N-Methyl-D-Aspartate Evokes Rapid Net Depolymerization of Filamentous Actin in Cultured Rat Cerebellar Granule Cells

Spencer L. Shorte

Institut National de la Santé et de la Recherche Médicale (INSERM), Unité 29, Laboratoire de Neurobiologie et Physiopathologie du Développement, Hôpital de Port-Royal, 75014 Paris; and INSERM Unité 261, Institut Pasteur, 75015 Paris, France

    ABSTRACT
Abstract
Introduction
Methods
References

Shorte, Spencer L. N-methyl-D-aspartate evokes rapid net depolymerization of filamentous actin in cultured rat cerebellar granule cells. J. Neurophysiol. 78: 1135-1143, 1997. Filamentous actin(F-actin) was measured in cultured rat cerebellum granule neurons with the use of fluorescently labeled phallotoxin as a site-specific probe for F-actin, and fluorescence microscopy. The averaged apparent intensity of soma-associated F-actin-derived fluorescence (Fapp) was measured from fixed cells after incubation in either 1) normal Krebs solution containing 2 mM extracellular calcium ([Ca2+]ex) or 2) normal Krebs solution plus N-methyl-D-aspartate (NMDA) for 2 min immediately before fixation. NMDA (10, 50, and 100 µM) decreased Fapp to 63 ± 5% (mean ± SE), 53 ± 4%, and 47 ± 2%, respectively, of that measured from control cells. This effect was mimicked by treatment of cells with ionomycin. The ability of NMDA to reduce the Fapp in the presence of [Ca2+]ex was abolished when cells were maintained in [Ca2+]ex-free medium. Cells first treated with NMDA for 2 min and then left in normal medium for 30 min before fixation gave Fapp fluorescence similar to control values (91 ± 12%). However, if the F-actin polymerization inhibitor cytochalasin D was added to cells immediately after NMDA was removed, the Fapp did not recover with time (36 ± 3%). Cells treated for 30 min with cytochalasin D alone showed a small reduction in staining (~20%). It is concluded that the actin polymerization state of rat cerebellar granule neurons is sensitive to changes in intracellular calcium, and that NMDA receptor activation evokes an initial rapid depolymerization of F-actin.

    INTRODUCTION
Abstract
Introduction
Methods
References

Changes in cytoskeleton protein architecture are thought to underlie many neuronal cell functions including neurite outgrowth (Lankford and Letourneau 1989), cell migration (Rakic and Komuro 1995; Rivas and Hatten 1995), axonal transport (Morris and Hollenbeck 1995), growth cone advance/collapse (Fan et al. 1993; Forscher and Smith 1988; Lin and Forscher 1995; Neely and Gesemann 1994; Tanaka and Sabry 1995), and neurotransmitter release (Bernstein and Bamburg 1989). One major cytoskeleton protein, actin, abundant in neurons (Matus et al. 1982), exists in a dynamic equilibrium comprising two basic states: a filamentous polymeric form (F-actin) and a globular monomeric form(G-actin). In vitro, F-actin is assembled from G-actin spontaneously via noncovalent interactions, and a steady state is reached when the G-actin concentration approaches the critical concentration (Fechheimer and Zigmond 1993; Schafer and Cooper 1995). In vivo, G-actin concentration is some 10-fold higher than predicted from the critical concentration (Symons and Mitchison 1991) because of stringent regulation exerted by an array of actin-binding proteins distinguished on the basis of their characteristic actin polymerization, depolymerization, capping, nucleation, and bundling activities (for review see Schafer and Cooper 1995). In turn, these activities are modulated by various physiological factors, i.e., cytosolic ions, secondary messenger metabolites, and protein phosphorylation (Agnew et al. 1995; Allen and Janmey 1994; Janmey and Stossel 1987; Lamb et al. 1993; for review see Schafer and Cooper 1995).

Activation of the N-methyl-D-aspartate (NMDA) receptor depolarizes neurons, leading to changes in the cytosolic concentrations of calcium ([Ca2+]i) (Mayer et al. 1987), protons (Endres et al. 1986), and magnesium (Brocard et al. 1993), and also alters the phosphorylation state of many cellular proteins (Scheetz and Constantine-Paton 1996). All of these parameters could directly effect the actin polymerization state in neurons, and because the receptor is modulated by drugs that alter actin polymerization state it has been suggested that NMDA receptor activation might cause depolymerization of F-actin by way of a positive feedback mechanism regulating subsequent channel activity (Rosenmund and Westbrook 1993). Indeed, recently an actin filament bundling protein, alpha -actinin, was reported to bind directly to the cytoplasmic tails of both the NR1 and NR2B NMDA receptor subunits, and in the case of the NR1 subunit alpha -actinin binding was competitively antagonized by binding of calmodulin (Wyszynski et al. 1997). Moreover, neuronal voltage-gated calcium channels (Johnson and Byerly 1993) and synaptic plasticity (Wang et al. 1996) are also altered by drugs that change actin polymerization state. These observations provide a basis to speculate that activity-dependent changes in the subplasmallema cortical actin cytoskeleton might serve as a mechanism for functional regulation of both receptor localization and channel activity. However, to date there has been no demonstration that receptor-mediated, activity-dependent changes in actin polymerization state occur in intact neurons. The present study directly addresses this question. Cultured rat cerebellar granule (RCG) cells were used as a model, and two semiquantitative fluorescence-based assays were utilized to estimate changes in the F-actin content of individual cells after different treatments. It is concluded that F-actin in these cells is highly stable under resting conditions, but NMDA receptor activation, in the first instance, rapidly and transiently shifts the F-/G-actin equilibrium far in favor of depolymerization.

