Angiotensin II Type 2 Receptor Stimulation Increases the Rate of ...

0013-7227/08/$15.00/0 Printed in U.S.A.

Endocrinology 149(6):2923–2933 Copyright © 2008 by The Endocrine Society doi: 10.1210/en.2007-0313

Angiotensin II Type 2 Receptor Stimulation Increases the Rate of NG108 –15 Cell Migration via Actin Depolymerization Peter Kilian, Shirley Campbell, Lyne Bilodeau, Marie-Odile Guimond, Claude Roberge, Nicole Gallo-Payet, and Marcel Daniel Payet De´partment de Physiologie et Biophysique (P.K., S.C., L.B., N.G.-P., M.D.P.) and Service d’Endocrinologie (M.-O.G., C.R., N.G.-P.), De´partement de Me´decine, Faculte´ de Me´decine et des Sciences de la Sante´, Universite´ de Sherbrooke, Sherbrooke, Que´bec, Canada J1H 5N4 Angiotensin II (Ang II) has been reported to induce migration in neuronal cell types. Using time-lapse microscopy, we show here that Ang II induces acceleration in NG108 –15 cell migration. This effect was antagonized by PD123319, a selective AT2 receptor antagonist, but not by DUP753, a selective AT1 receptor antagonist, and was mimicked by the specific AT2 receptor agonist CGP42112. This Ang II-induced acceleration was not sensitive to the inhibition of previously described signaling pathways of the AT2 receptor, guanylyl cyclase/cyclic GMP or p42/p44mapk cascades, but was abolished by pertussis toxin treatment and involved PP2A activation. Immunofluorescence studies indicate that Ang II or CGP42112 decreased the amount of filamentous actin at the leading edge of the cells. This decrease was accompanied by a concomitant increase in globular actin levels. Regulation of actin turnover

T

HE ANGIOTENSIN II (Ang II) octapeptide, the active component of the renin-angiotensin system, binds and activates two major types of seven-transmembrane domain G protein-coupled receptors, namely type 1 (AT1) and type 2 (AT2). Much of the classical actions of Ang II, including modulation of blood pressure, control of fluid/electrolyte balance, and cellular proliferation, are associated with activation of the AT1 receptor (1, 2). In the adult, activation of the AT2 receptor is known to modulate negatively the effects associated with AT1 receptor activation (for review, see Refs. 2– 6). One of the most striking features of the AT2 receptor is its high level of expression in several fetal tissues, including the brain (7–9). At the cellular level, the AT2 receptor has been localized in neurons, but not in astrocytes or glial cells (10, 11). The presence of AT2 receptors in restricted brain areas of First Published Online March 6, 2008 Abbreviations: ADF, Actin depolymerizing factor; ANG II, angiotensin II; Cfl1, cofilin-1; cGMP, cyclic GMP; F-actin, filamentous actin; FBS, fetal bovine serum; FN, fibronectin; G-actin, globular actin; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; K-S Dist, Kolmogorov-Smirnov normality test; L-NAME, NG-nitro-l-arginine methyl ester; MAP, microtubule-associated protein; MEK, MAPK kinase; NGF, nerve growth factor; OA, okadaic acid; P-ADF, phosphorylated actin depolymerizing factor; pNPP, p-nitrophenyl phosphate; PTX, pertussis toxin; SDS, sodium dodecyl sulfate; siRNA, small interfering RNA. Endocrinology is published monthly by The Endocrine Society (http:// www.endo-society.org), the foremost professional society serving the endocrine community.

in actin-based motile systems is known to be mainly under the control of the actin depolymerizing factor and cofilin. Basal migration speed decreased by 77.2% in cofilin-1 small interfering RNA-transfected NG108 –15 cells, along with suppression of the effect of Ang II. In addition, the Ang II-induced increase in cell velocity was abrogated in serum-free medium as well as by genistein or okadaic acid treatment in a serumcontaining medium. Such results indicate that the AT2 receptor increases the migration speed of NG108 –15 cells and involves a tyrosine kinase activity, followed by phosphatase activation, which may be of the PP2A type. Therefore, the present study identifies actin depolymerization and cofilin as new targets of AT2 receptor action, in the context of cellular migration. (Endocrinology 149: 2923–2933, 2008)

the adult and its broad distribution in fetal tissue supports a role for this receptor subtype in neuronal development and function (12). Neuronal cell lines such as NG108 –15 (13–15), PC12W (16), and N1E-115 (17) express the AT2 receptor at high levels and have thus far been used to investigate the role and mechanisms of action of AT2. Specific activation of the AT2 receptor induces neurite outgrowth in NG108 –15 cells (18) and PC12W cells (16), as well as in primary cultures of rat cerebellar granule cells (19). A role of the AT2 receptor in cell migration was also demonstrated, both in primary cultures of rat cerebellar granule cells (19) and in dorsal root ganglia neurons during axonal regeneration after injury in the adult (20, 21). In our endeavor to elucidate AT2 receptor involvement in Ang II-induced neuronal differentiation, we and others have investigated signaling mechanisms activated by this receptor. This seven-transmembrane domain receptor is not coupled to any of the classical, well-established second messengers such as cAMP or inositol phosphates (for review, see Refs. 1, 11, and 22–24). In both NG108 –15 (25) and PC12W cells (26), Ang II-induced p42/p44mapk activation appears essential for inducing neurite outgrowth. Associated mechanisms involve the activation of the Rap1/B-Raf cascade of signaling and a decrease in the p21ras/Raf-1 pathway (27). Nitric oxide and cyclic GMP (cGMP) are also involved in the effect of the AT2 receptor in neurite outgrowth and branching, through a pertussis toxin (PTX)-sensitive inhibitory G

2923

2924

Endocrinology, June 2008, 149(6):2923–2933

protein (Gi) (28). On the other hand, the sustained activation of p42/p44mapk and corresponding neurite outgrowth (29) by the AT2 receptor are mediated by phosphorylation of the nerve growth factor (NGF) tropomyosin-related kinase A receptor (30). During migration and neurite outgrowth, cells are characterized by advancing, retracting, turning, and branching behavioral patterns. In neurite outgrowth and elongation, the long-term activation of the AT2 receptor appears to converge on microtubules (16, 18, 19). However, dynamics are also driven by actin cytoskeleton, and reorganization of the actin and microtubular cytoskeleton. In particular, during the process of migration, actin filaments play a major role and may be considered as the primary target of guidance cues because they are localized at the cell periphery and in filopodium in the growth cone (31). Indeed, actin polymerization at the leading edge of migrating cells (32) or protruding growth cones is considered to be the driving force for the forward extension of the cell membrane (33, 34). However, the implication of the actin cytoskeleton and associated molecules in AT2-induced migration of neuronal cells has yet to be investigated. Therefore, we explored the possibility that Ang II, through its AT2 receptor, may modulate the migration of NG108 –15 cells by altering the dynamics of the actin cytoskeleton. In addition, actin-binding proteins of the actin depolymerizing factor (ADF)/cofilin family are important in the regulation of actin-based motility in response to signaling (35, 36). Cofilin is a small 19-kDa protein that binds to both filamentous actin (F-actin) and monomeric actin [globular actin (G-actin)]. Its most important physiological function is to increase actin dynamics by depolymerizing filaments from their pointed ends, whereas the rate of dissociation from the barbed ends is not affected. The dissociation of actin monomers from the filament pointed end is the rate-limiting step of the treadmilling cycle and is increased 25-fold by ADF/ cofilins (37). Overexpression of cofilin enhances cell motility (38), whereas genetic ablation of ADF/cofilin (38) or small interfering RNA (siRNA) knockdown (39) impairs cell motility. Thus, the aim of the present study was to investigate: 1) whether Ang II, through the AT2 receptor, stimulates migration of NG108 –15 cells; 2) how actin cytoskeleton may be involved in this process; and 3) the mechanisms involved in the AT2-receptor mediating effect of actin dynamics. We show here that Ang II increases the rate of migration of NG108 –15 cells in a PTX-sensitive manner and that this acceleration is mediated by an enhanced activity of cofilin. Materials and Methods Chemicals Chemicals were obtained from the following sources: DMEM, fetal bovine serum (FBS), hypoxanthine, aminopterin, thymidine supplement, and gentamycin from Life Technologies, Inc. (Burlington, Ontario, Canada); [Val5]-Angiotensin II from Bachem (Marina Delphen, CA); PD123319 from RBI (Natick, MA); CGP42112 from Ciba-Geigy (Basel, Switzerland); DUP753 from DuPont Merck Pharmaceutical (Paris, France); Hibernate-A medium from BrainBits (Springfield, IL); antiactin antibody clone MAB1501R and anti-tubulin antibody from CHEMICON International, Inc. (Temecula, CA); PD98059 and total p42/p44mapk antibody from New England Biolabs (Beverly, MA); okadaic acid (OA) and

