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Originally published online as doi:10.1189/jlb.0406251 on January 16, 2007

Published online before print January 16, 2007
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(Journal of Leukocyte Biology. 2007;81:1127-1136.)
© 2007 by Society for Leukocyte Biology

Signaling requirements for translocation of P-Rex1, a key Rac2 exchange factor involved in chemoattractant-stimulated human neutrophil function

Tieming Zhao*, Perihan Nalbant*, Mikio Hoshino{dagger}, Xuemei Dong{ddagger}, Dianqing Wu{ddagger} and Gary M. Bokoch*,1

* Departments of Immunology and Cell Biology, IMM14, The Scripps Research Institute, La Jolla, California, USA;
{dagger} Department of Pathology and Tumor Biology, Kyoto University Graduate School of Medicine, Kyoto, Japan; and
{ddagger} Program of Vascular Biology, Department of Pharmacology, Yale University School of Medicine, New Haven, Connecticut, USA

1 Correspondence: Departments of Immunology and Cell Biology, IMM14, The Scripps Research Institute, 10550 N. Torrey Pines Road, La Jolla, CA 92037, USA. E-mail: bokoch{at}scripps.edu


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
PI 3,4,5-trisphosphate [PI(3,4,5)P3; PIP3]-dependent Rac exchanger 1 (P-Rex1) is a Rac-specific guanine nucleotide exchange factor abundant in neutrophils and myeloid cells. As a selective catalyst for Rac2 activation, P-Rex1 serves as an important regulator of human neutrophil NADPH oxidase activity and chemotaxis in response to a variety of extracellular stimuli. The exchange activity of P-Rex1 is synergistically activated by the binding of PIP3and ß{gamma} subunits of heterotrimeric G proteins in vitro, suggesting that the association of P-Rex1 with membranes is a prerequisite for cellular activation. However, the spatial regulation of endogenous P-Rex1 has not been well defined, particularly in human neutrophils activated through G protein-coupled receptors. Upon stimulation of neutrophil chemoattractant receptors, we observed that P-Rex1 translocated from cytoplasm to the leading edge of polarized cells in a G protein ß{gamma} subunit- and PIP3-dependent manner, where it colocalized with F-actin and its substrate, Rac2. Redistribution of P-Rex1 to the leading edge was also dependent on tyrosine kinase activity and was modulated by cell adhesion. Furthermore, we observed that activation of cAMP-dependent protein kinase A (PKA), which phosphorylates and inactivates P-Rex1, inhibited its translocation. Our data indicate that endogenous P-Rex1 translocates to areas of Rac2 and cytoskeletal activation at the leading edge in response to chemoattractant stimuli in human neutrophils and that this translocation can be negatively modulated by activation of PKA and by cell adhesion.

Key Words: GTPases • chemoattractant receptors • signal transduction • chemotaxis • NADPH oxidase


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
In the mammalian innate immune response, neutrophils serve as a critical line of defense. These cells respond to chemoattractants acting primarily through G protein-coupled receptors (GPCR) and undergo chemotaxis toward invading microorganisms, whereupon they engulf them through the process of phagocytosis [1 2 3 ]. Through the action of a membrane-assembled NADPH oxidase, neutrophils generate reactive oxygen species (ROS) to contain and eliminate ingested pathogens [4 , 5 ]. Rac guanosinetriphosphatases (GTPases), particularly Rac2, are major regulators of these highly complex and coordinated processes, including chemotaxis, phagocytosis, and ROS production [3 ]. Like other small GTPases, Rac2 cycles between a guaosine 5'-diphosphate (GDP)-bound, inactive cytosolic form and a guanosine 5'-triphosphate (GTP)-bound, active membrane-associated form, which promotes neutrophil activation. The formation of Rac2-GTP in response to GPCR chemoattractants such as fMLF has been shown to be rapid, correlating with the onset of neutrophil activation [6 ]. The dissociation of GDP and binding of GTP to Rac are catalyzed by guanine nucleotide exchange factors (GEFs) [7 , 8 ].

PI 3,4,5-trisphosphate [PI(3,4,5)P3; PIP3]-dependent Rac exchanger 1 (P-Rex1) is a major Rac-specific GEF identified in neutrophils on the basis of its Gß{gamma} and PIP3-regulated exchange activity toward Rac GTPase [9 ]. It is evident from several recent studies that P-Rex1 serves as a critical regulator of chemoattractant-stimulated neutrophil motility and ROS production. Treatment of myeloid cells with P-Rex1 antisense oligonucleotide resulted in a substantial decrease in C5a-stimulated ROS production [9 ]. P-Rex1-knockout neutrophils exhibited severe deficiency in chemoattractant-stimulated ROS formation and a reduction of cell speed during chemotaxis [10 , 11 ]. P-Rex1, expressed primarily in myeloid cells and in nervous tissue, has also been implicated in neurotrophin-initiated signaling and neuronal migration [12 ]. These studies indicate the important role of P-Rex1 in host defense and the regulation of inflammation.