    METHODS
Abstract
Introduction
Methods
References

Preparation of cultured RCG neurons

The brain was removed from 7- to 8-day-old Wistar rats killed by cervical dislocation and rapid decapitation. The cerebellum formation was dissected from the isolated brain tissue in calcium- and magnesium-free phosphate-buffered saline (0.02 M, pH 7.4) supplemented with 0.6% (wt/vol) glucose. The dissected tissue was incubated for 20 min in phosphate-buffered saline containing 0.2% (wt/vol) trypsin (bovine pancreas, Sigma) plus 0.1% (wt/vol) DNAse (type I, Sigma) at room temperature. The enzymatic digestion was quenched by addition of 1% (wt/vol) soybean trypsin inhibitor. The partially digested tissue was then centrifuged gently. This was immediately followed by trituration of tissue through a fire-narrowed glass Pasteur pipette. The tissue was allowed to settle and then was resuspended in culture medium(D-MEM, GIBCO, containing 20 g/l glucose, 5 U/ml penicillin, and 5 µg/ml streptomycin plus 10% NU Serum, Collaborative Research) to which DNAse was added (0.05%, wt/vol). After a brief centrifugation (90 g, 10 min) the cell pellet was resuspended in culture medium without DNAse, and the cells were plated on poly-l-lysine-coated (Sigma, 30-70 kDa, 0.1 µg/ml, 12 h) round coverslips (1 cm diam) at a density of ~150,000 cells/cm2 in nontissue culture treated Petri dishes (1 cm diam, multiwell). Cells were incubated in a cell culture oven at 37°C and under an atmosphere of 95% O2-5% CO2. Cells were used for experiments between 5 and 14 days after preparation.

Treatment of cultured cells before staining

For experiments, the cells were removed from the culture environment and the culture medium was gently diluted over 1 min by addition of an equivalent volume of experimental medium. The latter was a standard extracellular solution prepared by 10-fold dilution of stock solution [1.9 M NaCl, 10 mM KCl, 50 mMN-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES)-NaOH, 50 mM HEPES, and 100 mM glucose], yielding a working experimental buffer that was adjusted to 330 mosM by dilution (5-10%) at pH 7.4. Before use, the following additions were made (final concentrations in parentheses): tetrodotoxin (0.5 µM), 6-cyano-7-nitroquinoxaline-2,3-dione (1 µM), and bicuculline (1 µM). CaCl2 (2 mM) was also added unless stated otherwise. Tetrodotoxin was used to abolish spontaneous electrical activity, bicuculline blocked gamma -aminobutyric acid-receptor-mediated inhibitory activity, and 6-cyano-7-nitroquinoxaline-2,3-dione blocked kainate receptor activity. Magnesium was omitted to allow full receptor activation by NMDA (Nowak et al. 1984). This basic external bathing solution was prepared at the start of each day's experiments. To ensure that observations were not affected by differences in osmolarity between solutions, all drugs were diluted into and all cell manipulations were performed with the use of aliquots taken from the same fresh stock solution. After 10-15 min, cells were completely exchanged into the standard extracellular medium, and drug treatments were carried out with the use of the specified agents diluted into experimental medium at the final concentrations stated in figures. Addition of drugs was by a single medium exchange (total volume 1 ml) with the use of a hand-held pipette. Where NMDA was used, glycine was also included at 10 µM. All control samples were treated in parallel to drug-treated samples by exchanging medium containing no drugs. Control samples fixed after 2 or 30 min were found to be identical (data not shown), and so data from these samples were pooled where appropriate. Each condition described in figures represents data collected from between two and seven independent cell preparations as stated. All drugs and reagents were research-grade high purity and were (unless otherwise stated) supplied by Sigma (Saint Quentin Fallavier, France), Calbiochem (Meudon, Europe), Aldrich Chemical (Saint Quentin Fallavier, France), and Tocris Cookson (Bristol, UK).

