrrolidine-1 were compared using a 5-bromo-2'-deoxyuridine (BrdU) incorporation-based ELISA (Roche Diagnostics, Lewes, East Sussex, United Kingdom). Cells were seeded at 1 x 10³ cells per well (0.55 x 10³ cells/cm²) in 96-well plates and cells, grown for 24 h, and then processed according to manufacturer's instructions. The BrdU incorporation period was fixed at 16 h.

SDS-PAGE and Immunoblotting
Samples (20 or 10 µg protein) were resolved for 60 min at 30 mA/gel on 10% SDS-PAGE mini-gels using a discontinuous buffer system (Laemmli, 1970). For immunoblotting, protein was transferred onto nitrocellulose membranes for 3 h at 300 mA (Towbin et al., 1979). Membranes were blocked in 5% (wt/vol) nonfat milk in phosphate-buffered saline (PBS) for 30 min and then incubated overnight with primary antibody (1:500) at room temperature. After incubation with HRP-conjugated anti-goat IgG (1:3000) for 1 h, immunoreactive bands were visualized using a West Pico–enhanced chemiluminescence (ECL) detection kit (Pierce, Rockford, IL). A Fuji Film Intelligent dark box II image reader using Fuji Las-1000 Prosoftware (Tokyo, Japan) was used to capture images. Band intensities were determined densitometrically using Aida (Advanced Image Data Analyzer, Raytest, Straubenhardt, Germany) 2.11 software. For comparison of nonconfluent and confluent HUVEC protein expression, cells were grown on noncoated plastic to control for misleading protein estimation due to gelatin-coated surfaces.

Immunofluorescence Microscopy
This technique was adapted from previous protocols (Heggeness et al., 1977; Barwise and Walker, 1996). Cells were seeded on 0.1% (wt/vol) gelatin-coated coverslips and grown to the required level of confluence. Medium was aspirated and cells were fixed in 10% (vol/vol) Formalin (in neutral buffered saline; Sigma) for 5 min at 37°C. Before fixation some cells were stimulated for 1 min with 5 µM A23187 [GenBank] as previously described (Grewal et al., 2003, 2004). All ensuing steps were performed at 25°C. After permeabilization with 0.1% (vol/vol) Triton X-100 in PBS for 5 min, cells were refixed for 5 min, washed three times with PBS, and then incubated in 50 mM ammonium chloride in PBS for 10 min. After three further PBS washes, nonspecific binding sites were blocked with 5% (vol/vol) donkey serum in PBS for 3 h. Cells were incubated overnight with primary antibody followed by the appropriate Alexafluor-conjugated secondary antibodies, Alexafluor 594–conjugated ConA or rhodamine-conjugated WGA for 3 h. For antigenic adsorption, antibodies to cPLA₂ were incubated with blocking peptide (1:5 ratio of µg antibody to µg peptide) for 30 min before incubation, as previously described (Grewal et al., 2003). Finally cells were washed eight times in PBS and mounted on microscope slides in prolong mounting medium (Molecular Probes).

Deconvolution Imaging and Quantitation
Deconvolution fluorescence microscopy was performed using an Olympus IX-70 inverted fluorescence microscope and DeltaVision deconvolution system (Applied Precision, Issaquah, WA). Individual optical sections of 0.2 µm were generated from 15 iterative cycles of deconvolution. Quantification of colocalization was determined using the IMARIS software suite (Bitplane AG, Zurich, Switzerland) on selected Golgi regions. Background was eliminated by excluding gray scale values lower than 10% of the maximum pixel intensity. Colocalized pixels were recorded and expressed as percentages of the total pixels selected. Mean pixel intensities were determined using Image J software (http://rsb.info.nih.gov/ij/). Regions of interest were selected, mean pixel intensity calculated then mean background pixel intensity subtracted from this. No images analyzed were pixel saturated.

Brefeldin A Treatment
HUVECs were grown to confluence on 0.1% (wt/vol) gelatin-coated coverslips. Cells were washed twice with prewarmed (37°C) PBS and then incubated with 5 µg/ml brefeldin A (BFA; Roche Diagnostics, Sussex, United Kingdom) in serum-free media for 30 min and processed for immunofluorescence microscopy as described above.