Kilian et al. • AT2 Receptor Increases the Rate of Cell Migration

genistein from Calbiochem VWR Canlab (Mississauga, Ontario, Canada); NG-nitro-l-arginine methyl ester (L-NAME), PTX, p-nitrophenyl phosphate (pNPP), fibronectin (FN) from bovine plasma, and anticofilin antibody clone C8736 from Sigma-Aldrich Canada Ltd. (Oakville, Ontario, Canada); and anti-PP2A (4B7) was from Cell Signaling Technology, Inc. (Danvers, MA). Alexa Fluor-546 phalloidin, DNase I Alexa Fluor 488, 4⬘, 6⬘-diamino-2-phenylindole, and secondary conjugated anti-IgG antibody coupled with Alexa Fluor 488 nm were from Molecular Probes, Inc. (Eugene, OR), and Vectashield mounting medium was from Vector Laboratories (Burlington, Ontario, Canada). Anti-phosphoADF/cofilin was a generous gift from Professor James R. Bamburg (Colorado State University, Fort Collins, CO). Protein G-Sepharose 4 fast Flow beads, horseradish peroxidase-conjugated antirabbit, and antimouse antibodies were from GE Healthcare (Oakville, Ontario, Canada). Complete protease inhibitor, polyvinylidene difluoride membranes, and the enhanced chemiluminescence detection system were from Roche Molecular Biochemicals (Laval, Quebec, Canada). The RNAqueous4PCR system and DNase I were from Ambion, Inc. (Austin, TX). Oligo(deoxythymidine)15 (oligo(dT)15) primers, rRNasin Ribonuclease inhibitor, and Moloney murine leukemia virus reverse transcriptase were from Promega Corp. (Madison, WI). Deoxynucleotide triphosphates were from Amersham Pharmacia Biotech (Oakville, Ontario, Canada). All other chemicals were of grade A purity.

Cell culture Mouse neuroblastoma X rat glioma NG108 –15 hybrid cells were cultured (passage 18 –24) in DMEM high glucose with 10% FBS, hypoxanthine, aminopterin, thymidine supplement, and 50 mg/liter gentamycin plus l-glutamine at 37 C in 25 cm2 Nunclon ␦ flasks (VWR Intl., Quebec, Canada) in a humidified atmosphere of 93% air and 7% CO2 as previously described (15, 18). The medium was changed every 2 d. Subcultures were performed at subconfluency. Under these conditions, cells mainly express the AT2 receptor subtype (18). For cell migration experiments, the cells were plated onto 22-mm coverslips (50,000 cells per dish) coated with 25 ␮g/ml FN and cultured under the same conditions as previously described (18) for 1 or 2 d before use. This number of cells yields the optimum density for tracking individual cells. Higher densities make single-cell tracking difficult because cells tend to form agglomerates.

Cell dynamics experiments Because experiments were performed for up to 4 h, the culture medium was changed for a CO2 independent medium, Hibernate-A. The coverslip with the adhered cells was transferred to a Peltier element cell culture chamber containing 1 ml fresh Hibernate-A medium supplemented with 10% of FBS. During the experiments, the cells were maintained at approximately 35 C. To avoid evaporation of the culture medium, pure mineral oil (⬃250 ␮l) was used to create a thin oil film on the surface of the medium in the chamber. This film also kept the cells free of bacterial contamination during the experiment. Cell locomotion was recorded during 3– 4 h, with drugs added at midpoint, and the behavior of individual cells compared before and after drug addition. In this manner, the first half of every experiment represents a control for the given stimulation. For image acquisition, a Nikon Eclipse TE300 microscope (Nikon, Mississauga, Ontario, Canada) equipped with CoolSNAP fx digital camera (Roper Scientific, Tucson, AZ) connected to a personal computer running MetaMorph version 5.0 software (Universal Imaging Corp., West Chester, PA) was used. Images were acquired at 30-sec intervals. The final time-lapse video recordings were stored locally on a personal computer hard disk, and the trajectory of individual cells was analyzed using the TrackObject feature of the MetaMorph software.

G- and F-actin preparations To measure monomeric G-actin, cells were briefly washed with icecold PBS, placed on ice, and harvested into Triton lysis buffer containing protease inhibitors [50 mm Tris (pH 7.5), 150 mm NaCl, 1% wt/vol Triton X-100, Complete mixture of inhibitors]. After 10 min, the lysates were centrifuged at 4 C for 10 min at 15,000 ⫻ g to separate the Triton soluble fraction corresponding to monomeric G-actin from the insoluble fraction


containing F-actin filaments (40). The supernatant was used to measure protein content. To extract phospho-ADF/cofilin and cofilin, cells were rinsed with ice-cold PBS and scraped into sodium dodecyl sulfate (SDS) lysis buffer containing phosphatase inhibitors [2% SDS, 10 mm Tris (pH 7.5), 2 mm EGTA, 5 mm dithiothreitol, and 1 mm Na2VO3], transferred to 1.5-ml microcentrifuge tubes, and boiled immediately for 5 min. To decrease solution viscosity, the samples were sonicated to shear genomic DNA and centrifuged 5 min at 15,000 ⫻ g.

Immunofluorescence studies Cytoskeleton immunofluorescence studies were performed as previously described (41, 42). Cells were plated on 22-mm coverslips coated with FN and stimulated with 10 nm CGP42112 for 15 min. Cells were fixed with 3.7% (vol/vol) formaldehyde in Hanks’ balanced salt solution buffer for 15 min at 4 C, and permeabilized for 10 min with 0.2% Triton X-100/Hanks’ balanced salt solution. Cells were then incubated for 60 min at room temperature with Alexa Fluor-546 phalloidin (1:60) (for visualization of microfilaments) or with antitubulin (0.25 ␮g/ml) for 60 min at room temperature. After washes, cells were further incubated for 60 min at room temperature with a secondary conjugated anti-IgG antibody coupled with Alexa Fluor 488 nm. For G-/F-actin assays, cells were incubated with DNase I Alexa Fluor 488 (1:2000) (G-actin) and Alexa Fluor-546 phalloidin (1:60 dilution) for 20 min at room temperature. Dnase I was used to detect and measure unpolymerized actin; this enzyme binds to G-actin with an affinity of about 50 nm. Cells were then incubated with 4⬘, 6⬘-diamino-2-phenylindole (1:300) for 5 min to visualize nuclei. After washes, cells were mounted in Vectashield mounting medium and images acquired with an ORCA-ER digital camera (Hamamatsu, Bridgewater, NJ) mounted on a Nikon Eclipse TE-2000 inverted microscope equipped for epi-illumination. Images were acquired using a 100⫻ objective. Each staining was acquired as separate images using the proper filter set, and care was taken to maintain the exact same exposure time and camera settings for individual color channels during the entire experiment. To calculate the G-/F-actin ratio, an area was traced around each individual cell, and the average intensity of the green (G-actin) and red (F-actin) channel was measured and the ratio expressed in arbitrary units as mean ⫾ se.


2925

RT-PCR analysis Total RNA was extracted, treated with deoxyribonuclease I (to digest contaminating genomic DNA), and reverse transcribed as previously described (45). For standard qualitative RT-PCR, the annealing temperature was 55 C for rat cofilin-1 (Cfl1) primers (QIAGEN, Mississauga, Canada) and 57 C for rat glyceraldehyde-3-phosphate dehydrogenase (GAPDH) primers (46). Sizes of amplicons obtained were 112 bp for rat Cfl1 and 176 bp for rat GAPDH.