Fixation, extraction, and staining of cells with fluorescent phallotoxin and fluorescein

The method and order of fixation are important factors in studies of the actin cytoskeleton. For example, in fish keratocyte lamellipodium there are two populations of F-actin filaments, a Triton-detergent-soluble and an insoluble fraction; if Triton extraction precedes fixation, the former is not preserved or labeled during subsequent phalloidin staining (Small et al. 1995). Thus, in the present study, after drug treatments of RCG neurons the medium in incubation wells was removed and immediately replaced with 1 ml of fixation buffer. The latter contained 4% paraformaldehyde (wt/vol) diluted into "cytoskeleton preservation buffer" (CPB) (based on Rivas and Hatten 1995) comprising 0.02 M phosphate-buffered saline, 3 mM MgCl2, 2 mM ethylene glycol-bis(beta -aminoethyl ether)-N,N,N',N'-tetraacetic acid, and 5 mM piperazine-N,N'-bis(2-ethanesulfonic acid), pH 6.9. This buffer served to rapidly fix cells, to quench calcium activated processes, and to preserve the actin cytoskeleton. After 30 min, the fixation buffer was exchanged for CPB plus Triton X-100 detergent (0.5% vol/vol) but without paraformaldehyde, and the cells were extracted by careful triplicate exchange of the incubation volume at three 10-min intervals with CPB. All subsequent treatments were carried out in CPB plus Triton X-100 (0.5%, vol/vol). The concentration of Triton X-100 in the preservation buffer was based on the observation that this agent preserves and stabilizes the disposition of cytoskeletal proteins at this concentration (Fulton and L'Ecuyer 1993).

The fixed and extracted cells were stained with rhodamine-conjugated phalloidin (TRITC-Ph; Molecular Probes and Sigma) on the basis of the method described by Haughland and colleagues (Haughland 1992; Haughland et al. 1994). The TRITC-Ph was stored at -20°C in a methanolic stock solution at either 6.6 or 77 µM. However, methanol disrupts F-actin filaments (Cassimeris et al. 1990; Haughland 1992), and the solvent was therefore evaporated. The TRITC-Ph was then resuspended in CPB (0.3 µM) and cells on coverslips were incubated in this solution for 30 min to 1 h. The cells on coverslips were then washed three times to ensure the removal of unbound phalloidin and then inverted and mounted on microscope slides under a solution containing 50% CPB:glycerol. The samples were sealed airtight with varnish and either immediately visualized (see below) or stored at -20°C until visualization.

For some of the experiments a second fluorescein (fluorescein isothiocyanate, FITC)-based cross-staining procedure (Fan et al. 1993) was used in combination with TRITC-Ph. In this case the procedure was identical to that described above, except that after TRITC-Ph was removed, cells were finally incubated in a fresh 0.001% solution of 5-[4,6-dichlorotriazin-2-yl]-amino-fluorescein (DTAF, Sigma), a fluorescein derivative, in CPB for 30 min and then washed thoroughly preceding mounting.

Confocal visualization of cell-associated fluorescence

Each sample was visualized on a scanning confocal laser system (BIORAD MRC-600, argon-krypton laser) connected to an upright microscope (Axioskop, Karl Zeiss) equipped with a ×40 water-immersion objective (numerical aperture 0.7). To avoid nonspecific breakdown of proteins at room temperature, stained samples were not left on the microscope for >60 min. The intensity of staining was recorded from individual cell somata according to strict criteria. Cells that had characteristics of healthy RCG neurons were visually selected with the use of the normal light microscope mode. The fluorescence image was then visualized and the focal plane was adjusted to the level at which the greatest signal was apparent. The fluorescence image was then recorded. The sensitivity of the photomultipliers used to detect fluorescence was fixed for each batch of samples with the use of a linear signal and background gain. In choosing the necessary range for gain controls, both control samples and treated samples were initially visualized to ensure that the dynamic range of the 8-bit signal was fully employed. Having "fixed" the gains, the samples were then visualized at ×6.0 zoom with the use of an integrated slow scan mode and were kalman averaged from four independent passes. To avoid sample bleaching, a neutral-density filter was used that allowed either 1 or 3% transmission of excitation light. The confocal aperture was fixed at 10%, giving a confocal field depth of ~3 µm. It should be noted that because the brightest fluorescence signal was used to establish the focal plane for each sample, the majority of the measured signal was from the uppermost part of the cell bodies. For samples stained only with TRITC-Ph, the excitation wavelength used was 568 nm, and fluorescent emission was measured at 585 nm. For cells cross-stained with DTAF, the samples were treated exactly as above except that a second independent optimized measurement was made subsequently. The excitation light was switched to 488 nm and a second photomultiplier was used to measure fluorescent emission at 522 nm. The same confocal aperture and neutral-density filters were used, and the gains of a second amplifier were similarly fixed for final measurements. When cross-stained cells were excited at 488 nm there was no signal detected on the TRITC photomultiplier set to maximum gain. Equally, fluorescence from cells stained with TRITC-Ph and excited at 568 nm did not give a detectable signal on the FITC photomultiplier. Thus the sequential independent measurement of the TRITC-/FITC-derived signals made crossover between these fluorophores negligible. Equally, cellular autofluorescence was not detected above background levels when the excitation wavelengths and gain settings that were routinely used for acquiring experimental images were employed. Finally, recorded images were then saved on a computer for later analysis.