AA Release
This technique was adapted from previous protocols (Whatley et al., 1994; Bailleux et al., 2004). HUVECs were cultured to the required cell density in six-well culture dishes and labeled for 24 h with 1 µCi/ml [³H]AA. Cells were washed three times with PBS and then incubated with 10 µM bromoenol lactone for 30 min to inhibit iPLA₂ activity. Medium was then aspirated and cells were incubated with serum-free medium supplemented with 5 µM A23187 [GenBank] and 0.3% (wt/vol) fatty acid–free bovine serum albumin. Aliquots of media were removed at times indicated, centrifuged at 16,000 x g for 5 min, and supernatant was counted for radioactivity by liquid scintillation. Cells were lysed in 0.1% Triton X-100 for 5 min and also counted by liquid scintillation for radioactivity.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGMENTS REFERENCES

 
cPLA₂ Mediates Endothelial Cell Proliferation
cPLA₂ activity but not sPLA₂ activity was essential for maximal HUVEC proliferation (Figure 1). The contribution of cPLA₂ to HUVEC proliferation was assessed by determining BrdU incorporation in the presence of varying concentrations of pyrrolidine-1 (Figure 1A). Pyrrolidine-1 specifically inhibits cPLA₂ and has no effect on iPLA₂ or sPLA₂ (type IIA, X and V) activity (Ghomashchi et al., 2001). Pyrrolidine-1 acted in a dose-dependent manner, inhibiting proliferation by 28.7 ± 4.6, 49.9 ± 4.2, and 70.7 ± 2.9% at concentrations of 5, 10, and 20 µM (Figure 1A). At concentrations above 20 µM, pyrrolidine-1 was toxic, whereas at concentrations of 10 µM and below, cell viability was unaffected, as determined by trypan blue exclusion. Varying concentrations of the sPLA₂ inhibitor, ONO-RS-082 (Hashimoto et al., 2003), had no significant effect on BrdU incorporation (Figure 1B). The inhibitory effects of pyrrolidine-1 could be significantly reversed by addition of exogenous free AA (Figure 1C). AA, 10 µM, reversed the effect of 10 µM pyrrolidine-1, reaching 75.1 ± 2% of control. We decided to investigate further the properties of cPLA₂ under different growth conditions.

View larger version (20K): [in this window] [in a new window]   Figure 1. Inhibition of HUVEC proliferation by cPLA2 inhibition. Quantitation of HUVEC growth in the presence of varying concentrations of pyrrolidine-1 (A) and ONO-RS-082 (B) for 16 h (n = 6, ±SEM). Quantitation of HUVEC growth in the presence of 10 µM pyrrolidine-1 with and without 10 µM arachidonic acid supplementation (C; n = 5, ±SEM). Proliferation was determined using a colorimetric ELISA based on BrdU incorporation, as described in Materials and Methods. * p < 0.02 versus control. ** p < 0.001 versus control. p < 0.001 versus 10 µM pyrrolidine-1 without AA.

cPLA₂ Expression and Localization in Confluent and Nonconfluent HUVECs
The lower levels of AA release seen with confluent endothelial cells are not due to reduced cPLA₂ expression (Figure 2, A and B). It is well established that nonconfluent endothelial cells release much greater levels of AA and eicosanoids than confluent endothelial cells and that the mechanical wounding of confluent monolayers results in elevated AA production (Evans et al., 1984; Whatley et al., 1994). Lower levels of AA release could be due to reduced cPLA₂ expression because it is the rate-limiting enzyme in AA liberation. The protein levels of cPLA₂ were determined in subconfluent proliferating and in confluent nonproliferating HUVECs (Figure 2, A and B). cPLA₂ levels were assessed by Western blotting of HUVEC total protein using a well-characterized affinity-purified antibody specific to the C-terminal region of cPLA₂ (Grewal et al., 2002, 2003, 2004). Densitometric analysis demonstrated no significant difference in total cPLA₂ levels between subconfluent and confluent cells when using 10 µg (Figure 2B) and 20 µg (unpublished data) of total protein. The amounts of total cell protein used for Western blot detection of cPLA₂ gave responses that reside in the linear region of a saturation curve. Antigenic absorption with a blocking peptide, corresponding to the C-terminal 20 amino acids unique to cPLA₂, eliminated detection by immunoblotting (unpublished data).