Cell transfection with Cfl1 siRNAs Because the mouse Cfl1 gene is less expressed than the rat Cfl1 gene in the rat/mouse NG108 –15 hybrid cell line (data not shown), predesigned siRNA duplexes directed against rat Cfl1 (Rn_Cfl1_2 and Rn_Cfl1_3) were purchased from QIAGEN. As indicated by QIAGEN, these rat Cfl1 siRNAs have 100% homology with the mouse Cfl1 gene and may thus also repress mouse Cfl1 gene expression. The sequence of these commercial siRNAs duplexes is not of public access. Transfection with Cfl1 siRNAs or nonspecific control (scrambled) siRNA (50 nm; QIAGEN) was performed with 5 ␮l Lipofectamine 2000 (Invitrogen, Carlsbad, CA) according to the manufacturer’s recommendations in NG108 –15 cells at 60% confluence. Gene silencing was confirmed 24 h after transfection by RT-PCR and Western blot analysis. Cotransfection of Cfl1 siRNA and pEYFP-c1 (Clontech, Mountain View, CA) in NG108 –15 cells yielded a transfection efficiency of more than 80% for pEYFP-c1 (data not shown).

Data analysis The trajectory data of each cell obtained with the TrackObject feature of the MetaMorph software were transferred to Microsoft Excel (Mi-

Phosphatase activity Phosphatase assay was performed as described by Yusa and Campbell (43). Briefly, cells were treated at indicated times with 100 nm Ang II, and lysis was performed with lysis buffer [100 mm Tris (pH 7.5), 150 mm NaCl, 1% Igepal (Sigma-Aldrich Canada Ltd.), and Complete protease inhibitor]. After centrifugation, 350 ␮g proteins were incubated with agitation for 2 h at 4 C with 2 ␮g anti-PP2A in 1 ml total volume of lysis buffer. Thereafter, 20 ␮l protein G-Sepharose beads were added for 1 h at 4 C under agitation. Beads were washed three times in washing buffer [100 mm Tris (pH 7.5), 150 mm NaCl, and 0.1% Igepal] and three times with washing buffer without Igepal [50 mm Tris (pH 7.5), 150 mm NaCl]. Beads were then incubated overnight at 37 C in phosphatase buffer [20 mm Tris (pH 7.5), 100 mm NaCl, 1 mm EGTA, 10 mm dithiothreitol, and 12 mm pNPP]. Reaction was stopped with addition of a final concentration of 0.5 n NaOH, and absorbance was read at 410 nm. pNPP is a nonspecific substrate for tyrosine and serine/threonine phosphatase (44).

Western blotting After stimulation with appropriate conditions, cell lysis (see appropriate section for each type of specific extraction) and Western blotting were performed as previously described (25, 28, 42). Equal amounts of protein (12–20 ␮g) were separated on 10% SDS-polyacrylamide gels. Western blotting was performed, using polyvinylidene difluoride membranes. Membranes were blocked with 5% nonfat milk and probed with primary antibodies (antiactin 1:8,000, anticofilin 1:10,000, and anti-phospho-ADF/cofilin 1:5,000). Detection was performed by reaction with horseradish peroxidase-conjugated secondary antibody, and visualization by chemiluminescence with enhanced chemiluminescence system and analyzed by densitometry.

FIG. 1. Morphology and time-lapse imaging of NG108 –15 cell dynamics. A, Time progression of NG108 –15 cells during a 4-h experiment. Only six images at 20-min intervals are presented. The trajectory of one cell is identified by an arrow. B, The migration paths of cells in control condition were traced during a 140-min period. The intersection of the x- and y-axes was considered the starting point for each cell path. Scale bar, 30 ␮m.



crosoft Corp., Redmond, WA), in which the average velocity before and after addition of respective drugs was calculated for individual cells. The differences between the average velocity before and after drug addition were then expressed in percentage, averaged, and reported as a value ⫾ sem for the number of cells obtained from at least three independent experiments. One-way ANOVA with the Tukey post hoc test was used to compare pairwise the experimental data for statistical differences. A value of P ⱕ 0.05 was considered statistically significant.

Results

In control conditions, NG108 –15 cells displayed a relatively round or polygonal cell body, with short protrusions, as previously described by our group (28). As shown in Fig. 1A, these cells exhibited a very high migratory potential, even in control conditions. Using time-lapse video microscopy, cell displacements were recorded by acquiring an im-

number or events

A

B

40

control 100 nM Ang II

30 20 10 0 0

40 80 120 160 average velocity (µm/h)

200

60

*

40

*

30 20 10 0

50

* *

*

40

*

30 20 10

0 Ang II CGP42112 PD123319 DUP753

D

50

CGP42112 L-NAME PD98059 PTX

60

+ -

-

50

velocity change (%)

velocity change (%)

C

age every 30 sec for a period of 4 h. The trajectories are represented as lines and show a random orientation (Fig. 1B). The random walk fashion of cell locomotion was confirmed by fitting the mean squared displacement curve of several cells (n ⫽ 22) to a simple diffusion model (47) (data not shown). The migration speed in control conditions varied from 31.5–110.9 ␮m/h with an average of 72.4 ⫾ 4.1 ␮m/h (n ⫽ 22). Upon addition of 100 nm Ang II to the bath medium, the mean velocity of the cells increased from 72.4 ⫾ 4.1 to 98.5 ⫾ 6.0 ␮m/h, an increase of 36 ⫾ 7% (n ⫽ 22; P ⬍ 0.01). Velocities before and after Ang II treatment displayed a normal distribution as verified by the Kolmogorov-Smirnov normality test (K-S Dist) (K-S Dist ⫽ 0.075, P ⫽ 0.098 vs. K-S Dist ⫽ 0.124 P ⬎ 0.2, control vs. Ang II, respectively) (Fig. 2A). To inves-

velocity change (%)

2926

+ -

*

40

+ + -

+ +

+ +

*

*

30 20 10 0

-

+ -

+ + -

+ + -

+ +

-10

Ang II w/o FBS Gen OA

-

+ -

+ + -

+ + -

+ +

FIG. 2. Modulation of the Ang II-induced migration velocity of NG108 –15 cells. A, Histograms showing velocity distribution in control and Ang II-stimulated cell populations. B, Comparison of velocity changes in cells in control conditions and in cells stimulated for 2 h with 100 nM Ang II, 10 nM CGP42112 (an AT2 receptor agonist), Ang II plus 1 ␮M PD123319 (an AT2 receptor antagonist), Ang II or CGP42112 plus 1 ␮M DUP753 (an AT1 receptor antagonist). C, Putative transduction pathways implicated in AT2-receptor-mediated acceleration. Cells were stimulated with CGP42112 while preincubated, or not, with pharmacological inhibitors of key signaling pathways, namely L-NAME (0.2 mM, 10-min preincubation), PD98059 (10 ␮M, 10-min preincubation), and PTX (100 ng/ml, overnight preincubation). D, Role of serum-induced phosphorylation in AT2-receptor-mediated cell motility. Cells were stimulated with Ang II in a serum-free medium or in serum-containing medium in the presence of genistein (Gen) (10 ␮M, 10-min preincubation) or OA (10 nM, 60-min preincubation). Values are presented as mean ⫾ SE. *, P ⱕ 0.01, statistical significance compared with control or respective control conditions. w/o, Without.