Calculation of average apparent somatic F-actin fluorescence and relative F-actin content

For analysis of all images, the standard quantitative image analysis package CoMOS (BIORAD) was used. Individual cells in each image were selected, and for each a circumference outline was drawn along the outer edge of the plasma membrane. This area was measured in pixels and converted to µm2 (1 pixel square = 0.074 µm2), and the average intensity of fluorescence was calculated from the sum of the intensities of the detected pixels within the delineated area. For any given batch of conditions measured on the same day, data were arbitrarily comparable, because the measurement gains were fixed. However, to combine data from different experimental days it was necessary to normalized the apparent fluorescence measured from each cell on any given day. This was done by expressing the apparent signal from each cell as a percentage of the mean measured from the corresponding controls on the same day: the apparent F-actin-dependent fluorescence (Fapp). The data from different days and individual cells could then be combined and collated to calculate mean Fapp values.

The second method used to quantify the TRITC-Ph signal was based on the cells cross-stained with DTAF. The second FITC-based fluorescence signal was analyzed sequentially and as described above. The relative F-actin content (RA) was calculated for each given cell in each batch of experiments according to the method already described by Fan et al. (1993) and shown below
Apparent Ratio
 = <FR><NU>average pixel intensity for cell-associated TRITC-Ph fluorescence</NU><DE>average pixel intensity for cell-associated DTAF (FITC) fluorescence</DE></FR>
RA = 100⋅[(Apparent Ratio)/(Mean Ratio for Control Cells)]
Thus the ratio from each cell was normalized to the calculated mean ratio measured from corresponding control samples. The RA value was taken to be a generic indicator of F-actin levels in individual cells. Where appropriate, data were collated from many experiments to find the mean values for each treatment tested and compared with the use of a standard independent t-test (P = 0.01).

    RESULTS AND DISCUSSION

NMDA stimulation causes a decrease in total cellular F-actin

The percentage normalized apparent intensity of TRITC-Ph fluorescence (Fapp) has been used previously as an arbitrary quantitative method for assessing the content of F-actin in fixed cells (Cassimeris et al. 1990; Fan et al. 1993; Howard and Oresajo 1985a,b; Perrin et al. 1992). This approach also formed the basis for the studies presented here. First, the effect of NMDA stimulation on somatic (cell body)-associated TRITC-Ph stain intensity (see Fig. 2) was investigated. Treatment of cells (as described in METHODS) with NMDA at 10, 50, and 100 µM for 2 min caused a reduction in the mean Fapp to 63 ± 5% (mean ± SE), 53 ± 4%, and 47 ± 2%, respectively, compared with cells treated with control medium (Figs. 1 and 2).


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FIG. 2. Relative cellular localization of somatic TRITC-Ph fluorescence. Cells were cross-stained with TRITC-Ph and 5-[4,6-dichlorotriazin-2-yl]-amino-fluorescein (DTAF). TRITC-Ph fluorescence images were acquired from field of control cells (A) and field of cells treated with NMDA (100 µM, 2 min, C). Both images are comparable because they were collected with the use of identical parameters (see METHODS), and "true" intensity of staining is shown with the use of linear pseudocolor lookup table (red is low fluorescence, and white is high fluorescence). Note very bright fluorescence is associated with cortical region of cell body. Some fluorescence is also detected in central regions because of wide confocal plane selected, which allowed detection of filamentous actin (F-actin)-dependent fluorescence from the uppermost part of the cell cytoplasm (see METHODS). Images shown in A and C were then ratioed (TRITC/FITC) on a pixel-by-pixel basis; corresponding images are shown in B and D. Pseudocolor representation of ratioed data are scaled to 256 gray levels corresponding to arbitrary ratio range of 0-9.0.


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FIG. 1. Dose-dependent effect of N-methyl-D-aspartate (NMDA) treatment on cell-associated rhodamine-conjugated phalloidin (TRITC-Ph)fluorescence. TRITC-Ph fluorescence was measured from individual cell somata as described in METHODS. Each value was percentage normalized to respective mean value measured from corresponding control cells (apparent F-actin-dependent fluorescence, Fapp). Calculated mean Fapp values are shown from cells treated for 2 min with NMDA at 0, 10, 50, and 100 µM. Numbers in parentheses: numbers of cells sampled for each condition.