View larger version (46K): [in this window] [in a new window]   Figure 2. cPLA2 expression and localization in nonconfluent and confluent HUVECs. cPLA2 was detected in nonconfluent and confluent HUVEC total protein (10 µg) by immunoblotting using a goat polyclonal antibody (A). Relative amounts of cPLA2 immunoreactivity were determined using Aida densitometry software and plotted (±SEM) (B). The localization of cPLA2 in nonconfluent (C) and confluent cells (D) was determined by immunofluorescence microscopy. HUVECs were grown to nonconfluence or confluence on 0.1% gelatin coated coverslips, fixed, permeabilized, and cPLA2 detected with a goat antibody. In E, the percentage of total cells with juxta-nuclear cPLA2 localization was quantified for confluent, subconfluent, and nonconfluent cells (n = 3, ±SEM). The localization of cPLA2 in nonconfluent cells upon A23187 [GenBank] stimulation was determined by immunofluorescence microscopy (F). HUVECs were grown to nonconfluence or confluence on 0.1% gelatin-coated coverslips, fixed, and permeabilized, and cPLA2 was detected with a goat antibody. All images represent 0.2-µm sections through cell nuclei. Scale bar, 25 µm. * p < 0.001 versus confluent cells. All results are representative of at least 3 separate experiments.

cPLA₂ redistributes to a compact juxtanuclear reticulum in confluent HUVECs (Figure 2, C–E). Indirect immunofluorescence microscopy was used to determine the subcellular localization of cPLA₂ in HUVECs exhibiting differing levels of confluence (Figure 2, C and D). As previously described for endothelial cells (Sierra-Honigmann et al., 1996; Grewal et al., 2002), nonconfluent HUVECs displayed homogeneous cPLA₂ staining throughout the cytoplasm and nucleus (Figure 2C). High-resolution deconvolution microscopy shows that cPLA₂ was present both as diffuse and structured pools of staining in nonconfluent cells, similar to observations with fibroblasts (Bunt et al., 1997). At confluence, cPLA₂ was redistributed to a compact juxtanuclear reticulum (Figure 2D). HUVECs of differing levels of confluence were scored for the presence of juxtanuclear cPLA₂ staining (Figure 2E). Our criteria defined confluent HUVECs as those completely surrounded by other cells. Cells not contacting any other were categorized as nonconfluent. Cells contacting one or more cells but not entirely surrounded were classed as subconfluent. A distinct correlation between confluence and the presence of juxtanuclear cPLA₂ staining was evident. A high percentage of confluent cells (87.5 ± 3.2%) displayed juxtanuclear cPLA₂ staining, whereas only 37.8 ± 4.7% of subconfluent and 6.2 ± 3.5% of nonconfluent cells displayed this localization pattern.

On stimulation of nonconfluent cells with the Ca²⁺ ionophore, A23187 [GenBank] , cPLA₂ translocates to the peri-nuclear membrane and endoplasmic reticulum (Figure 2F). These regions are classical sites of cPLA₂-mediated AA release in response to Ca²⁺ elevation (Hirabayashi et al., 1999). Stimulation of confluent endothelial cells with A23187 [GenBank] did not induce cPLA₂ translocation and cPLA₂ staining remained restricted to the juxtanuclear region (unpublished data). Antigenic absorption with a blocking peptide, corresponding to the C-terminal 20 amino acids unique to cPLA₂, eliminated detection by immunofluorescence microscopy (unpublished data).

cPLA₂ Redistributes to Membrane Components of the Distal Golgi Apparatus at HUVEC Confluence
cPLA₂ redistributes to the Golgi apparatus in confluent endothelial cells under standard cell culture conditions (Figure 3, A and B). Because the juxtanuclear staining pattern for cPLA₂ resembled the location of the Golgi apparatus, possible colocalization was investigated by counterstaining confluent HUVECs with rhodamine-conjugated WGA (Figure 3A). This lectin specifically binds N-acetyl--D-glucosaminyl residues found predominantly in the Golgi apparatus, with lower levels at the plasma and nuclear membranes (Virtanen et al., 1980). cPLA₂ was seen to colocalize extensively with WGA-positive juxtanuclear structures corresponding to the Golgi apparatus (yellow indicates overlap), but no significant overlap was seen with the endoplasmic reticulum (ER) marker lectin, ConA (Virtanen et al., 1980; Figure 3B). A smaller pool of cPLA₂ staining that did not colocalize with WGA-positive or ConA-positive structures was also evident (Figure 3, A and B).