2927

FIG. 3. Effect of CGP42112 on immunofluorescence labeling of actin filaments and microtubules. NG108 –15 cells were plated on plastic petri dishes (1 ⫻ 105 cells) as described in Materials and Methods and subsequently stimulated without (A–C) (control) or with 10 nM CGP42112 for 15 min (D–F). After formaldehyde fixation and permeabilization with 0.1% Triton X-100, cells were processed for immunofluorescence labeling using phalloidin coupled to Alexa Fluor 546 nm for visualization of F-actin (A and D) (red) and with antitubulin antibody coupled to Alexa Fluor 488 nm for visualization of tubulin (B and E) (green). Merged images are shown in C and F with inset magnifications shown in panels G–J. Images are representative illustrations of more than 20 cells originating from five different experiments. Scale bars, 8 ␮m for panels A–F and 3 ␮m for panels G–J. Arrowheads indicate cortical ring of actin, and arrows the parallel bundles of F-actin in filopodia.

tigate whether this effect was specific to the activation of the AT2 receptor, two strategies were used. First, application of 10 nm CGP42112, a selective agonist of the AT2 receptor, in the bath caused acceleration of the cells by 34 ⫾ 5% (n ⫽ 12; P ⬍ 0.01 when compared with control cells), a value similar to that observed with Ang II (Fig. 2B). Second, when Ang II was applied 10 min after preincubation with 1 ␮m PD123319, a selective AT2 receptor antagonist, no acceleration was observed (2 ⫾ 7%, n ⫽ 9). To exclude further any implication of AT1 receptors in the observed Ang II-induced cell acceleration, cells were preincubated 10 min with DUP753, a selective AT1 receptor antagonist. In these conditions, 100 nm Ang II or 10 nm CGP42112 induced the same effect as Ang II or CGP42112 alone; the cells accelerated by 44 ⫾ 7% (n ⫽ 12) and 45 ⫾ 7% (n ⫽ 19), respectively, a value significantly different when compared with control experiments (P ⬍ 0.01) (Fig. 2B). Previous results from our laboratories have shown that the signaling mechanisms inducing neurite outgrowth by the AT2 receptors in NG108 –15 cells include at least two distinct but complementary pathways involving the PTX-sensitive nitric oxide synthase/nitric oxide/soluble guanylyl cyclase/ cGMP (28), and the Rap1/B-Raf/MAPK kinase (MEK)/p42/ p44mapk cascades (11). Therefore, we investigated whether these pathways could be implicated in the acceleration effect observed upon AT2 receptor stimulation. Neither the inhibition of the nitric oxide synthase with 0.2 mm L-NAME nor the inhibition of MEK activity with 10 ␮m PD98059, could reverse Ang II (data not shown) or CGP42112-induced acceleration (Fig. 2C). Nevertheless, the cascade leading from AT2 receptor to an increase in NG108 –15 cell migration rate involved a PTX-sensitive Gi protein. Indeed, overnight preincubation of NG108 –15 cells with 100 ng/ml PTX abolished

the acceleration induced by either 100 nm Ang II (data not shown) or 10 nm CGP42112 (Fig. 2C). Because PP2A activity was also increased by AT2 receptor activation in primary neuronal cultures in a PTX-sensitive fashion (29, 48), we, therefore, tested PP2A activity after stimulation of NG108 –15 cells with Ang II. After 10 min, relative PP2A activity significantly increased from 1.0 ⫾ 0.1 in control conditions to 2.1 ⫾ 0.4 in 100 nm Ang II-stimulated conditions (n ⫽ 6, P ⫽ 0.026). Recovery to basal level was reached after 60 min (0.94 ⫾ 0.23; n ⫽ 6; P ⫽ 0.832 with control). Preincubation (60 min) of the cells with 10 nm OA, a relatively specific PP2A phosphatase inhibitor at this particular concentration (44, 49), greatly decreased the acceleration induced by 100 nm Ang II (Fig. 2D) (Ang II: 35.6 ⫾ 7.4% n ⫽ 22; OA plus Ang II: 7.5 ⫾ 5.2% n ⫽ 32; P ⬍ 0.01). In addition, because the effects of Ang II on neurite outgrowth occurred in the presence of serum, motility was verified in cells maintained in a serum-free medium during the experimental period. Cell velocity in a serum-containing medium (72.4 ⫾ 4 ␮m/h, n ⫽ 22) was not different (P ⫽ 0.377) from that recorded in a serum-free medium (66.7 ⫾ 4.8 ␮m/h, n ⫽ 23). As shown in Fig. 2D, Ang II did not increase cell velocity in serum-free conditions (Ang II: 35.6 ⫾ 7.4% n ⫽ 22; Ang II serum-free: ⫺0.8 ⫾ 5% n ⫽ 23; P ⬍ 0.01). Furthermore, in a serum medium containing 10 ␮m genistein (a tyrosine kinase inhibitor), the acceleration induced by 100 nm Ang II was practically abolished (Ang II: 35.6 ⫾ 7.4% n ⫽ 22; genistein plus Ang II: 17.3 ⫾ 6.3% n ⫽ 32; P ⬍ 0.01). Actin filaments are known to play a central part in cell motility (31), involving cycles of net assembly of actin filaments at the leading edge, retrograde movement of actin networks, and disassembly of filament proximally (33, 50, 51). Thus, the actin cytoskeleton is a potential candidate for

2928



FIG. 4. Effect of CGP42112 on immunofluorescence labeling of F- and G-actin. A, NG108 –15 cells were plated on plastic petri dishes (1 ⫻ 105 cells) as described in Materials and Methods and subsequently stimulated without (Aa–Ac) (control) or with 10 nM CGP42112 for 15 min (Ad–Af). After formaldehyde fixation and permeabilization with 0.1% Triton X-100, cells were processed for immunofluorescence labeling using Alexa Fluor-546 phalloidin (1:60 dilution) for visualization of F-actin (Aa and Ad) (red) and with DNase I Alexa Fluor 488 (1:2000) for visualization of G-actin (Ab and Ae) (green). Merged images are shown in Ac and Af with inset magnifications shown in panels Ag–Aj. Images are representative illustrations of more than 398 cells originating from 10 different experiments. Scale bars, 8 ␮m for panels Aa–Af and 3 ␮m for panels Ag–Aj. B, G-/F-actin ratio in central and CGP4212 stimulated cells. *, P ⱕ 0.01, statistical significance compared with control.

the convergence of signals resulting from the activation of the AT2 receptor, leading to acceleration of NG108 –15 cellular migration. In control conditions, actin appeared as straight actin filaments (F-actin) forming a cortical ring in proximity to the cell membrane (arrowheads) and parallel bundles in membrane filopodia (arrows) (Fig. 3, A, C, G, and I). In contrast, microtubules were loosely distributed throughout the cytoplasm (Fig. 3, B, C, E, and F), occasionally interacting with F-actin at the cell periphery, as evidenced by the yellow color in overlay images (Fig. 3, C, G, and I). After a 15-min stimulation with 10 nm CGP42112 (Fig. 3, D–F), the number of actin filaments in filopodia decreased along with the disappearance of the submembrane labeling as well as the colocalization with tubulin (Fig. 3, H and J compared with G and I). It is traditionally well accepted that the actin pool within a cell is relatively constant. To verify whether activation of AT2 receptors altered the balance between F-actin vs. G-actin

through an increase in the unpolymerized actin pool, two distinct fluorescent ligands were used, namely DNaseI conjugated to Alexa Fluor 488 for labeling G-actin and phalloidin coupled to Alexa Fluor 546 to identify F-actin. As previously shown, control cells exhibited F-actin as a ring adjacent to the cell membrane and as short stress fibers in filopodia (Fig. 4, A, C, G, and I), whereas G-actin (green) appeared diffusely distributed throughout the cytoplasm (Fig. 4, B and C). After a 15-min stimulation with 10 nm CGP42112, F-actin labeling (red) decreased, whereas the level of G-actin increased, mainly at the periphery of the cell (Fig. 4, D–F, H, and J). These observations revealed a shift from F-actin to G-actin, as indicated by quantification of the G-/F-actin ratio (1.43 ⫾ 0.009, n ⫽ 398 in control cells compared with 1.38 ⫾ 0.009, n ⫽ 398 in CGP42112-treated cells; P ⱕ 0.001) (Fig. 4B). These results were further corroborated by Western blotting. Stimulation of cells with CGP42112 increased the level of unpolymerized G-actin. This increase was observed as


A


2929

10 nM CGP42112 0

1

3

5

10

15

30

60

min

actin G actin F

B

100 nM PTX 10 nM CGP42112 0

1

3

5

10

15

30

60

min

actin G actin F

Amount of G-actin (%)

C

350 PTX without PTX

300 250 200

FIG. 6. Effect of AT2 receptor stimulation on ADF/cofilin expression. A, Cells were stimulated with 100 nM Ang II for the indicated time periods. Representative Western blots show the P-ADF/cofilin and total ADF/cofilin after activation of the AT2 receptor. B, Cells were stimulated with 10 nM CGP42112 for the indicated time periods. C, Cells were preincubated overnight with 100 ng/ml PTX and stimulated with 100 nM Ang II for the indicated time periods.