This effect was unlikely to have arisen from nonspecific changes, because phallotoxins specifically bind F-actin in preference to other proteins including G-actin (TRITC-Ph has a dissociation constant for F-actin of ~40 nM) (Haugland 1992; Huang et al. 1992) and the effect of NMDA was also observed in cells treated with 3% bovine serum albumin after fixation and extraction (data not shown). It was assumed, therefore, that Fapp was a representative index of the amount of F-actin present in each cell (Bernstein and Bamburg 1989; Cassimeris et al. 1990; Fan et al. 1993; Howard and Oresajo 1985a,b; Perrin et al. 1992). Because NMDA treatment caused a decrease in Fapp measured from RCG neurons, it was concluded that the amount of F-actin present in cell somata was specifically decreased by NMDA stimulation. The magnitude and time course of the NMDA-induced F-actin breakdown observed here compares well with that reported for parathyroid cells, where parathyroid hormone causes a 50-60% calcium-dependent reduction in F-actin levels within minutes of agonist stimulation (Egan et al. 1991). Equally rapid, calcium-dependent, secretagogue-induced changes have also been reported for other nonneuronal cells, including chromaffin cells, mediated by nicotinic acetylcholine receptor activation (Vitale et al. 1991), and pancreatic acinar cells, mediated by muscarinic acetylcholine receptor activation (Perrin et al. 1992). The current study provides the first report of such a phenomenon in neuronal cells.

Dependence of NMDA-induced F-actin decreases on extracellular calcium

Exchanging cells into medium containing no added extracellular calcium (for 30 min before fixation) resulted in a Fapp decrease of ~20-25% compared with controls (Table 1). This was not unexpected. In platelet cells, for example, stringent calcium chelation reduces thrombin-induced actin nucleation and polymerization activities (Hartwig 1992). In RCG neurons, maintenance of basal F-actin levels might also be dependent on intracellular calcium concentration.

 
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TABLE 1. Mean percentage normalized Fapp, cell areas, and estimated cell volumes in rat cerebellar granule cells after different treatments

Cells incubated in calcium-free medium and then subsequently with calcium-free medium containing NMDA (100 µM, 2 min) showed no change as compared with non-NMDA-treated cells in calcium-free medium (Table 1). The most obvious conclusion from this observation is that the NMDA-induced decrease in F-actin levels was dependent on extracellular calcium; this suggests that calcium influx leading to a cytosolic calcium increase was necessary to the effect. The source of the calcium fueling the [Ca2+]i rise was probably through the open NMDA channel protein itself (Mayer et al. 1987), but NMDA-induced depolarization can also cause entry of calcium through voltage-gated calcium channels also present in these cells (Pearson et al. 1995). Furthermore, these observations do not exclude the possibility that NMDA receptor activation caused a [Ca2+]i rise propagated, in part, by calcium-induced calcium release from intracellular stores depleted during the 30-min preexposure to calcium-free medium. However, it is reasonable to suggest that the mechanism of the NMDA-induced decrease in F-actin levels involves a calcium-dependent activation of F-actin fragmenting activity present in these cells.

Dependency of F-actin recovery following NMDA stimulation on actin polymerization

Cells were treated with NMDA for only 2 min, at which time it was removed (by exchanging cells into control medium), and cells were left for 30 min in normal medium before fixation. This resulted in an apparent recovery of Fapp, which reached ~90% of its value recorded from control cells (Table 1). To establish whether the apparent recovery of F-actin levels was dependent on polymerization, cells treated in parallel with NMDA for 2 min were left for 30 min in the continued presence of cytochalasin D (20 µM), a specific inhibitor of actin polymerization, although its effects are complex (Cooper 1987). Under these conditions the apparent F-actin recovery was not observed. Indeed, the Fapp was less than that recorded from cells treated with NMDA or ionomycin (Table 1). From this it was concluded that the apparent recovery of cellular F-actin levels following NMDA treatment was dependent on actin polymerization, and the simplest explanation for the effect of NMDA was that it caused actin depolymerization.

An actin-depolymerizing factor has been isolated from brain, but its ability to depolymerize actin does not appear to be calcium dependent (Petrucci et al. 1983). A calcium-dependent actin-depolymerizing protein has yet to be isolated from neurons but will presumably share some characteristics with the calcium-dependent actin-depolymerizing proteins already isolated from nonneuronal cells (Schafer and Cooper 1995), notably gelsolin (Yin and Stossel 1979). However, the data presented here do not exclude the possibility that NMDA evoked a net F-actin polymerization through limited proteolysis of G-actin. Indeed, there is evidence to suggest that calcium-activated neutral proteinases help to regulate the normal functional reorganization of cytoskeletal proteins in response to secondary messengers under nonpathological conditions (Nixon 1989), and NMDA receptor activation causes accumulation of spectrin breakdown products in hippocampal slices (Seubert et al. 1988). Proteolysis would also result in a G-/F-actin equilibrium favoring depolymerization. Under these conditions the apparent recovery could have been by rapid de novo synthesis of G-actin monomers (Fulton and L'Ecuyer 1993) followed by polymerization. This possibility has yet to be tested with the use of protein synthesis inhibitors.