A number of biochemical investigations about the regulation of the GEF activity of P-Rex1 have been carried out, providing some molecular insights into how the P-Rex family is regulated. P-Rex1 is dually activated by PIP3, a product of activated PI-3Ks, and ß{gamma} subunits of heterotrimeric G proteins to achieve its full GEF activity in vitro [9 ]. The activation is conferred by the binding of PIP3 to the Plekstrin homology (PH) domain and the Gß{gamma} subunit to the Dbl homology (DH) domain of P-Rex1. Moreover, the DEP, PDZ, and IP4P domains have some modulatory effects on the basal and/or stimulated GEF activity of P-Rex1, suggesting a role for intramolecular interactions between domains of P-Rex1 in the regulation of activity [13 ]. The ability of different Gß{gamma} dimer combinations to activate P-Rex1 has been assessed, with Gß{gamma} dimers containing the ß1 subunit or {gamma}2 subunit being most efficacious, indicating that P-Rex1 is regulated differentially by Gß{gamma} subunits [14 ].

Although the regulation of P-Rex1 GEF activity has been examined extensively through in vitro biochemical studies, the signaling mechanisms regulating its activity in intact cells are poorly delineated to date. As neutrophils express abundant P-Rex1 endogenously, and their activation by chemoattractant stimuli sets in motion an array of signaling pathways leading to Rac-dependent chemotaxis, phagocytosis, and ROS production, the regulation of P-Rex1 in these important inflammatory cells requires examination. Furthermore, as the known in vitro activators of P-Rex1, Gß{gamma} subunits, and PIP3 are tightly associated with the plasma membrane, it would be expected that P-Rex1 must translocate to the site of generation of these regulators during cell activation. In a study where this was investigated in a transfected, nonmyeloid cell line, P-Rex1 was observed to localize largely to the cytoplasm, although a small, variable accumulation of P-Rex1 at the plasma membrane and with subcortical actin was observed in unstimulated- and platelet-derived growth factor (PDGF)-stimulated cells [9 ].

In the present study, we examined the localization of endogenous P-Rex1 in response to chemoattractant stimuli in intact, primary human neutrophils. Our results indicate that P-Rex1 translocates from cytosol to plasma membrane within the leading pseudopod upon cell activation by GPCR agonists, correlating with Rac2 activation. Translocation is dependent on G protein ß{gamma} subunits and PI-3K activation and is modulated by cell adhesion and the action of tyrosine and ser/thr kinases.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Preparation of neutrophils
Human blood was collected from the The Scripps Research Institute (TSRI) General Clinical Research Center (La Jolla, CA, USA) donor pool by venipuncture, and the neutrophils were isolated (95% purity) by dextran sedimentation, hypotonic lysis of erythrocytes, and centrifugation through Ficoll-Paque as described previously [15 ]. The cells were washed with 0.9% NaCl and resuspended in Krebs-Ringer Hepes buffer (118 mM NaCl, 4.8 mM KCl, 25 mM Hepes, 1.2 mM KH2PO4, 1.2 mM MgSO4) containing 5.5 mM glucose.

Membrane translocation—biochemical fractionation assay
Membranes and cytosol of unstimulated and fMLF-stimulated human neutrophil samples were separated by ultracentrifugation as described previously [15 , 16 ]. Briefly, 2.5 x 108 neutrophils were disrupted by two 7 s pulses of sonication using a microtip probe sonicator (Heat Systems, Farmingdale, NY, USA) at low amplification. Unbroken cells and nuclei were removed by low-speed centrifugation, and the resulting supernatant loaded onto 6.7 ml 15% sucrose layered over 4 ml 34% sucrose. After centrifugation for 40 min at 4°C and 195,000 g, cytosol was removed from the top of the gradient, and the light membrane fraction was collected from the interface of the 15% and 34% sucrose solution. Total yields averaged 150 µg membrane and 3400 µg cytosol. The cytosol (100 µg) and membranes (100 µg) were then subjected to SDS-PAGE followed by immunoblotting with 1:1000 dilution of rat anti-P-Rex1 mAb [12 ], 1:10,000 dilution of rabbit anti-Rho guanine dissociation inhibitor (RhoGDI) R922 antibody [17 ], or 1:2000 dilution of rabbit anti-Rac2 R786 antibody [16 ] to assess translocation of P-Rex1 or Rac2 to the membrane. In some experiments, a rabbit polyclonal antibody directed against P-Rex1 amino acids 1–57, generated in our laboratory, was used to verify results. Both P-Rex1 antisera were specific for P-Rex1, as indicated by the loss of the immunoreactive band in lysates from P-Rex1 knockout mouse neutrophils [10 ]. As the intensity of the cytosolic P-Rex1 band on the immunoblot (see Fig. 1A ) is similar to that of membrane-associated P-Rex1, we calculate in this experiment that 4.4% of P-Rex1 was translocated.