View larger version (99K): [in this window] [in a new window]   Figure 3. Association of cPLA2 with membrane components of the Golgi apparatus. HUVECs were grown to confluence on 0.1% gelatin-coated coverslips, fixed, and then permeabilized, and cPLA2 was detected with a goat antibody (A and B). Cells were then incubated with Alexafluor 488–conjugated anti-goat antibodies and either rhodamine-conjugated WGA (A) or Alexafluor 594–conjugated ConA (B). Images represent 0.2-µm sections through cell nuclei. Disruption of the Golgi apparatus with brefeldin A (BFA; C and D). Confluent HUVECs were grown on 0.1% gelatin-coated coverslips and directly fixed (C) or treated for 30 min with 5 µg/ml the fungal metabolite BFA (D) before being fixed and permeabilized. Cells were then incubated with goat anti-cPLA2 and rabbit anti-ManII antibodies. Then cells were incubated with Alexfluor 488–conjugated anti-goat and Alexfluor 594–conjugated anti-rabbit antibodies. Images represent projected stacks of whole cells. Scale bar, 10 µm. All results are representative of three separate experiments.

In confluent HUVECs, cPLA₂ colocalized with components of the distal Golgi apparatus, i.e., the medial-Golgi, trans-Golgi, and trans-Golgi network (Figure 4). The Golgi complex consists of five spatially distinct subcompartments: the ER-Golgi intermediate compartment (ERGIC), cis-Golgi, medial-Golgi, trans-Golgi, and TGN. Antibodies specific to subcompartment resident proteins were used in costaining microscopy experiments to determine more specifically the localization of cPLA₂ in confluent HUVECs (Figure 4). ERGIC-53 (Figure 4A), GM130 (Figure 4B), ManII (Figure 4C), -1,4-galactosyltransferase (GalT; Figure 4D) and TGN46 (Figure 4E) are specific markers for the ERGIC (Schweizer et al., 1988, 1990), cis-Golgi (Nakamura et al., 1995), medial-Golgi (Novikoff et al., 1983), trans-Golgi (Roth et al., 1985; Nilsson et al., 1993), and TGN (Prescott et al., 1997), respectively. cPLA₂ appeared to colocalize most extensively with ManII- and GalT-positive structures (yellow indicates overlap; Figure 4, C and D). Extensive overlap was also seen with TGN46 (Figure 4E) but not to the extent of ManII and GalT. cPLA₂ only appeared closely associated and not superimposed with ERGIC-53 (Figure 4A) and GM130 (Figure 4B). Quantification of the colocalization of cPLA₂ with Golgi subcompartment markers was determined (Figure 4F). Percentage colocalization of cPLA₂ with ManII, GalT, and TGN-46 was seen to be 68.4 ± 3.5, 59.7 ± 3.8, and 42.2 ± 4.5%, respectively, whereas ERGIC-53 and GM130 displayed only 19.8 ± 5.6 and 23.8 ± 3.1% colocalization. All secondary antibody controls gave no staining (unpublished data).

View larger version (47K): [in this window] [in a new window]   Figure 4. Colocalization of cPLA2 with Golgi compartment markers. Confluent HUVECs were grown on 0.1% gelatin-coated coverslips, fixed, and permeabilized. Cells were incubated with goat anti-cPLA2 antibody and either mouse anti-ERGIC-53 (A), rabbit anti-GM130 (B), rabbit anti-ManII (C), mouse anti-GalT (D), or rabbit anti-TGN46 (E) antibodies. Cells were then incubated with anti-goat Alexafluor 488–conjugated and either anti-rabbit or anti-mouse Alexafluor 594–conjugated antibodies. Images represent 0.2-µm sections through cell nuclei. Scale bar, 10 µm. Results are representative of three separate experiments. Quantification of cPLA2 overlap with ERGIC-53, GM130, ManII, GalT, and TGN46 (F). Percentage colocalization was calculated using the IMARIS computer package as described in Materials and Methods (n = 9, ±SEM). * p < 0.02 versus GM130. ** p <0.001 versus GM130.