150 100 50 0 0

10

20

30

40

50

60

Duration of stimulation (min) FIG. 5. Effect of AT2 receptor stimulation on G-actin accumulation. A, Western blots of G-actin content in NG108 –15 cells stimulated with 10 nM CGP42112 for the indicated time periods. B, Western blots of G-actin content in cells preincubated overnight with 100 ng/ml PTX and stimulated with 10 nM CGP42112 for the indicated time periods. C, Time course of G-actin accumulation after CGP42112 stimulation. Comparative densitometric analyses of experimental data showing the mean ⫾ SE of three independent experiments, expressed as fold increase over control or PTX pretreatment at time zero.

early as 1 min after stimulation (150% of control nonstimulated cells), with a maximal response at 15 min (260% of control nonstimulated cells) and sustained for at least 1 h (Fig. 5, A and C). The involvement of a PTX-sensitive G protein was observed in this effect because preincubation of NG108 –15 cells with 100 ng/ml PTX abolished actin depolymerization induced by 10 nm CGP42112 (Fig. 5, B and C). Actin depolymerization is also known to be regulated by the ADF/cofilin protein family. Therefore, we verified the hypothesis that ADF/cofilin activity could be modulated by AT2 receptor activation. Stimulation of the AT2 receptor with 100 nm Ang II (Fig. 6A) or 10 nm CGP42112 (Fig. 6B) decreased the level of phosphorylated ADF (P-ADF)/cofilin, an effect measurable after 10 min. Overnight preincubation of NG108 –15 cells with 100 ng/ml PTX abolished the P-ADF/ cofilin response induced by 100 nm Ang II (Fig. 6C). The role of cofilin in the acceleration induced by Ang II was further assessed using NG108 –15 cells in which expression of Cfl1

was suppressed by siRNA plasmid transfection. Cfl1 expression, at both mRNA (Fig. 7A) and protein (Fig. 7B) levels, was markedly blunted in Cfl1 siRNA-transfected cells. Using the predesigned siRNA duplexes directed against rat Cfl1 (Rn_Cfl1_2 and Rn_Cfl1_3), Fig. 7C shows that the expression of Cfl1 was significantly decreased in both conditions (44.9 ⫾ 5.6%, n ⫽ 3, P ⬍ 0.01 and 40.7 ⫾ 9.8%, n ⫽ 3, P ⬍ 0.01, compared with the control value). The consequence of Cfl1 blunting was subsequently studied on cell migration using Cfl1-siRNA-transfected cells (Cfl1_3). Cfl1 siRNA-transfected cells were rounded or polygonal without neurites, and their displacement was restricted, if any (Fig. 8A). When analyzed over the same time-lapse period, Cfl1 siRNA-transfected cells covered a shorter distance compared with scramble-transfected cells (Fig. 8, B and C). This reflected a slower migration speed that was reduced from 80.03 ⫾ 4.2 ␮m/h in cells transfected with a scramble Cfl1 siRNA (n ⫽ 9) to 16.5 ⫾ 3.3 ␮m/h in cells transfected with Cfl1 siRNA (n ⫽ 12) (Fig. 8C), the difference between the two groups being statistically significant (P ⬍ 0.001) (Fig. 8E). In cells transfected with a scramble Cfl1 siRNA, the migration speed (80.03 ⫾ 4.18 ␮m/h; n ⫽ 9) was not significantly affected (P ⫽ 0.283) compared with control nontransfected cells (72.4 ⫾ 4.1 ␮m/h; n ⫽ 22) (Fig. 8 B compared with Fig. 1B). As observed in control cells, velocities of Cfl1 siRNA-transfected cells displayed a normal distribution (Fig. 8D) as verified by the K-S Dist (0.18; P ⬎ 0.2). In the process of migration, cells also frequently change direction; this behavior can be quantified by the J index, expressed as the ratio of the net translocation


distance over the cumulative length of migration. The J index of Cfl1 siRNA-transfected cells (0.12 ⫾ 0.027, n ⫽ 12) was significantly lower (P ⬍ 0.001) than that of control cells (0.44 ⫾ 0.03, n ⫽ 28). Addition of 100 nm Ang II to Cfl1 siRNA-transfected cells did not affect the speed of migration (17.8 ⫾ 5.8 ␮m/h, n ⫽ 6; P ⫽ 0.836) compared with siRNAtransfected cells in control medium (Fig. 8E). Discussion

In the present study, we demonstrate that activation of the AT2 receptor by Ang II or CGP42112 increases the migration velocities of NG108 –15 cells. Results indicate that Ang II decreases the amount of F-actin in filopodium and increases the pool of unpolymerized actin, through a PTX-sensitive increase in ADF/cofilin activity. Thus, these results are in agreement with previous observations whereby the AT2 receptor was shown to enhance neuronal cell migration in microexplant cultures of the rat cerebellum (19). The effects of Ang II are mediated by the type 2 receptor, the predominant subtype present in nondifferentiated NG108 –15 cells (10, 18). This is supported by the observations that the increase in the rate of migration was abolished by PD123319, a selective AT2 receptor antagonist, but not by DUP753, a selective AT1 receptor antagonist. Moreover, these Ang II effects were able to be reproduced using CGP42112, a selective AT2 receptor agonist. Previous studies from our laboratories have shown that Ang II mediates the morphological neuronal differentiation of NG108 –15 cells through at least two independent signaling pathways, namely nitric oxide synthase/nitric oxide/soluble guanylyl cyclase/cGMP (PTX sensitive), and Rap1/B-Raf/ MEK/p42/p44mapk cascade of signaling (27). None of these well-characterized cascades, except for PTX sensitivity, appears to be involved in the Ang II-induced acceleration of NG108 –15 cell movement. Indeed, neither blocking of nitric oxide synthase with L-NAME nor inhibition of the MAPK pathway at the MEK level with PD98059 could reverse the effect of AT2 receptor activation on the rate of NG108 –15 cell migration. Several studies clearly place a central role of actin in cell motility (31). Indeed, advancing, retracting, turning, and branching are all regulated by reorganization and dynamics of both actin and microtubule cytoskeleton. This involves cycles of net assembly and disassembly of actin filaments (33, 50 –52). Here, we confirm that in NG108 –15 cells, the process of AT2receptor induced cell acceleration of migration affects these dynamics. This was clearly illustrated in immunofluorescence studies. First, we show that in control cells, F-actin is abundant, appearing as spikes in filopodia at the leading edge of cells, and that Ang II or CGP42112 decreases the amount of F-actin at the leading edge of the cells. This decrease in actin filament labeling is accompanied by a concomitant increase in the level of G-actin, demonstrated both by immunofluorescence and Western blotting. In addition, there is also some colocalization of F-actin with microtubules, corroborating recent observations, which disappear during Ang II-induced migration. It has been suggested that such actin-microtubule interactions may modulate motility (31). Among the candidate molecules that possibly cross-link actin filaments and microtubules are microtubule-associated proteins (MAPs), such as MAP2c and MAP1B (53, 54), proteins


that we have previously shown to be affected during the process of neurite outgrowth stimulated by the AT2 receptor, both in NG108 –15 cells and in cerebellar granule cells (18, 19). Regulation of actin turnover in actin-based motile systems is mainly under the control of the ADF and cofilin (36, 55). To assess the role of cofilin in the acceleration process induced by Ang II, expression of Cfl1 was suppressed by transfection of siRNA plasmids. Migration activity was profoundly affected in Cfl1 siRNA NG108 –15 cells, as shown by a 77.2% decrease in migration speed. Similar results have also been obtained in several other cell types, including yeast, Drosophila melanogaster, and mammalian cells (38, 39, 56, 57). Moreover, the migration velocity of Cfl1 siRNA cells was no longer affected by AT2 cofilin 1

