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Originally published In Press as doi:10.1074/jbc.M800752200 on March 22, 2008

J. Biol. Chem., Vol. 283, Issue 20, 13992-14001, May 16, 2008
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Phospholipase C-{delta}1 Expression Is Linked to Proliferation, DNA Synthesis, and Cyclin E Levels*Formula

Jonathan D. Stallings{ddagger}, Yue X. Zeng§, Francisco Narvaez§, and Mario J. Rebecchi§1

From the {ddagger}Department of Clinical Investigation, Madigan Army Medical Center, Tacoma, Washington 98431 and the §Department of Anesthesiology, State University of New York, Stony Brook, New York 11794

Received for publication, January 29, 2008 , and in revised form, March 21, 2008.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
We previously reported that phospholipase C-{delta}1 (PLC-{delta}1) accumulates in the nucleus at the G1/S transition, which is largely dependent on its binding to phosphatidylinositol 4,5-bisphosphate ( Stallings, J. D., Tall, E. G., Pentyala, S., and Rebecchi, M. J. (2005) J. Biol. Chem. 280, 22060-22069[Abstract/Free Full Text] ). Here, using small interfering RNA (siRNA) that specifically targets rat PLC-{delta}1, we investigated whether this enzyme plays a role in cell cycle control. Inhibiting expression of PLC-{delta}1 significantly decreased proliferation of rat C6 glioma cells and altered S phase progression. [3H]Thymidine labeling and fluorescence-activated cell sorting analysis indicated that the rates of G1/S transition and DNA synthesis were enhanced. On the other hand, knockdown cultures released from the G1/S boundary were slower to reach full G2/M DNA content, consistent with a delay in S phase. The levels of cyclin E, a key regulator of the G1/S transition and DNA synthesis, were elevated in asynchronous cultures as well as those blocked at the G1/S boundary. Epifluorescence imaging showed that transient expression of human phospholipase C-{delta}1, resistant to these siRNA, suppressed expression of cyclin E at the G1/S boundary despite treatment of cultures with rat-specific siRNA. Although whole cell levels of phosphatidylinositol 4,5-bisphosphate were unchanged, suppression of PLC-{delta}1 led to a significant rise in the nuclear levels of this phospholipid at the G1/S boundary. These results support a role for PLC-{delta}1 and nuclear phospholipid metabolism in regulating cell cycle progression.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Phosphoinositides are metabolized by a multifaceted and highly regulated set of phosphoinositide-specific enzymes (2-4). Kinases sequentially phosphorylate the inositol head-group of phosphatidylinositol, and phosphatases reverse this process (5-7); both can generate phosphatidylinositol 4,5-bisphosphate (PIP2),2 the principle substrate of phospholipase C (PLC) (2-4). PLC cleaves PIP2 to generate key second messengers, inositol 1,4,5-trisphosphate and diacylglycerol (2-4), that mobilize internal calcium stores (8) and activate protein kinase C (9), respectively. In mammals, the PLC family consists of at least 13 isoforms that fall into six subtypes: β, {gamma}, {delta}, {epsilon} (2-4), {zeta} (10), and {eta} (11, 12).

PIP2 hydrolysis is vital to a wide range of cellular responses, including cytoskeletal remodeling (13), membrane trafficking (14), and gene transcription (15), proliferation, and differentiation (3, 4). PIP2 metabolism and PLC activity (particularly in the nucleus) play prominent roles in cell cycle progression and ultimately influence global decisions, such as differentiation and proliferation (6, 16-18). Indeed, homozygous deletion of PLCβ3 (19) or PLC{gamma}1 (20) is embryonic lethal. Although PLC{delta}1 is not essential, homozygous deletion in mice results in aberrant expression of terminal differentiation markers in several types of skin cells as well as the development of alopecia and spontaneous skin tumors (21). These effects appear to result from increased expression of proinflammatory cytokines (22). Mice that lack both PLC{delta}1 and PLC{delta}3, however, die between embryonic days 11.5 and 13.5 due to abnormal cellular proliferation and apoptosis in placental trophoblasts (23).