The Subcellular Localization of cPLA₂ Is Associated with HUVEC Proliferation
HUVEC confluence and proliferation are directly linked (Figure 5A). Subconfluent cultures of endothelial cells proliferate until they reach confluence then cell-cell contacts inhibit cellular proliferation (Chen et al., 2000). HUVECs were seeded at equivalent densities (0.55 x 10³/cm²) and cultured for 7 d. From days 2–7, cell numbers and proliferation per cell were determined and compared (Figure 5A). Cell density rose exponentially to reach a plateau after 5 d. BrdU incorporation per cell declined in correlation to also plateau at 5 d. At this point a confluent HUVEC monolayer is formed, cell-cell contacts inhibit proliferation, cells enter the G₀ phase of quiescence, and cell numbers cease to increase.

View larger version (20K): [in this window] [in a new window]   Figure 5. The subcellular localization of cPLA2 is associated with HUVEC proliferation. HUVEC cell density and growth were quantified over a culture period of 7 d (A). HUVECs (0.55 x 103/cm2) were seeded in 0.1% gelatin-coated 24-well culture dishes. They were counted after trypsinization each day from day 2 to day 7 after seeding (logarithmic scale, solid line with diamonds). Results represent the number of cells per cm2 (n = 3, ±SEM). HUVEC proliferation was determined using a colorimetric ELISA based on BrdU incorporation, as described in Materials and Methods (dotted line with squares). HUVECs (0.55 x 103/cm2) were seeded in 0.1% gelatin-coated 96-well culture dishes and grown for varying periods of time. Cells were labeled for 16 h with BrdU before processing. Results represent the BrdU incorporation per cell (n = 6, ±SEM). The subcellular localization of cPLA2 was quantified over a culture period of 7 d and compared with the BrdU incorporation per cell (B). HUVECs were seeded in 0.1% gelatin-coated coverslips at an equivalent density to those seeded in 96-well culture dishes. From day 2 to day 7 they were fixed, permeabilized, and processed for immunofluorescence microscopy and the localization of cPLA2 determined (solid line with diamonds). Results represent the percentage of total cells with juxta-nuclear localized cPLA2 at each time point (n = 3, ±SEM). Results were plotted alongside the quantification of BrdU incorporation per cell from Figure 5A. Arachidonic acid release in response to A23187 [GenBank] stimulation was determined for confluent (dotted line) and nonconfluent (solid line) HUVECs (C). Cells were loaded with 1 µCi/ml [3H]AA, stimulated with 5 µM A23187 [GenBank] , and released AA was determined at the times indicated (n = 3, ±SEM). Results are expressed as a percentage of the total [3H]AA incorporated. * p < 0.01 versus confluent.

It was important to determine whether association of cPLA₂ with the Golgi apparatus affected its enzyme activity. Consequently, nonconfluent, proliferating, and confluent nonproliferating cultures of cells were assayed for AA release in response to elevation of cytosolic Ca²⁺ (Figure 5C). cPLA₂ activity was reduced by 87% in confluent endothelial cells (Figure 5C); therefore, Golgi associated cPLA₂ was inactivated.