A

2

3

4 112 bp

1

2

GAPDH

3

4 176 bp

B

cofilin 19 kDa MAPK 44 kDa 42 kDa

C 100

Cofilin levels (% of control)

2930

80 60

**

40

**

20 0

Control Scramble Cfl_2

Cfl_3

FIG. 7. Effect of siRNA-induced gene silencing of cofilin on NG108 –15 cell motility. A, PCR analysis of mRNA cofilin levels in NG108 –15 cells from control, scramble siRNA-, Cfl_2 siRNA-, and Cfl_3 siRNA-transfected cells, for lanes 1– 4, respectively. B, Western blot analysis of cofilin levels in NG108 –15 cells from control, scramble siRNA-, Cfl_2 siRNA-, and Cfl_3 siRNA-transfected cells, for lanes 1– 4 in A, respectively. Cell lysates containing equal concentrations of protein were subjected to Western blot analyses with antibodies against cofilin-protein (upper panels). Lower panels represent the same blots reprobed for total p42/p44mapk. C, Densitometry analysis of cofilin levels in control and siRNA-transfected cells. Note that the cofilin/p42/p44mapk ratio was more strongly reduced in siRNA-transfected cells, although the expression of the p42/p44mapk was also attenuated. **, P ⬍ 0.001, statistical significance compared with control or respective control conditions.


A


0 min

40 min

80 min

120 min

160 min

200 min

B

C

-90

-60

-30

80

80

40

40

0

0

30

-40

60

m

-150 -120

m

m

-150 -120

-90

-60

-30

0

0

-40

-80

-80

-120

-120

-160

-160

30

m

60

E

D

100

4

Number of events

2931

80

3

60

2 40

1 0

**

20

0

10

20

30

40

50

0

Scramble

**

siRNA siRNA+Ang II

Velocity ( m/h) FIG. 8. Effect of cofilin siRNA on NG108 –15 cell migration. A, Phase-contrast morphology of Cfl_3 siRNA-transfected cells illustrated over a 200-min period. B, Migration paths of cells in control (scramble) condition were traced for 140 min. The intersection of the x- and y-axes was considered the starting point for each cell path. C, Migration paths of cofilin siRNA-transfected cells. D, Histogram showing velocity distribution in the population of cofilin siRNA-transfected cells. E, Mean velocity in control cells, cofilin siRNA cells, and cofilin siRNA cells after addition of 100 nM Ang II in the bath. Mean cell velocity was not significantly different between cofilin siRNA cells before and after Ang II addition. **, P ⬍ 0.001, statistical significance compared with control. Scale bar, 30 ␮m.

receptor stimulation, which could infer the role of cofilin in Ang II-induced acceleration of NG108 –15 cells. Cofilin, like ADF [these two proteins having overlapping roles (57)], plays a critical role in actin polymerization by severing F-actin and exposing free barbed ends for polymerization (58). A second role, different from that of an actin-depolymerizing factor, was also

recently demonstrated for cofilin. Indeed, by using caged cofilin, Ghosh et al. (58) showed that protrusions were formed at the site of uncaging and that these protrusions determined the direction of cell migration. Moreover, cofilin knockdown cells are round, flat, and have lost their polarity, indicating that cofilin is required to set the direction of cell migration (57).

2932


Directionality can be quantified by the J index: low values of J indicating a random motion, whereas higher values highlight a more directed path (59). In the present study, the J index for Cfl1 siRNA NG108 –15 cells was decreased by 72.7% compared with control cells, therefore, indicating a loss of directional migration. The question remains as to what are the mechanisms by which AT2 receptor is coupled to ADF/cofilin dephosphorylation. Growth factors such as NGF and insulin induce a rapid dephosphorylation of ADF/cofilin in PC12 cells (60). Based on the dual observations that a serum-containing medium is required for the effect of AT2 receptor (61) and that the AT2 receptor-induced sustained activation of p42/p44mapk is mediated by phosphorylation of the tropomyosin-related kinase A receptor of NGF (30), we propose that dephosphorylation of ADF/cofilin by Ang II binding to the AT2 receptor could involve a tyrosine kinase activity. Indeed, we demonstrate here that the Ang II-induced increase in cell velocity is abrogated in a serum-free medium or by genistein in a serum-containing medium. Dephosphorylation of ADF/cofilin may occur by more than one class of phosphatases, depending on cell type and the signaling pathway that is activated (60). These include the PP family of phosphatases (PP1, PP2A, PP2B, and PP2C) (60, 62, 63), Slingshot family phosphatases (64, 65), and chronophin (66). PP2A phosphatase is indeed linked to AT2 receptor signaling in NG108 –15 cells in a PTX-sensitive manner (29, 48). In the present study, PP2A activity was found increased by Ang II, and preincubation with OA (10 nm, 60 min) abolished the acceleration induced by 100 nm Ang II. This indicates that PP2A phosphatase is most likely involved in the signaling pathway of the AT2 receptor in modulating cell migration. Whether PP2A acts directly or indirectly on P-ADF/cofilin is not known in NG108 –15 cells. Indeed, stimulation of PC12 cells with growth factors has revealed that ADF/cofilin is dephosphorylated by PP1, but not PP2A (60). In conclusion, a central role for AT2 receptors in neuronal development has long been suspected given that the expression of the AT2 receptor in the central nervous system peaks during the developmental period and decreases drastically after birth and in the adult (9, 67). We have indeed previously shown that Ang II, through AT2 receptor activation, induces neurite elongation both in NG108 –15 cells and in primary cultures of neuronal cerebellar granule cells. Here, we demonstrate that shortterm stimulation of the AT2 receptor increases the rate of NG108 –15 cell migration by affecting actin polymerization, thereby thus indicating a new role for this receptor in actin dynamics. Together, the present studies further document that Ang II function, via its AT2 receptor, plays a key role in two distinct phases of neuronal differentiation, namely migration and neurite outgrowth. Thus, as described for serotonin (68) and for pituitary adenylyl cyclase activating peptide (69), Ang II, through the AT2 receptor, could be considered as a guidance cue (70) in promoting neurite outgrowth and cytoskeletal dynamics. Furthermore, the role of cofilin and actin cytoskeleton in neuronal migration disorders has recently been outlined in vivo through the use of mouse mutants for cofilin and ADF, whereby mutations affecting actin filament formation were shown to contribute to cell cycle and neuronal migration defects in the cerebral cortex (71).


Acknowledgments We thank Professor James R. Bamburg (Colorado State University, Fort Collins, CO) for the generous gift of the anti-phospho-actin depolymerizing factor/cofilin antibody. Received March 6, 2007. Accepted February 25, 2008. Address all correspondence and requests for reprints to: Marcel Daniel Payet, Ph.D., De´partement de Physiologie et Biophysique, Faculte´ de Me´decine et des Sciences de la Sante´, Universite´ de Sherbrooke 3001, 12e Avenue Nord, Sherbrooke, Que´bec, Canada J1H 5N4. E-mail: [email protected]. This work was supported by grants from the Canadian Institute for Health Research (to N.G.-P. and M.D.P.) (MOP27912). P.K. is the recipient of a Ph.D. studentship “Bourse d’Excellence” from the Ministe`re de l’E´ducation du Que´bec. N.G.-P. is the recipient of a Canada Research Chair in Endocrinology of the Adrenal Gland. Present address for S.C.: Department of Pharmacology and the Vermont Cancer Center, University of Vermont, E-213 Given Building, 89 Beaumont Avenue, Burlington, Vermont 05405. Disclosure Statement: The authors have nothing to declare.