Figure 1
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  		Figure 1. P-Rex1 translocation from cytosol to plasma membrane in human neutrophils. (A) Neutrophils (2.5x108) were incubated with or without 1 x 10–7 M fMLF for 1 min, and membrane and cytosol fractions were prepared as described in Materials and Methods. Samples of 100 µg cytosol or membrane fraction were run on SDS-PAGE and immunoblotted with anti-P-Rex1 or anti-Rac2 antibody to analyze the presence of P-Rex1 and Rac2 on the cell membrane or with RhoGDI antibody as control for cytosol, respectively. The last lane of the GDI panel represents 10 µg cytosol as a RhoGDI antibody standard. Molecular weight marker sizes are indicated at the left of each panel. Results shown are representative of two out of four experiments (see text). (B) Neutrophils (3x108 cells) were incubated with 1 x 10–7 M fMLF (+FP) or buffer control (–FP) for 2 min and cavitated in an N2 cavitation chamber in the presence of 1 mM PMSF and 100 U/ml aprotinin, and cell lysates were fractionated on discontinuous sucrose gradients, as detailed in refs. [18 , 19 ] and in Materials and Methods. Aliquots (40 µl) of the indicated fractions were run on SDS-PAGE and immunoblotted with anti-P-Rex1, anti-Rac2; anti-RhoGDI; or anti-Gß antibodies (Gb), as in Materials and Methods. The resulting immunoblots were quantified by densitometry and plotted as arbitrary units. The location of major neutrophil subcellular organelles on such gradients has been determined in detail using various marker enzymes (see refs. [18 19 20 21 ]). The plasma membrane peak, identified using alkaline phosphatase (Alk Phos), is shown. RhoGDI served as a cytosolic marker. Results shown are representative of two similar experiments.

 
For analysis of cell fractions by discontinuous sucrose gradients, samples were prepared and analyzed on 15%/40%/60% sucrose gradients, as described thoroughly [18 ]. In these studies, a specific Gß antibody (R3,4) was used for immunoblotting at 1:1000 dilution [19 ]. The localization of the major neutrophil subcellular organelles on such gradients is determined using various marker enzymes characteristic of each organelle [18 19 20 21 ]. Recoveries of total activity on the gradients averaged ~80%. In the current study, we used RhoGDI as a cytoplasmic marker and alkaline phosphatase as a plasma membrane marker. Granule-containing fractions (past Fraction 14) are not shown here, as they were devoid of the proteins of interest.

Immunofluorescence microscopy
Cells were treated with 1 x 10–7 M fMLF, 1 µg/ml PMA, 10 nM C5a, 100 ng/ml LPS, or 10 ng/ml GM-CSF (all from Sigma Chemical Co., St. Louis, MO, USA) for various times, the reactions were stopped on ice, and the cells were mounted gently onto cover glass (Fisher Scientific, Pittsburgh, PA, USA) using a Shandon Cytospin Cytocentrifuge (Thermo Electron Corp., Waltham, MA, USA). Cytocentrifugation did not change the morphological appearance or behavior of the neutrophils in our studies. After fixing in 4% of paraformaldehyde for 15 min, the cells were permeabilized in 0.5% Triton X-100 for 10 min. The cover glass was incubated for 1 h with 5 µg/ml rat anti-P-Rex1 mAb, followed by incubation with 1:500 secondary, Alexa 488-conjugated anti-rat IgG (Molecular Probes, Eugene, OR, USA) for 1 h. The subcellular distribution of P-Rex1 was then examined with a 40x/0.6 aperture objective lens under a Nikon Eclipse TE2000-U microscope (Nikon, Tokyo, Japan). The specificity of P-Rex1 immunofluorescence with the P-Rex1 mAb was verified by comparative staining of P-Rex1 null mouse neutrophils versus wild-type cells after fMLF stimulation (data not shown). Images were acquired with the Photometrics CoolSnap HQ camera (Photometrics, Tucson, AZ, USA) and Metamorph software (Molecular Devices, Sunnyvale, CA, USA) and processed using Adobe Photoshop (Adobe Systems, San Jose, CA, USA). The percentage of cells with P-Rex1 membrane translocation was obtained by counting 200 cells under each experimental condition in each experiment (given as mean ± SE). In some experiments, fixed cells were incubated with 1:100 Alexa 568-conjugated phalloidin (Molecular Probes) for 1 h to assess the distribution of F-actin or 1:200 rabbit anti-Rac2 R786 antibody [16 ] for 1 h, followed by 1:500 secondary, Alexa 568-conjugated anti-rabbit IgG (Molecular Probes) for 1 h to detect Rac2 distribution.

Confocal images were obtained on a Rainbow Radiance 2100 multilaser-scanning confocal system (Bio-Rad Zeiss, Hercules, CA, USA), attached to a Nikon TE2000 U microscope with a 63x/1.4 oil objective. Confocal slice thickness was 0.54 µm. Image processing was performed with MetaMorph, ImageJ (http://rsb.info.nih.gov/ij/), and Adobe Photoshop.