Mechanical Wounding Induces cPLA₂ Relocation in HUVECs
On mechanical wounding of confluent HUVEC monolayers, cPLA₂ reversibly redistributes from the Golgi apparatus to the diffuse cytoplasmic and nuclear location of nonconfluent HUVECs (Figure 6A–D). Mechanical wounding of confluent endothelial monolayers induces exit from G₀, entry into the cell cycle, proliferation, and migration of cells to fill denuded areas (Chen et al., 2000). HUVECs were mechanically wounded to investigate the effects on Golgi-localized cPLA₂ upon conversion from a confluent, quiescent to a subconfluent, proliferating state. Cells were fixed at various time points after initial wounding and cPLA₂ was detected by immunofluorescence microscopy (Figure 6A–C). Areas to the right side of the panels depict wound borders at the edge of denuded zones. At 0 h after wounding, HUVECs retained Golgi localized cPLA₂ (Figure 6A). However, 8 h (Figure 6B) and 24 h (Figure 6C) after wounding, HUVECs at the wound border displayed progressively less Golgi-associated cPLA₂. Cells invading the wound revert to the diffuse nuclear and cytoplasmic localization characteristic of nonconfluent cells. From 2 h after wounding, the percentage of HUVECs at wound borders displaying Golgi-localized cPLA₂ was significantly reduced (Figure 6D). After 24 h, only 11.4 ± 1.7% of cells at the wound border retained Golgi-localized cPLA₂. The Golgi apparatus remained intact (ManII immunofluorescence staining) throughout the experiment. Association of cPLA₂ with the Golgi apparatus is thus reversible upon conversion from confluent to subconfluent cells.

View larger version (87K): [in this window] [in a new window]   Figure 6. The Golgi association of cPLA2 is reversible upon wounding of confluent HUVEC monolayers. HUVECs were grown to confluence on 0.1% gelatin-coated coverslips and subjected to mechanical wounding. HUVECs were allowed to recover for varying periods of time before fixation. The localization of cPLA2 in cells at the wound border (right of panels) after 0 h (A), 8 h (B), and 24 h (C) of recovery was determined by immunofluorescence microscopy using a goat polyclonal antibody. Images represent projections of whole cells; scale bar, 100 µm. Results are representative of three separate experiments. In D, the percentage of cells at the wound border with Golgi-localized cPLA2 were quantified after 0, 1, 2, 4, 8, and 24 h of recovery (n = 3, ±SEM). * p < 0.05 versus 0 h, ** p < 0.001 versus 0 h.

cPLA₂ Activity Is Required for Induction of Ki67 Expression and Entry Into the Cell Cycle

View larger version (70K): [in this window] [in a new window]   Figure 7. Ki67 upregulation induced by mechanical wounding is associated with cPLA2 relocation. Confluent HUVECs were grown on 0.1% gelatin-coated coverslips and either directly fixed (A) or mechanically wounded and allowed to recover for 24 h before fixing (B). Cells were permeabilized and incubated with goat anti-cPLA2 antibody and rabbit anti-Ki67 antibodies. Cells were then incubated with anti-goat Alexafluor 488–conjugated and either anti-rabbit Alexafluor 594–conjugated antibodies. Images represent a confluent monolayer (A) or wound border (B) and are projections of whole cells. Scale bar, 100 µm. In C, the percentage of total cells with nucleoli localized Ki67 was quantified in confluent monolayers of cells and at wound borders after mechanical wounding and 24 h of recovery (n = 3; ±SEM). In D, the subcellular localization of cPLA2 in cells at the wound border with nucleoli localized Ki67 was determined. * p < 0.001 versus confluent monolayer. p < 0.001 versus Golgi. All results are representative of at least three separate experiments

cPLA₂ activity is vital for exit from G₀ and entry into the S phase of the cell cycle as pyrrolidine-1 inhibits Ki67 induction upon wounding (Figure 8). Confluent monolayers of HUVECs were mechanically wounded and recovered for 24 h in the absence or presence of 10 µM pyrrolidine-1. Cells that underwent cPLA₂ relocation were analyzed by immunofluorescence microscopy to assess the level of cPLA₂ and Ki67 expression (Figure 8, A and B). Levels of cPLA₂ expression do not significantly change (Figure 8C) but a 48.2% (±4.7%) reduction in Ki67 expression was observed after recovery in pyrrolidine-1 (Figure 8D). Ki67 expression is essential for cell cycle progression and proliferation. Thus, induction of HUVEC proliferation requires cPLA₂ activity and occurs upon relocation of cPLA₂ to the cytoplasm and nucleus.