References 1. Inagami T, Kambayashi Y, Ichiki T, Tsuzuki S, Eguchi S, Yamakawa T 1999 Angiotensin receptors: molecular biology and signalling. Clin Exp Pharmacol Physiol 26:544 –549 2. de Gasparo M, Catt KJ, Inagami T, Wright JW, Unger T 2000 International union of pharmacology. XXIII. The angiotensin II receptors. Pharmacol Rev 52:415– 472 3. Horiuchi M, Lehtonen JY, Daviet L 1999 Signaling mechanism of the AT2 angiotensin II receptor: crosstalk between AT1 and AT2 receptors in cell growth. Trends Endocrinol Metab 10:391–396 4. Stoll M, Unger T 2001 Angiotensin and its AT2 receptor: new insights into an old system. Regul Pept 99:175–182 5. Volpe M, Musumeci B, De Paolis P, Savoia C, Morganti A 2003 Angiotensin II AT2 receptor subtype: an uprising frontier in cardiovascular disease? J Hypertens 21:1429 –1443 6. Reudelhuber T 2005 The continuing saga of the AT2 receptor: a case of the good, the bad, and the innocuous. Hypertension 46:1261–1262 7. Millan MA, Jacobowitz DM, Aguilera G, Catt KJ 1991 Differential distribution of AT1 and AT2 angiotensin II receptor subtypes in the rat brain during development. Proc Natl Acad Sci USA 88:11440 –11444 8. Tsutsumi K, Saavedra JM 1991 Characterization of AT2 angiotensin II receptors in rat anterior cerebral arteries. Am J Physiol 261(3 Pt 2):H667–H670 9. Nuyt AM, Lenkei Z, Palkovits M, Corvol P, Llorens-Cortes C 1999 Ontogeny of angiotensin II type 2 receptor mRNA expression in fetal and neonatal rat brain. J Comp Neurol 407:193–206 10. Bottari SP, Obermuller N, Bogdal Y, Zahs KR, Deschepper CF 1992 Characterization and distribution of angiotensin II binding sites in fetal and neonatal astrocytes from different rat brain regions. Brain Res 585:372–376 11. Gendron L, Payet MD, Gallo-Payet N 2003 The angiotensin type 2 receptor of angiotensin II and neuronal differentiation: from observations to mechanisms. J Mol Endocrinol 31:359 –372 12. Lucius R, Gallinat S, Busche S, Rosenstiel P, Unger T 1999 Beyond blood pressure: new roles for angiotensin II. Cell Mol Life Sci 56:1008 –1019 13. Speth RC, Mei L, Yamamura HI 1989 Angiotensin II receptor binding and actions in NG108 –15 cells. Pept Res 2:232–239 14. Tallant EA, Diz DI, Khosla MC, Ferrario CM 1991 Identification and regulation of angiotensin II receptors subtypes on NG108 –15 cells. Hypertension 17:1135–1143 15. Buisson B, Bottari SP, De Gasparo M, Gallo-Payet N, Payet M-D 1992 The angiotensin AT2 receptor modulates T-type calcium current in non-differentiated NG108 –15 cells. FEBS Lett 309:161–164 16. Meffert S, Stoll M, Steckelings UM, Bottari SP, Unger T 1996 The angiotensin II AT2 receptor inhibits proliferation and promotes differentiation in PC12W cells. Mol Cell Endocrinol 122:59 – 67 17. Nahmias C, Cazaubon SM, Briend-Sutren MN, Lazard D, Villageois P, Strosberg AD 1995 Angiotensin AT2 receptors are functionally coupled to protein tyrosine dephosphorylation in N1E-115 neuroblastoma cells. Biochem J 306(Pt 1):87–92 18. Laflamme L, Gasparo M, Gallo JM, Payet MD, Gallo-Payet N 1996 Angiotensin II induction of neurite outgrowth by AT2 receptors in NG108 –15 cells. Effect counteracted by the AT1 receptors. J Biol Chem 271:22729 –22735 19. Coˆte´ F, Do TH, Laflamme L, Gallo JM, Gallo-Payet N 1999 Activation of the AT(2) receptor of angiotensin II induces neurite outgrowth and cell migration in microexplant cultures of the cerebellum. J Biol Chem 274:31686 –31692 20. Lucius R, Gallinat S, Rosenstiel P, Herdegen T, Sievers J, Unger T 1998 The


21.

22. 23.

24. 25.

26. 27. 28. 29. 30.

31. 32. 33. 34. 35. 36. 37.

38. 39. 40. 41.

42. 43. 44. 45.

angiotensin II type 2 (AT2) receptor promotes axonal regeneration in the optic nerve of adult rats. J Exp Med 188:661– 670 Reinecke K, Lucius R, Reinecke A, Rickert U, Herdegen T, Unger T 2003 Angiotensin II accelerates functional recovery in the rat sciatic nerve in vivo: role of the AT2 receptor and the transcription factor NF-␬B. FASEB J 17:2094 – 2096 Nouet S, Nahmias C 2000 Signal transduction from the angiotensin II AT2 receptor. Trends Endocrinol Metab 11:1– 6 Senbonmatsu T, Saito T, Landon EJ, Watanabe O, Price Jr E, Roberts RL, Imboden H, Fitzgerald TG, Gaffrey FA, Inagami T 2003 A novel angiotensin II type 2 receptor signaling pathway: possible role in cardiac hypertrophy. EMBO J 22:6471– 6482 Landon EJ, Inagami T 2005 Beyond the G protein: the saga of the type 2 angiotensin II receptor. Arterioscler Thromb Vasc Biol 25:15–16 Gendron L, Laflamme L, Rivard N, Asselin C, Payet MD, Gallo-Payet N 1999 Signals from the AT2 (angiotensin type 2) receptor of angiotensin II inhibit p21ras and activate MAPK (mitogen-activated protein kinase) to induce morphological neuronal differentiation in NG108 –15 cells. Mol Endocrinol 13: 1615–1626 Stroth U, Blume A, Mielke K, Unger T 2000 Angiotensin AT(2) receptor stimulates ERK1 and ERK2 in quiescent but inhibits ERK in NGF-stimulated PC12W cells. Brain Res Mol Brain Res 78:175–180 Gendron L, Oligny J, Payet M, Gallo-Payet N 2003 Cyclic AMP-independent involvement of Rap1/B-Raf in the angiotensin II AT2 receptor signaling pathway in NG108 –15 cells. J Biol Chem 278:3606 –3614 Gendron L, Cote F, Payet MD, Gallo-Payet N 2002 Nitric oxide and cyclic GMP are involved in angiotensin II AT(2) receptor effects on neurite outgrowth in NG108 –15 cells. Neuroendocrinology 75:70 – 81 Huang XC, Richards EM, Sumners C 1995 Angiotensin II type 2 receptormediated stimulation of protein phosphatase 2A in rat hypothalamic/brainstem neuronal cocultures. J Neurochem 65:2131–2137 Plouffe B, Guimond MO, Beaudry H, Gallo-Payet N 2006 Role of tyrosine kinase receptors in angiotensin II AT2 receptor signaling: involvement in neurite outgrowth and in p42/p44mapk activation in NG108 –15 cells. Endocrinology 147:4646 – 4654 Kalil K, Dent EW 2005 Touch and go: guidance cues signal to the growth cone cytoskeleton. Curr Opin Neurobiol 15:521–526 Meyer G, Feldman EL 2002 Signaling mechanisms that regulate actin-based motility processes in the nervous system. J Neurochem 83:490 –503 Gallo G, Letourneau PC 2004 Regulation of growth cone actin filaments by guidance cues. J Neurobiol 58:92–102 Rodriguez OC, Schaefer AW, Mandato CA, Forscher P, Bement WM, Waterman-Storer CM 2003 Conserved microtubule-actin interactions in cell movement and morphogenesis. Nat Cell Biol 5:599 – 609 Moon A, Drubin DG 1995 The ADF/cofilin proteins: stimulus-responsive modulators of actin dynamics. Mol Biol Cell 6:1423–1431 Bamburg JR 1999 Proteins of the ADF/cofilin family: essential regulators of actin dynamics. Annu Rev Cell Dev Biol 15:185–230 Carlier MF, Laurent V, Santolini J, Melki R, Didry R, Xia GX, Hong Y, Chua NH, Pantaloni D 1997 Actin depolymerizing factor (ADF/cofilin) enhances the rate of filament turnover: implication in actin-based motility. J Cell Biol 136:1307–1322 Chen J, Godt D, Gunsalus K, Kiss I, Goldberg M, Laski FA 2001 Cofilin/ADF is required for cell motility during Drosophila ovary development and oogenesis. Nat Cell Biol 3:204 –209 Nishita M, Tomizawa C, Yamamoto M, Horita Y, Ohashi K, Mizuno K 2005 Spatial and temporal regulation of cofilin activity by LIM kinase and Slingshot is critical for directional cell migration. J Cell Biol 171:349 –359 Gu YY, Zhang HY, Zhang HJ, Li SY, Ni JH, Jia HT 2006 8-Chloro-adenosine inhibits growth at least partly by interfering with actin polymerization in cultured human lung cancer cells. Biochem Pharmacol 72:541–550 Coˆte´ M, Payet MD, Gallo-Payet N 1997 Association of ␣ S-subunit of the GS protein with microfilaments and microtubules: implication during adrenocorticotropin stimulation in rat adrenal glomerulosa cells. Endocrinology 138: 69 –78 Otis M, Gallo-Payet N 2006 Differential involvement of cytoskeleton and rho-guanosine 5⬘-triphosphatases in growth-promoting effects of angiotensin II in rat adrenal glomerulosa cells. Endocrinology 147:5460 –5469 Yusa S, Campbell KS 2003 Src homology region 2-containing protein tyrosine phosphatase-2 (SHP-2) can play a direct role in the inhibitory function of killer cell Ig-like receptors in human NK cells. J Immunol 170:4539 – 4547 Prickett TD, Brautigan DL 2006 The ␣4 regulatory subunit exerts opposing allosteric effects on protein phosphatases PP6 and PP2A. J Biol Chem 281: 30503–30511 Salzmann C, Otis M, Long H, Roberge C, Gallo-Payet N, Walker CD 2004 Inhibition of steroidogenic response to adrenocorticotropin by leptin: impli-