Effect of artificially raised [Ca2+]i on F-actin

Ionomycin is an ionophore that works as a calcium-proton exchanger, dissipating calcium gradients in favorable ionic environments (pH > 7.0) (Liu and Hermann 1978). It is expected to release large proportions of intracellularly stored calcium pools and to increase calcium entry over the plasma membrane, resulting in increased [Ca2+]i and destabilization of the actin cytoskeleton (Lankford and Letourneau 1989). Cells treated with ionomycin (5 µm, 5 min) in the presence of extracellular calcium showed a 50% reduction in Fapp (Table 1). The ability of ionomycin to reduce F-actin levels in RCG neurons lends support to the conclusion that F-actin in these cells is indeed susceptible to calcium-dependent destabilization. However, when cells incubated in medium containing no added calcium were challenged with ionomycin in the continued absence of extracellular calcium, a rise in Fapp was observed (Table 1). It is possible that this observation arose from an artifact of the technique, and this is discussed below.

Relationship between F-actin, Fapp, and cell volume

Volume regulation in many cell types is altered by treatments that affect [Ca2+]i and/or the cytoskeleton (for review see Pierce and Politis 1990). Moreover, cell volume changes have been reported to accompany F-actin disruption in epithelial cells (De Filippo et al. 1995). The consideration of cell volume changes is important to the interpretation of Fapp measurements in the current study because total cellular protein concentration is proportional to cell volume. Consequently the intensity of fluorescent protein markers will also be effected by cell volume. Thus an increase in cell volume would necessarily decrease the cellular Fapp, and a decrease in cell volume would result in the opposite effect.

The real (non-normalized) mean detected cell area for control cells was 51 ± 15 µm2 as calculated from fluorescence images (see METHODS). Assuming that the cell bodies are nearly spherical (a fact that was confirmed by making 1-µm optical slices of cell somata with the use of the confocal microscope), this corresponds to an average cell radius of 4.03 µm. The total excluded cell volume was estimated to be 274 µm3. Treatment with NMDA increased this to 282 µm3 (Table 1), an increase that was not significantly different from values in control cells (P = 0.01). Thus treatment of cells with NMDA did not change cell volume, suggesting that the NMDA-induced decrease in Fapp was not an artifact in this case.

On the other hand, ionomycin caused a significant reduction in cell volume (Table 1). The same was true for cells treated in medium containing no extracellular calcium with or without NMDA (Table 1). This, however, is the opposite effect to that expected to reduce Fapp, and suggests that the decreased Fapp observed as a result of these treatments was unlikely to be an artifact. This was not true for cells treated with ionomycin in the absence of extracellular calcium; these cells' volume was also significantly reduced, and in this case the Fapp was increased as might be expected if an artifact were responsible for the effect. Likewise, cells treated under the two conditions in which cytochalasin D was used both showed significantly increased cell volumes and decreased Fapp signals (Table 1), which also raises the possibility that an artifact was involved. This necessitated the development of a second approach to measure Fapp independently of cell volume changes.

Effect of NMDA and cytochalasin D on RA

The RA of cells can be determined by using a nonspecific protein stain as a reference and calculating the ratio of the total cell-associated TRITC-Ph fluorescence against the total cell-associated nonspecific protein stain (Fan et al. 1993). One advantage of this approach is that it rules out possible artifacts due to volume changes. It also provides an intrinsic correction for variations in sample thickness, and the optical efficiency of the microscope's excitation/emission-light pathways. This method was used to further quantify the effects of NMDA and cytochalasin D.

Cells were cross-stained with both TRITC-Ph and DTAF, and were then visualized as described in METHODS. An example of the typical pattern of TRITC-Ph fluorescence observed for control cells is illustrated in the image shown in Fig. 2A. The pattern of DTAF staining was always homogeneous across the entire cell soma (data not shown), whereas the distribution of TRITC-Ph staining was anisotropic: there were high levels of staining across the cytoplasm but the highest levels were detected in a broad cortex region (Fig. 2A). The fluorescence detected across the central part of the cells is due to detection of F-actin-dependent fluorescence in the uppermost part of the cell cytoplasm lying in the focal plane, whereas the brighter cortical staining is due to cytoplasmic regions perpendicular to the focal plane.