View larger version (49K): [in this window] [in a new window]   Figure 8. cPLA2 inhibition prevents Ki67 induction in response to mechanical wounding. Confluent HUVECs were grown on 0.1% gelatin-coated coverslips, mechanically wounded, and recovered for 24 h in the absence (A) and presence (B) of 10 µM pyrrolidine-1. Cells were fixed, permeabilized, and incubated with goat anti-cPLA2 antibody and rabbit anti-Ki67 antibodies. Cells were then incubated with anti-goat Alexafluor 488–conjugated and either anti-rabbit or anti-mouse Alexafluor 594–conjugated antibodies. Only cells displaying non-Golgi localized cPLA2 at wound borders were imaged. Images represent 0.2-µm sections through cell nuclei. Scale bar, 25 µm. Mean cellular cPLA2 pixel intensity (C) and nuclear Ki67 pixel intensity (D) were determined using Image J software as described in Materials and Methods. Results represent cells displaying non-Golgi localized cPLA2 at wound borders for those recovered with and without 10 µM pyrrolidine-1 (n = 20). * p < 0.001 versus control. All results are representative of at least three separate experiments.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGMENTS REFERENCES

Our report is the first to document the importance of cPLA₂ activity for controlling endothelial cell proliferation and entry into the cell cycle. Previously AACOCF₃, an inhibitor of both cPLA₂ and iPLA₂ (Ackermann et al., 1995; Street et al., 1993) was seen to inhibit bovine aortic endothelial cell proliferation (Antoniotti et al., 2003) but that study lacked specific inhibitor studies. Here we use the specific cPLA₂ inhibitor, pyrrolidine-1 (Ghomashchi et al., 2001), to directly implicate cPLA₂ in endothelial cell proliferation. Reduced AA liberation by cPLA₂ is at least partially responsible for the inhibitory effects of pyrrolidine-1. Despite this, exogenous AA does not augment the proliferation of nonconfluent HUVECs (unpublished data) although they may proliferate at an optimal rate in our culture conditions so exogenous AA has no effect. In addition, exogenous AA does not induce entry into the cell cycle and proliferation of confluent endothelial cells (unpublished data). This demonstrates that although cPLA₂ activity is essential for cell cycle entry, it is insufficient by itself to override the contact-inhibition of proliferation. Our report suggests that inhibition of cPLA₂ could be a viable antiangiogenic route because the induction of cell cycle entry and proliferation in resting endothelial cells is crucial to angiogenesis (Carmeliet, 2000) and cPLA₂ activity is essential to this process.

We describe the novel confluence-dependent association of cPLA₂ with membrane components of the distal Golgi apparatus. When associated with the Golgi apparatus, cPLA₂ activity is inhibited by 87%. This is in contrast to previous studies that demonstrate increased cPLA₂ activity upon association with the Golgi apparatus (Pettus et al., 2004; Grimmer et al., 2005). Association with the Golgi apparatus and cPLA₂ inactivation may occur to differing extents depending on cell type, cell confluence, or the ability of cells to undergo contact inhibition of proliferation. We propose that Golgi association and inactivation of cPLA₂ is a feature of the contact-inhibition of endothelial cells.

Our results suggest that the Golgi association and inactivation of cPLA₂ is important to the regulation of its function in endothelial cells. cPLA₂ associates with the Golgi apparatus at HUVEC confluence when proliferation is inhibited and cells exit the cell cycle. Entry back into the cell cycle occurs upon relocation of cPLA₂ back to the diffuse cytoplasmic and nuclear location seen in subconfluent HUVECs. Thus, cPLA₂ is mediating HUVEC proliferation when it is found throughout the cytoplasm and nucleus so the localization of cPLA₂ in proliferating endothelial cells is essential for targeting it to areas where it functions to aid proliferation. AA is evolved in response to Ca²⁺ elevation and cPLA₂ relocation to membrane substrate (Hirabayashi et al., 1999) as occurs in response to growth factor stimulation. In nonconfluent endothelial cells in response to Ca²⁺ elevation, cPLA₂ associates with perinuclear membrane substrate, a known site of cPLA₂-derived AA production (Hirabayashi et al., 1999). This does not occur in confluent endothelial cells because cPLA₂ remains immobilized on the Golgi apparatus and cPLA₂ activity remains inhibited. Thus, we outline a novel mechanism for the inhibition of cPLA₂ activity. We propose that association with the Golgi apparatus is a mechanism to partition cPLA₂ from sites of action in quiescent endothelial cells, excluding it from substrate. This model is also consistent with the regulation of MEK/ERK by Sef, a molecular scaffold that resides in the Golgi complex. Binding to Sef allows MEK/ERK signaling to cytosolic substrates but excludes nuclear targets (Torii et al., 2004).