46. 47. 48. 49. 50. 51. 52. 53. 54. 55. 56.

57.

58. 59. 60. 61. 62. 63. 64. 65. 66. 67. 68. 69.

70. 71.

2933

cations for the adrenal response to maternal separation in neonatal rats. Endocrinology 145:1810 –1822 Battista MC, Roberge C, Otis M, Gallo-Payet N 2007 Seladin-1 expression in rat adrenal gland: effect of adrenocorticotropic hormone treatment. J Endocrinol 192:53– 66 Qian H, Sheetz MP, Elson EL 1991 Single particle tracking. Analysis of diffusion and flow in two-dimensional systems. Biophys J 60:910 –921 Huang XC, Richards EM, Sumners C 1996 Mitogen-activated protein kinases in rat brain neuronal cultures are activated by angiotensin II type 1 receptors and inhibited by angiotensin II type 2 receptors. J Biol Chem 271:15635–15641 Bialojan C, Takai A 1988 Inhibitory effect of a marine-sponge toxin, okadaic acid, on protein phosphatases. Specificity and kinetics. Biochem J 256:283–290 Lin CH, Forscher P 1995 Growth cone advance is inversely proportional to retrograde F-actin flow. Neuron 14:763–771 Dent EW, Gertler FB 2003 Cytoskeletal dynamics and transport in growth cone motility and axon guidance. Neuron 40:209 –227 Forscher P, Smith SJ 1988 Actions of cytochalasins on the organization of actin filaments and microtubules in a neuronal growth cone. J Cell Biol 107:1505– 1516 Dehmelt L, Smart FM, Ozer RS, Halpain S 2003 The role of microtubuleassociated protein 2c in the reorganization of microtubules and lamellipodia during neurite initiation. J Neurosci 23:9479 –9490 Dehmelt L, Halpain S 2004 Actin and microtubules in neurite initiation: are MAPs the missing link? J Neurobiol 58:18 –33 Ono S 2003 Regulation of actin filament dynamics by actin depolymerizing factor/cofilin and actin-interacting protein 1: new blades for twisted filaments. Biochemistry 42:13363–13370 Gunsalus KC, Bonaccorsi S, Williams E, Verni F, Gatti M, Goldberg ML 1995 Mutations in twinstar, a Drosophila gene encoding a cofilin/ADF homologue, result in defects in centrosome migration and cytokinesis. J Cell Biol 131:1243– 1259 Hotulainen P, Paunola E, Vartiainen MK, Lappalainen P 2005 Actindepolymerizing factor and cofilin-1 play overlapping roles in promoting rapid F-actin depolymerization in mammalian nonmuscle cells. Mol Biol Cell 16: 649 – 664 Ghosh M, Song X, Mouneimne G, Sidani M, Lawrence DS, Condeelis JS 2004 Cofilin promotes actin polymerization and defines the direction of cell motility. Science 304:743–746 Distasi C, Ariano P, Zamburlin P, Ferraro M 2002 In vitro analysis of neuronglial cell interactions during cellular migration. Eur Biophys J 31:81– 88 Meberg PJ, Ono S, Minamide LS, Takahashi M, Bamburg JR 1998 Actin depolymerizing factor and cofilin phosphorylation dynamics: response to signals that regulate neurite extension. Cell Motil Cytoskeleton 39:172–190 Hansen JL, Servant G, Baranski TJ, Fujita T, Iiri T, Sheikh SP 2000 Functional reconstitution of the angiotensin II type 2 receptor and G(i) activation. Circ Res 87:753–759 Okada K, Takano-Ohmuro H, Obinata T, Abe H 1996 Dephosphorylation of cofilin in polymorphonuclear leukocytes derived from peripheral blood. Exp Cell Res 227:116 –122 Takuma T, Ichida T, Yokoyama N, Tamura S, Obinata T 1996 Dephosphorylation of cofilin in parotid acinar cells. J Biochem 120:35– 41 Niwa R, Nagata-Ohashi K, Takeichi M, Mizuno K, Uemura T 2002 Control of actin reorganization by Slingshot, a family of phosphatases that dephosphorylate ADF/cofilin. Cell 108:233–246 Soosairajah J, Maiti S, Wiggan O, Sarmiere P, Moussi N, Sarcevic B, Sampath R, Bamburg JR, Bernard O 2005 Interplay between components of a novel LIM kinase-slingshot phosphatase complex regulates cofilin. EMBO J 24:473– 486 Gohla A, Birkenfeld J, Bokoch GM 2005 Chronophin, a novel HAD-type serine protein phosphatase, regulates cofilin-dependent actin dynamics. Nat Cell Biol 7:21–29 Arce ME, Sanchez S, Seltzer A, Ciuffo GM 2001 Autoradiographic localization of angiotensin II receptors in developing rat cerebellum and brainstem. Regul Pept 99:53– 60 Zhou FQ, Cohan CS 2001 Growth cone collapse through coincident loss of actin bundles and leading edge actin without actin depolymerization. J Cell Biol 153:1071–1084 Zhang W, Duan W, Cheung NS, Huang Z, Shao K, Li QT 2007 Pituitary adenylate cyclase-activating polypeptide induces translocation of its G-protein-coupled receptor into caveolin-enriched membrane microdomains, leading to enhanced cyclic AMP generation and neurite outgrowth in PC12 cells. J Neurochem 103:1157–1167 Huber AB, Kolodkin AL, Ginty DD, Cloutier JF 2003 Signaling at the growth cone: ligand-receptor complexes and the control of axon growth and guidance. Annu Rev Neurosci 26:509 –563 Bellenchi GC, Gurniak CB, Perlas E, Middei S, Ammassari-Teule M, Witke W 2007 N-cofilin is associated with neuronal migration disorders and cell cycle control in the cerebral cortex. Genes Dev 21:2347–2357

Endocrinology is published monthly by The Endocrine Society (http://www.endo-society.org), the foremost professional society serving the endocrine community.