In some experiments, neutrophils were treated with 2 µg/ml latrunculin A (Biomol, Plymouth Meeting, PA, USA) for 10 min, 0.5 µg/ml pertussis toxin (PTX; List Biologicals, Campbell, CA, USA) for 30 min, 100 nM wortmannin (Sigma Chemical Co.) for 20 min, 50 µM LY294002 (Calbiochem, San Diego, CA, USA) for 20 min, 10 µM M119 or 10 µM M119B [22 ] for 5 min, 100 µM genistein (Sigma Chemical Co.) for 30 min, 10 µM forskolin (Sigma Chemical Co.) for 15 min, 10 µM H-89 (Calbiochem) for 20 min, 2 mM 6-Bnz-cAMP (Biolog, Bremen, Germany) for 20 min, 2.6 µM tyrphostin A9 (Calbiochem) for 30 min, 100 µM piceatannol (Sigma Chemical Co.) for 10 min, 10 µM PP1 (Biomol) for 30 min, or 2 µM SU6656 (Calbiochem) for 30 min, before they were treated with various stimuli. For adherent cells, the cover glass (Fisher Scientific) was coated with 50 µg/ml human plasma fibronectin (Sigma Chemical Co.) overnight at 4°C. Freshly prepared neutrophils were then warmed for 10 min at 37°C on the cover glass and treated with 1 x 10–7 M fMLF for the indicated times (as in ref. [23 ]). Cells were then fixed as in ref. [23 ], and P-Rex1 distribution was detected as above.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
P-Rex1 translocates to the plasma membrane in chemoattractant-stimulated human neutrophils
In response to human neutrophil activation by the chemotactic peptide fMLF, a heterotrimeric GPCR agonist, the Rac2 GTPase is activated and translocates from its localization in the cytosol of resting cells to the plasma membrane [15 , 16 ]. It has been suggested that this Rac2 membrane translocation is coupled directly to its activation by membrane-associated GEFs [24 ]. P-Rex1 has been identified recently as a major Rac2 GEF in fMLF-stimulated neutrophils [10 , 11 ]. Disruption of the P-Rex1 gene in mice results in nearly complete absence of fMLF-induced Rac2 activation, accompanied by defects in chemotaxis and NADPH oxidase activation. G protein ß{gamma} subunits and phosphoinositides such as PIP3 have been shown to synergistically regulate the GEF activity of P-Rex1 in vitro and in an in vivo porcine aortic endothelial (PAE) cell model [9 ]. However, P-Rex1 was found primarily in the cytosol in these cells, and cell stimulation by PDGF did not induce the association of P-Rex1 with the cell membrane, where the regulatory ß{gamma} subunits and PIP3 are found. Nor has the regulation of P-Rex1 function by GPCR chemoattractant been verified in neutrophils or other myeloid cells in which the function of P-Rex1 has been shown to be critical.

We examined whether P-Rex1 underwent translocation from the cytosol, where it resides in resting neutrophils, to the plasma membrane in response to GPCR chemoattractant stimuli. Freshly prepared human neutrophils were stimulated with the chemoattractant fMLF, and membrane fractions were obtained using sucrose density gradient fractionation. The cytosol and membrane fractions were then subjected to Western blot to assess the presence of proteins of interest. As shown previously [16 ], 5–10% of total cellular Rac2 was observed consistently to translocate to the membrane fraction of fMLF-stimulated cells, indicating that the neutrophils were activated properly (Fig. 1A ). RhoGDI served as a cytosolic protein control, indicating there was no contamination of the membrane fractions with cytosol. Similar to Rac2, we estimated that ~5% of total P-Rex1 translocated to the membrane fraction (Fig. 1A) . However, this association was not stable, as we observed clear and substantial membrane association in only two out of four experiments, suggesting that the interaction of P-Rex1 with the plasma membrane could be disrupted by the fractionation and membrane separation procedure.

To preserve the association of P-Rex1 with the plasma membrane fraction, we fractionated neutrophils using milder procedures (Fig. 1B) : Lysates were prepared by nitrogen cavitation from unstimulated or fMLF-stimulated neutrophils, and subcellular fractions were separated on discontinuous sucrose gradients, as described previously [18 19 20 21 ]. In fMLF-stimulated cells, we observed in two out of two experiments that a fraction of the P-Rex1 (5.1% and 10% in respective experiments) became associated with the peak of plasma membrane (marker enzyme, alkaline phosphatase). G protein ß{gamma} subunits were also present in this plasma membrane fraction, as reported previously [19 ]. The remainder of P-Rex1 remained localized in the cytosolic fraction along with RhoGDI. These data confirm biochemically the translocation of P-Rex1 to the plasma membrane in fMLF-stimulated human neutrophils.

To examine the requirements for P-Rex1 translocation further, we used immunofluorescence microscopy to assess the subcellular localization of endogenous human neutrophil P-Rex1 in response to chemoattractant stimulation. Immunostaining with P-Rex1 mAb revealed that in resting neutrophils, P-Rex1 was distributed throughout the cytoplasm. Upon stimulation with fMLF, the neutrophils developed a polarized morphology with an F-actin-rich, single leading edge. Confocal microscopy demonstrated that endogenous P-Rex1 became localized rapidly at the leading edge (Fig. 2A and B ), where it colocalized with F-actin (see Fig. 3A ). Using this nondisruptive method, P-Rex1 membrane translocation was observed consistently in greater than 70% of polarized, fMLF-stimulated neutrophils. Control Western blots showed that the P-Rex1 antibody used for these studies did not react with lysates from P-Rex1 null mouse neutrophils nor did they show significant immunofluorescence in non-P-Rex1-containing human embryo kidney-293 cells or in P-Rex1 null mouse neutrophils (data not shown).


Figure 2
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  		Figure 2. P-Rex1 localization in response to different stimuli. (A) Neutrophils were treated with 1 x 10–7 M fMLF for 1 min, 1 µg/ml PMA for 10 min, 10 nM C5a for 5 min, or 100 ng/ml LPS for 60 min. In some experiments, neutrophils were pretreated with 100 ng/ml LPS before stimulation by 1 x 10–7 M fMLF for 1 min. Cells were then fixed and immunostained with anti-P-Rex1 antibody and visualized using confocal microscopy. A representative image from four independent experiments is shown. Original bar, 8 µm. (B) Neutrophils were treated with 1 x 10–7 M fMLF for indicated times, and the percentage of cells with polarized distribution of P-Rex on plasma membrane was assessed and plotted as described in Materials and Methods.