We are the first to document the confluence-dependent relocation of cPLA₂ in any cell type. Association of cPLA₂ with distal components of the Golgi apparatus has been documented previously in epithelial cell lines but only in response to agents that elevate intracellular free Ca²⁺ (Evans et al., 2001; Grewal et al., 2003). Similar experiments using the Ca²⁺ ionophore, A23187 [GenBank] , or histamine to elevate HUVEC intracellular free Ca²⁺, does not induce cPLA₂ relocation to the Golgi apparatus but to the nuclear envelope (unpublished data). Stimulation-dependent relocation of cPLA₂ to the Golgi apparatus of epithelial cells is seen to place cPLA₂ in close apposition with the AA-metabolizing enzyme COX-1 (Grewal et al., 2003). In HUVECs, costaining revealed no such functional association with COX-1, COX-2, or prostacyclin I synthase upon HUVEC confluence (unpublished data).

Golgi-associated cPLA₂ may play a role in Golgi architecture maintenance or protein trafficking. The PLA₂ family of enzymes are strongly implicated in these processes, specifically via membrane tubule formation (Brown et al., 2003). Previous reports document disruption of the Golgi apparatus upon treatment of cells with PLA₂ inhibitors (de Figueiredo et al., 1998, 1999; Drecktrah and Brown, 1999; Kuroiwa et al., 2001). We see that in HUVECs, the sPLA₂ inhibitor, ONO-RS-082 (Hashimoto et al., 2003), induces Golgi dispersal but pyrrolidine-1 does not (unpublished data). A previous study did report selective blocking of constitutive trafficking to the plasma membrane and retention of proteins in the endoplasmic reticulum of canine kidney cells overexpressing cPLA₂ (Choukroun et al., 2000). But the possible role of cPLA₂ in specific trafficking events in confluent HUVECs has yet to be investigated.

cPLA₂ has previously been linked to cellular proliferation, as seen with studies in macrophages (Tashiro et al., 2004). Although no studies of cPLA₂ activity were conducted, cPLA₂ was seen to regulate expression of the c-myc protooncogene in a B-Myb–dependent manner (Tashiro et al., 2004). AA metabolites also induce endothelial cell proliferation (Dethlefsen et al., 1994; Nie et al., 2000; Antoniotti et al., 2003), and cPLA₂ activity is involved in gene regulation (Pawliczak et al., 2002), but little is known of how these processes are regulated. cPLA₂ is the rate-limiting enzyme of AA evolution and is thus a prime candidate for controlling the synthesis of proproliferative AA metabolites in proliferating and nonproliferating cells. We find that in HUVECs, cPLA₂ activity is essential for proliferation and cell cycle entry and that these functions are mediated by its subcellular localization.

	ACKNOWLEDGMENTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGMENTS REFERENCES

 
We thank M. H. Gelb for providing pyrrolidine-1 and G. J. Howell for help with bioimaging. This work was funded by a Biotechnology and Biological Sciences Research Council PhD studentship (S.P.H), a British Heart Foundation project grant (S.P., J.H.W.), and a Wellcome Trust equipment award to the Leeds Bioimaging Facility (S.P.).

	Footnotes

 
This article was published online ahead of print in MBC in Press (http://www.molbiolcell.org/cgi/doi/10.1091/mbc.E05-02-0164) on June 1, 2005.

Abbreviations used: AA, arachidonic acid; BFA, brefeldin A; cPLA₂, cytosolic phospholipase A₂-; ERGIC-53, endoplasmic reticulum-Golgi intermediate compartment-53; GalT, -1,4-galactosyltransferase; HUVEC, human umbilical vein endothelial cell; iPLA₂, calcium-independent phospholipase A₂; ManII, mannosidase II; sPLA₂, secretory phospholipase A₂

Address correspondence to: J. H. Walker (j.h.walker{at}leeds.ac.uk).

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