The nonnormalized TRITC-Ph/FITC ratio image for the same cells as shown in Fig. 2A is shown in Fig. 2B. In the ratiometric image the cortex plasma membrane signal is clearly visible because the ratio enhanced the signal measured from cytoplasmic volumes relative to the volume regions also occupied by the nucleus, which is large in these cells. For comparison, a typical example of TRITC-Ph staining in cells after 2 min of NMDA treatment is shown in Fig. 2C. It was noted that both the cortex region staining and the cytoplasmic staining were reduced compared with control cells and this was also reflected in the ratiometric images (Fig. 2D). Comparison of the images from control cells and NMDA-treated cells showed that the largest signal change in the raw and ratiometric images was in the plasma-membrane-associated cortex region of cell somata.

For each cell in each experiment the TRITC-Ph/FITC ratio was normalized to the mean ratio value calculated from corresponding control cells: the RA was taken to be a semiquantitative indicator of the amount of actin in each cell (see METHODS). The RA mean value calculated from cells treated with NMDA (100 µM, 2 min) was 70 ± 4% (n = 97) of that measured for controls and this was significantly different(P = 0.01). Analysis of the population distribution of cellular RA contents showed control cells to be grouped in a single population that was best fitted by a single Gaussian curve (Fig. 3A). However, NMDA-treated cells were distributed into two distinct populations best fitted by two Gaussian curves (Fig. 3B): one population (n = 58 of 97 cells tested) gave a peak at 37% of that measured from controls, whereas the second population (n = 39 of 97 cells) appeared to be completely unchanged from controls (Fig. 3B).


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FIG. 3. Effect of NMDA and cytochalasin D on relative F-actin content (RA). RA value of individual cells was calculated after different treatments. A: control (no drugs). B: NMDA (100 µM, 2 min). C: NMDA (100 µM, 2 min) followed by 30-min recovery in normal medium. D: NMDA (100 µM, 2 min) plus cytochalasin D (20 µM, 30 min). E: cytochalasin D alone (20 µM, 30 min). Corresponding population distribution of RA values observed after each treatment are shown in A-E, respectively). Dashed lines: best-fit Gaussian curves for each population. Note: population (n = 33) shown in E does not include 3 cells that had % normalized RAs of 193, 202, and 205%, respectively (total number of cells examined was 36).

In parallel, other cells were treated with NMDA (100 µM, 2 min), which was removed, and the cells were left for 30 min in normal medium to recover. The mean RA of the total population of cells was 118 ± 12% (n = 59) of the control samples, and this was not significantly different from the control samples (P = 0.01). However, the population distribution of RA values measured from these cells was also best fitted by two Gaussian curves (Fig. 3C). One population had a peak at 72% (n = 42 of 59 cells tested), whereas the second was at 160% (n = 17 of 59 cells tested) compared with control cells. The "high" RA population was significantly higher than control cells, and the "low" RA population was significantly lower than the control population but also significantly higher than the NMDA-responsive population (P = 0.01).

When cells were treated with NMDA (100 µM, 2 min) and cytochalasin D (30 min), the calculated mean RA value was 47 ± 2% of that measured from control cells, and this was found to be a significant difference (P = 0.01). In this case the population distribution was best fitted by a Gaussian curve with a single peak at 48% (n = 63 cells tested) of that measured from parallel controls (Fig. 3D). Thus the effect of cytochalasin D was to enhance the effect of NMDA so that all cells showed a decreased RA value compared with controls, and the bimodal pattern of recovery was abolished.

Conversely, the mean RA value calculated from cells treated with cytochalasin D alone (20 µM, 30 min) was 89 ± 3% (n = 33 cells tested) of that recorded from control cells. The population distribution was best fitted by a single Gaussian curve with a peak at 81% of that measured from parallel controls (Fig. 3E). Neither of these values differed significantly from that of control samples (P = 0.01). It was therefore concluded that cytochalasin D treatment alone caused only a small decrease in the F-actin-dependent fluorescence of RCG neurons under the conditions described in this study. However, it does not rule out the possibility that cytochalasin D treatment caused a change in the organization of F-actin. In some other cell types (Cassimeris et al. 1990; Cooper 1987; Hartwig 1992), cytochalasins are effective F-actin fragmenting agents but their effects are complicated and do not necessarily cause F-actin depolymerization per se (Cooper 1987). Thus in RCG neurons the ability of cytochalasin D to fragment F-actin was limited but the recovery of F-actin (after NMDA treatment) was completely dependent on cytochalasin-D-sensitive polymerization processes.

Physiological significance of NMDA-induced actin depolymerization

The present study provides evidence for a functional signal-transduction link between NMDA receptor activation and the polymerization state of the integral cytoskeletal protein actin. A dynamic link between filamentous cytoskeletal protein polymerization (stability) and NMDA receptor activity has been postulated previously (Halpain and Greengard 1990; Montoro et al. 1993; Rosenmund and Westbrook 1993) but not directly demonstrated. The observations presented here provide the first direct evidence that actin gel-sol transitions in intact living neurons are evoked rapidly (s/min) by NMDA receptor activation.