 

Figure 3
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  		Figure 3. Colocalization of P-Rex1 with F-actin. (A) Neutrophils were treated with 1 x 10–7 M fMLF for 1 min or 10 nM C5a for 5 min and fixed. Cells were then stained with anti-P-Rex1 antibody and Alexa 568-conjugated phalloidin. (B) Neutrophils were treated with 2 µg/ml latrunculin A (Lat), followed by 1 x 10–7 M fMLF for 1 min. Cells were then stained with anti-P-Rex1 antibody and with Alexa 568-conjugated phalloidin. (C) Neutrophils were treated with 1 x 10–7 M fMLF for 1 min, fixed, and stained with anti-Rac2 R786 antibody and anti-P-Rex1 antibody. Confocal images were acquired as described in Materials and Methods. Representative datasets from three independent experiments are shown (A–C). Original bars, 8 µm.

 
We determined the time course of fMLF-induced P-Rex1 membrane translocation (Fig. 2B) . After 15 s of activation with fMLF, P-Rex1 translocation was apparent: About 60% of the total cells displayed substantial P-Rex1 membrane localization at 15 s, increasing to just over 70% by 1 min. It is interesting that P-Rex1 was still present in the membrane fraction in over 50% of the cells at 5 min after stimulation. This time course correlates well with the time course of fMLF-stimulated Rac2 activation and ROS production, as previously established [6 , 16 ]. We also treated neutrophils with the complement component C5a, another chemoattractant that signals through GPCR, and assessed the localization of endogenous P-Rex1. Using confocal microscopy, we verified the ability of C5a to stimulate the P-Rex1 translocation to the leading edge (Fig. 2A) , where it colocalized with F-actin (Fig. 3A ).

To test the idea that the primary association of P-Rex1 was with the underlying plasma membrane, we preincubated neutrophils with latrunculin to inhibit F-actin assembly [25 ] and then examined the translocation of P-Rex1. As seen in Figure 3B , stimulation by fMLF induced a substantial redistribution of P-Rex1 to the cell periphery, although the cells could no longer polarize and form an actin-rich leading edge. Along with the biochemical fractionation data (Fig. 1) , these results strongly support the chemoattractant receptor-stimulated translocation of a pool of P-Rex1 to the plasma membrane.

The time course of P-Rex1 translocation (Fig. 2B) correlates well with that of Rac2 activation in neutrophils, and Rac2 activation has been shown to be dependent on P-Rex1 activity in knockout mouse neutrophils [10 ]. We therefore examined the subcellular localization of Rac2 to determine its spatial relationship with P-Rex1. In fMLF-stimulated cells, a fraction (5–10%; see ref. [16 ]) of Rac2 translocated to and accumulated at the leading edge (Fig. 3C) , where it becomes activated [26 ]. A substantial fraction of this Rac2 at the leading edge colocalized with translocated P-Rex1 (Fig. 3C , Overlay), consistent with the role of P-Rex1 in Rac2 activation.

We next examined whether P-Rex1 translocation can be induced by other, non-GPCR neutrophil stimuli. PMA stimulates neutrophil function through the direct activation of protein kinase C (PKC), leading to activation of Rac2 and the NADPH oxidase [6 ]. However, PMA did not stimulate P-Rex1 translocation (Fig. 2A) . This is consistent with the lack of effect of P-Rex1 deficiency on PMA-induced superoxide formation [9 10 11 ]. LPS, a component of the outer cell wall of Gram-negative bacteria and an agonist for TLRs, primes neutrophils and renders an enhanced granule secretion and ROS production when neutrophils respond following activation by fMLF. LPS-primed neutrophils were particularly sensitive to the absence of P-Rex1, as P-Rex1-deficient cells were totally deficient in their ability to generate ROS in response to LPS priming and fMLF or C5a stimulation [11 ]. This result suggested the possibility that LPS-mediated priming could involve P-Rex1 translocation. However, we observed that LPS treatment did not induce P-Rex1 translocation (Fig. 2A) , implying that pretranslocation of P-Rex1 is not involved in LPS priming of neutrophils. The fact that LPS failed to cause P-Rex1 translocation, in combination with the observation that PMA did not induce P-Rex1 translocation, strongly indicates that P-Rex1 translocation requires, and might be regulated specifically by, the activation of GPCR.

P-Rex1 translocation is dependent on G protein ß{gamma} signaling and PI-3K activity
To provide direct evidence that P-Rex1 translocation requires the activation of heterotrimeric G protein by GPCR, neutrophils were pretreated with PTX, which is known to inactivate Gi in the fMLF signaling pathway of human neutrophils [27 ]. Stimulation with fMLF failed to induce P-Rex1 translocation in PTX-treated cells (Fig. 4A ). Recently, novel compounds were identified that bind to the G protein ß subunit and interfere with its interactions with effector targets [22 ]. We examined the ability of the active M119 and the corresponding, inactive M119B compounds to affect P-Rex1 translocation. Figure 4A shows that M119, but not M119B, effectively blocked P-Rex1 translocation. These data are thus consistent with the requirement for Gß{gamma} in P-Rex1 activation.