The ability of NMDA to decrease cellular F-actin levels could form a part of the hypothetical mechanism for calcium-dependent feedback modulation of NMDA channel activity suggested by Rosenmund and Westbrook (1993). These workers showed that repeated NMDA receptor activation in the presence of cytochalasin D over 25 min resulted in modulation of NMDA channel activity. This is comparable with the protocol that evoked actin depolymerization as reported here, but also resulted in cell volume destabilization probably due to the breakdown of the actin cytoskeleton. Changes in cell volume might lead to changes in membrane tension. In turn, this might allow mechanosensitive modulation of NMDA channel activity (Paoletti and Ascher 1994). Notwithstanding this, an actin filament bundling protein, alpha -actinin, binds the cytoplasmic tail the NR1 receptor subunits directly, and this is competitively antagonized by binding of calmodulin (Wyszynski et al. 1997). The data presented here suggest that NMDA directly alters the cortical actin cytoskeleton. This may in turn effect the ability of alpha -actinin to bind the receptor and therefore ultimately alter the ability of calmodulin to bind and modulate NMDA receptor activity (Ehlers et al. 1996). It is interesting that the activity and disposition of acetylcholine receptors are also effected by actin gel-sol transitions (Bencherif and Lukas 1993), and this may involve direct interaction of actin with G proteins that mediate signal transduction (Ibarrondo et al. 1995). Thus NMDA-evoked actin gel-sol transitions might alter the activity of signal transduction pathways facilitating receptor "cross talk" and/or feedback modulation.

Recent studies have provided strong evidence that dynamic morphological changes occur in dendritic spines during neuronal activity (Dailey and Smith 1996). Dendritic spines do not contain microtubules but do contain large amounts of actin (Matus et al. 1982), and the morphological changes are likely to involve actin gel-sol transitions. It is possible that the changes observed in somatic F-actin levels reported here could also occur in dendritic spines. Indeed, F-actin gel-sol transitions have been reported to accompany depolarization-evoked neurotransmitter release from synaptosomes (Bernstein and Bamburg 1989), and most recently F-actin disruption was demonstrated to effect short-term synaptic plasticity without effects on normal synaptic transmission (Wang et al. 1996). Wang et al. suggested that F-actin provides a barrier to the release from the reserve pool but not the immediately available pool of synaptic vesicles from active zones of frog neuromuscular junction. This is analogous to the role proposed for F-actin in regulation of exocytosis from activated mast cells (Cheek and Burgoyne 1991; Muallem et al. 1995; Vitale et al. 1991), and it makes reasonable the suggestion that actin gel-sol transitions might be fundamental to the mechanisms for generation, maintenance, and regulation of some forms of synaptic plasticity. Unfortunately, in the current study it proved technically unfeasible to visualize and quantify actin staining in the neurites and processes of RCG neurons, and future studies will necessitate new technical approaches to address such questions, ideally in living cells.

Finally, it has been shown that actin depolymerization has an excitoprotective effect against amyloid beta -peptide-induced neuronal cell death, probably through its ability to specifically limit glutamate-activated calcium influx that triggers apoptosis (Furukawa and Mattson 1995; Furukawa et al. 1995). The current study demonstrates that NMDA receptor activation leads directly to actin depolymerization, and could therefore provide a mechanism through which NMDA receptor activation induces excitoprotection against glutamate-induced excitotoxicity (Damschroder-Williams et al. 1995). Thus NMDA-receptor-induced changes in the polymerization state of the actin cytoskeleton might be an important phenomenon in neuronal cell signal transduction and pathophysiology.

    ACKNOWLEDGEMENTS

  The author thanks Dr. E. Tremblay for preparation of excellent rat cerebellar granule cell cultures, and Drs. E. der Terrosian, P. Legendre, and P. Bregestovski for invaluable advice and critical reading of the manuscript.

  This work was supported by the Institut National de la Santé et de la Recherche Médicale and the European Commission, through grants awarded to Dr. Y. Ben-Ari (INSERM U-29); and Le Ministère de L'Education Nationale de L'Enseignement Supérieur, de La Recherche et de L'Insertion Professionnelle, through an award to Dr. P. Bregestovski. The author is particularly grateful to the European Commission for receipt of an Individual Fellowship awarded through the Training and Mobility of Researchers Program, and the Fondation pour la Recherche Medicale.

    FOOTNOTES

  Present address and address for reprint requests: INSERM Unité 261, Neurobiologie Cellulaire et Moleculaire, Bâtiment des Biotechnologies, Institut Pasteur, 28 Rue du Docteur Roux, 75274, Paris Cedex 15, France.

  Received 9 December 1996; accepted in final form 19 March 1997.

    REFERENCES
Abstract
Introduction
Methods
References

0022-3077/97 $5.00 Copyright ©1997 The American Physiological Society