Figure 4
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  		Figure 4. Dependence of P-Rex1 translocation on GPCR and PI-3Ks. Neutrophils were pretreated with 0.5 µg/ml PTX (P. Toxin) or with 100 nM wortmannin (Wort.), 50 µM LY294002, 10 µM M119, or 10 µM M119B for the times indicated in Materials and Methods, followed by 1 x 10–7 M fMLF for 1 min (A) or with 10 ng/ml GM-CSF for the indicated times (B). Cells were then fixed and immunostained with anti-P-Rex1 antibody. The percentage of cells with polarized distribution of P-Rex on the plasma membrane was assessed and plotted as described in Materials and Methods. A representative dataset from three independent experiments is shown.

 
P-Rex1 activation also requires PI-3K-mediated formation of PIP3 [9 ]. Neutrophils were treated with the selective PI-3K inhibitors, wortmannin and Ly294002, before stimulation with fMLF. We observed that both inhibitors abolished P-Rex1 translocation effectively (Fig. 4A) . To evaluate whether the synergistic activation of P-Rex1 by Gß{gamma} subunits and PIP3 is also reflected in their roles in membrane translocation of P-Rex1, we assessed the ability of GM-CSF, which activates PI-3K through tyrosine kinase receptor signaling but does not stimulate the formation of free Gß{gamma}, to induce P-Rex1 translocation. GM-CSF, added for times ranging from 0 to 20 min, did not induce substantial P-Rex1 translocation (Fig. 4B) , suggesting that the formation of PIP3 alone may not be sufficient to induce P-Rex1 translocation. These data establish that P-Rex1 translocation correlates closely with P-Rex1 activation and that it may require the formation of PIP3 and release of free Gß{gamma}.

P-Rex1 translocation is regulated by cell adhesion
We have shown previously that adhesion to extracellular matrix inhibits Rac2 activation and subsequent ROS production at early times in chemoattractant-stimulated human neutrophils [23 ]. In the present study, we found that adhesion inhibited early fMLF-induced P-Rex1 translocation as well (Fig. 5A and B ). As had been observed with Rac2 activation and ROS production, at later times ranging from 30 to 90 min after stimulation, P-Rex1 translocation took place (Fig. 5B) . P-Rex1 translocation was correlated with restoration of fMLF-induced PI-3K activity, as assessed by visualizing the phosphorylation of the downstream mediator, Akt (Fig. 5C) .


Figure 5
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  		Figure 5. Suppression of P-Rex1 translocation by cell adhesion. Adherent or suspended neutrophils were treated with 1 x 10–7 M fMLF for 1 min (A), or adherent cells were treated with 1 x 10–7 M fMLF for the indicated times (B and C). Fixed cells were incubated with anti-P-Rex1 antibody and visualized using fluorescence microscopy. Akt phosphorylation (P-Akt; C) was determined as described in Materials and Methods at T = 0, 1, or 60 min. Representative datasets from two or three independent experiments are shown. Original bars, 8 µm.

 
P-Rex1 translocation is regulated by protein kinase activities
Rac2 activation, stimulated by fMLF, has been shown to be inhibited by tyrosine kinase inhibitors [6 ]. We evaluated whether P-Rex1 translocation depends on protein tyrosine kinases, which have been shown to be important regulators of several Rac-GEFs. Indeed, genistein treatment strongly blocked P-Rex1 translocation, suggesting the possibility that P-Rex1 may be tyrosine-phosphorylated (Fig. 6A ). We expanded our analysis of the role of tyrosine kinases in P-Rex1 activation (Fig. 6B) . Selective inhibitors of Pyk2 (Tyrphostin A9), Syk (piceatannol), and general Src family kinases (PP1) substantially blocked P-Rex1 translocation induced by fMLF. In contrast, a selective inhibitor of c-Src (SU6656) had little effect on P-Rex1. These data suggest complex regulation of P-Rex1 through various tyrosine kinase-mediated signaling pathways.


Figure 6
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  		Figure 6. Effects of protein tyrosine kinases and PKA on P-Rex1 translocation. Neutrophils were treated with 100 µM genistein, 10 µM forskolin, 10 µM H89, and 2 mM 6-Bnz-cAMP (A) and 2.6 µM tyrphostin A9, 100 µM piceatannol (Pic.), 10 µM PP1, or 2 µM SU6656 (B) for 10 min, followed by 1 x 10–7 M fMLF for 1 min. Cells were then fixed and immunostained with anti-P-Rex1 antibody. The percentage of cells with polarized distribution of P-Rex was assessed and plotted as described in Materials and Methods. **, Values were significantly different from the means of the respective controls at P < 0.01, n = 3.

 
A recent study showed that cAMP-dependent PKA phosphorylated P-Rex1 and inhibited its GEF activity toward Rac [28 ]. We observed that the general cAMP-elevating agent forskolin and the PKA activation-specific cAMP analog, 6-Bnz-cAMP [29 ], inhibited P-Rex1 translocation significantly (Fig. 6A) . These inhibitions were reversed substantially by preincubation with the PKA inhibitor H89. Our data indicate that PKA also inhibits P-Rex1 function in vivo by blocking its association with the plasma membrane, where activating Gß{gamma} subunits and PIP3 reside. It is interesting that Rac2 translocation appeared to be only affected minimally by PKA-mediated inhibition of P-Rex1 translocation (data not shown).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
P-Rex1 has been studied largely at the in vitro biochemical level since its initial isolation from porcine neutrophils as a Rac-specific GEF whose activity was synergistically regulated by Gß{gamma} and PIP3. Subsequent studies have shown the Gß{gamma} subunits to interact directly with the DH domain of P-Rex1, and PIP3 interacts via the adjacent PH domain; the DEP, PDZ and IP4P domains play some modulatory roles [13 ]. The basis for the synergistic action of these two regulatory molecules has not been defined. Such in vitro biochemical studies provide important clues about how P-Rex1 may be regulated, but there is little or no information about P-Rex1 regulation in intact cells, particularly in neutrophils where P-Rex1 plays such important functional roles. Although endogenous P-Rex1 is localized primarily within the cytosol in human neutrophils, its known regulators Gß{gamma} and PIP3 are plasma membrane-associated. P-Rex1 activation would consequently be predicted to correlate with the recruitment of P-Rex1 to the plasma membrane of stimulated cells.

We observed that P-Rex1 translocated to the plasma membrane in human neutrophils stimulated with the GPCR agonists fMLF or C5a (Figs. 1 2 3) . The timing of the translocation was correlated closely with the time course of Rac2 activation and translocation induced by these same stimuli [6 , 16 ]. Plasma membrane-associated P-Rex1 localized with G protein ß{gamma} subunits (Fig. 1B) . The translocation of P-Rex1 to the cell periphery in neutrophils pretreated with the F-actin-disrupting drug latrunculin (Fig. 3B) in combination with subcellular fractionation data (Fig. 1) provides evidence that at least a portion of P-Rex1 indeed localizes to the plasma membrane itself and not to the membrane-associated cytoskeleton. At the membrane, P-Rex1 colocalized with Rac2 and F-actin within the leading edge of polarized neutrophils (Figs. 3A and 3C) . Translocation was not induced by the toll-like receptor stimulus LPS under conditions where priming of the fMLF-stimulated NADPH oxidase was observed (not shown), indicating that induction of P-Rex1 translocation does not account for the priming effect of LPS (Fig. 2A) .

P-Rex1 translocation was blocked almost entirely after treatment of cells with PTX (Fig. 4A) , which blocks receptor-stimulated release of Gß{gamma} subunits from Gi [27 ]. Similarly, a recently described inhibitor of Gß{gamma} subunits (M119) also blocked P-Rex1 translocation [22 ]. Although signaling from pathways initiated downstream of Gß{gamma} cannot be ruled out, these data strongly implicate the participation of Gß{gamma} subunits in P-Rex1 activation in GPCR-stimulated human neutrophils. Indeed, this is supported by the association of P-Rex1 with Gß{gamma} subunits in the plasma membrane of fMLF-stimulated cells (Fig. 1B) . Translocation was also blocked effectively by the PI-3K inhibitors wortmannin and Ly294002 (Fig. 4A) . This is consistent with the in vitro regulation of P-Rex1 by phosphoinositides, which are formed and reside within the plasma membrane after activation of GPCR. Our data suggest that there is a close correlation between P-Rex1 membrane translocation and its activation, as the regulatory requirements are identical. We cannot, however, rule out the possibility that there is an additional activation step that occurs subsequent to membrane recruitment.

We examined whether the stimulation of PIP3 formation was sufficient to recruit P-Rex1 to the membrane. When we used the growth factor stimulus GM-CSF to selectively activate PI-3K in the absence of Gß{gamma} release, we observed little translocation of P-Rex1 (Fig. 4A and 4B) . In conjunction with the inhibitor data, this observation suggests that there is a requirement for free Gß{gamma} and PIP3 for P-Rex1 to associate effectively with the membrane. This may account for the prior failure to observe substantial membrane association of transfected P-Rex1 in PDGF-stimulated PAE cells [9 ]. It is possible, however, that the level of PIP3 formed in response to GM-CSF or PDGF is simply insufficient to induce translocation of P-Rex1. It is interesting to note that in nonpolarized, latrunculin-treated neutrophils, stimulation with fMLF induces translocation of P-Rex1 around the entire cell periphery (Fig. 3B) . In contrast, in normal polarized cells, the P-Rex1 localizes primarily at the leading edge (e.g., Fig. 3A ). It has been shown that latrunculin treatment abolishes the polarized distribution of PIP3 to the leading edge as well [30 ]. In contrast, the distribution of Gß{gamma} subunits does not appear to be localized to the leading edge in chemotaxing cells [31 ]. This suggests that although Gß{gamma} may provide most of the membrane-binding affinity, PIP3 might act in synergy to direct where P-Rex1 becomes localized and activated.

Our studies also identified other signaling pathways involved in regulating P-Rex1 membrane recruitment. Protein tyrosine kinase activity was required for fMLF-induced translocation (Fig. 6) . It has been observed that Rac2 activation by fMLF is also sensitive to genistein [6 ] and Tyrphostin A9 [32 ]. It is not clear whether this is a result of a direct phosphorylation of P-Rex1 or through an indirect action on another component such as Gß{gamma}. It is interesting that we found that the ability of fMLF to stimulate P-Rex1 translocation at short times was suppressed in adherent neutrophils (Fig. 5A 5B 5C) . We had demonstrated previously that Rac2 activation is inhibited at early times by neutrophil adherence and that this correlated with inhibition of tyrosine phosphorylation of the Vav1 Rac2 GEF [23 ]. Our current data suggest the possibility that adhesion may also be blocking Rac2 activation through effects on P-Rex1. As fMLF-induced Rac2 activation has been shown to require Vav1 [33 ] and P-Rex1 [10 , 11 ] proteins in knockout mice, it will be of interest to examine the functional relationship between these two regulatory GEFs in neutrophils.

It was recently shown that P-Rex1 can be phosphorylated by PKA and that this phosphorylation inhibits its Gß{gamma}- and PIP3-stimulated Rac GEF activity [28 ]. One mechanism for the inhibitory effect of phosphorylation was shown to be a 47-fold reduction in the potency of Gß{gamma} for activating P-Rex1 GEF activity. This study suggested that the phosphorylation of P-Rex1 by PKA reduced its ability to bind Gß{gamma} dimers. In the present study, we have shown that in fMLF-stimulated human neutrophils, elevation of cellular cAMP levels by the addition of forskolin or a membrane-permeable cAMP analog inhibited P-Rex1 membrane translocation effectively (Fig. 6A) . This effect was mediated by the activation of PKA, as it was blocked by pretreatment with the PKA-selective inhibitor H89 and was initiated by 6-Bnz-cAMP, which selectively activates PKA [29 ]. Our data are consistent with a PKA-mediated decrease in the binding affinity of P-Rex1 for Gß{gamma}, resulting in ineffective recruitment to the plasma membrane. These results support our conclusion that interaction of P-Rex1 with Gß{gamma} is a key step in mediating membrane association. It is interesting that Rac2 translocation did not appear to be reduced substantially when P-Rex1 translocation was inhibited by PKA. This suggests that the recruitment of Rac2 to the membrane is not mediated by P-Rex1 directly.

Dbl family GEFs are important regulators for cell growth, differentiation, and migration [34 ]. By catalyzing GDP exchange for GTP, they activate their substrate Rho GTPases and exert subsequent biological functions [7 , 8 ]. There is accumulating evidence that regulation of subcellular localization is a common theme during the activation of Dbl family GEFs. For example, T cell-lymphoma invasion and metastasis-1 (Tiam1) translocates from cytoplasm to plasma membrane in response to serum and platelet-derived growth factor stimulation [35 , 36 ]. This redistribution was required for the activation of Rac1 by Tiam1. Vav1 translocates to the plasma membrane in a PI-3K-dependent manner during Rac activation in macrophages [37 , 38 ]. The Rho GEF activity of GEF-H1 is regulated by its binding or release from microtubules [39 ]. Many of these translocation events are controlled by the interaction of the PH domains of these GEFs with membrane-generated phosphoinositides and can be modulated by phosphorylation [7 , 8 ].

This study reports four major findings with regard to the regulation of P-Rex1 activity. First, we establish that endogenous P-Rex1 does translocate from the cytosol of resting cells to the plasma membrane of cells stimulated through GPCR. This translocation is dependent primarily on the release of G protein ß{gamma} subunits but also requires PI-3K activity, perhaps to specify the area of localization. These data suggest there is a close correlation between P-Rex1 translocation and its activation. Second, we show that P-Rex1 primarily becomes localized to the leading edge of stimulated, polarized neutrophils, where it colocalizes significantly with Rac2 and F-actin. This places P-Rex1 in position to regulate localized Rac2 activity during chemotaxis. Third, we establish that P-Rex1 translocation is regulated by tyrosine kinase activity and by cell adhesion. The latter observation is consistent with the suppression of Rac2 activation and subsequent ROS formation in newly adherent neutrophils [23 ]. Fourth, we show that P-Rex1 activation is inhibited by elevation of cellular cAMP, acting through the activation of PKA. As cAMP is known to inhibit chemotaxis and NADPH oxidase activation in neutrophils [40 41 42 ], our results raise the possibility that one mechanism of action of cAMP to inhibit these Rac2-dependent inflammatory cell functions may be through the suppression of Rac2 activation by P-Rex1. Similarly, it has been noted that the spatial regulation of PKA activity in human neutrophils is involved in chemoattractant-induced polarization and directed cell motility [43 , 44 ]. Our observations suggest that this may in part be a result of modulation of localized P-Rex1 activity. The studies described here provide novel molecular insights into how P-Rex1 and its activation are regulated in vivo and how such regulation may contribute to inflammatory cell responsiveness.


   ACKNOWLEDGEMENTS
 
This work was supported by National Institutes of Health grants GM39434 and HL48008 (to G. M. B.) and HL70694 and HL80706 (to D. W.) and a fellowship from the Arthritis Foundation, San Diego Chapter (to T. Z.). The Gß{gamma} inhibitors were generous gifts from Alan Smrcka, University of Rochester (Rochester, NY, USA). The authors acknowledge the assistance of M. Crawford and H. Zhang (TSRI) with neutrophil preparations.

Received April 5, 2006; revised August 28, 2006; accepted December 4, 2006.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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