Saccharomyces cerevisiae that lack PLC1-1, a homolog to mammalian PLC{delta}1, missegregate chromosomes (24) and exhibit osmotic and temperature sensitivity and defects in metabolism and growth (25, 26). The extents to which these phenotypes are displayed depend on the genetic background of each yeast strain, suggesting that plc1-1 modulates complex multigene processes having significant redundancies. Transformation of these yeast mutants with rat PLC{delta}1 can rescue growth defects (26), consistent with a high degree of functional conservation. Furthermore, overexpression of cyclin-dependent kinase inhibitors, SPL1 or SPL2, rescues these same defects (27), suggesting that PLC1 is somehow linked to cell cycle regulation.

Yagisawa et al. (28) first reported that PLC{delta}1 harbored both nuclear export and import sequences that contribute to its shuttling between the cytoplasm and nucleus. We have demonstrated that PLC{delta}1 accumulates in the nucleus at the G1/S boundary in NIH-3T3 fibroblasts and C6 glioma (1), and many of these observations have been confirmed (29). Here we set out to determine whether this protein plays a role in the cell cycle. We find that suppression of PLC{delta}1 increases cyclin E levels, alters S phase progression, and inhibits cell proliferation.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Synchrony, Flow Cytometry, and Cell Cycle Analysis—Rat C6 glioma cells (American Type Culture Collection) were maintained in RPMI 1640 (Invitrogen) supplemented with 7.5% fetal bovine serum and 1 mM penicillin and streptomycin (all supplements from Invitrogen) at 37 °C in a 5% CO2 humidified incubator. To synchronize cells to the G1/S boundary, adherent glioma cells were twice blocked with 2 mM thymidine (1). Briefly, cultures were washed with RPMI 1640 and incubated in growth medium supplemented with 2 mM thymidine for 12 h, washed three times with growth medium, and incubated for an additional 10 h without thymidine. Cultures were again blocked with 2 mM thymidine for an additional 12 h. To release them from G1/S block, cells were washed three times in RPMI 1640 and fed normal growth medium. To analyze DNA content, cells were harvested by treatment with trypsin/EDTA, washed with phosphate-buffered saline containing 1.0 mM calcium chloride and 2.0 mM magnesium chloride (PBS-CaMg), and fixed in 70% ethanol in phosphate-buffered saline (PBS) with 0.1% fetal bovine serum at 4 °C overnight (1). To stain DNA, ethanolfixed glia were incubated in PBS with 50 mM citrate buffer, 50 µg/ml propidium iodide, and 50 µg/ml RNase A for 30 min at 37 °C (1). After washing with PBS, cell cycle analysis was immediately performed using a fluorescence-activated cell sorter (FACScan; BD Biosciences). Histogram data were analyzed using the program Cychlred and the Origin 7.5 (OriginLab Corp.) peak fit module.

siRNA, Expression Plasmids, and Cell Transfection—We targeted the mRNA sequence 151-172 bp (5'-ggA CCC Cag gCC gCU Cgg TT-3') of rat PLC{delta}1 and designed and synthesized a corresponding duplexed siRNA (Proligo) based on previously described protocols (30, 31). For these experiments, C6 glioma cells were plated on plastic tissue culture dishes or #1.0 borosilicate chambered glass coverslips (Nalge Nunc International) coated with poly-L-lysine (Sigma). Cells were transfected with PLC{delta}1-specific siRNA ({delta}1-siRNA) or a commercially available nonspecific control C-siRNA (Ambion), ranging from 0.01 to 320 nM, using FuGene6 (Roche Applied Science) or RNAiMax (Invitrogen) according to manufacturer's protocol. Three commercially available rat PLC{delta}1-specific siRNAs (Ambion numbers 200652, 49731, and 49541; designated here as siRNA1, siRNA2, and siRNA3, respectively) were also tested for their capacity to knock down expression, and the resulting phenotypes were also assessed. In some experiments, siRNA was delivered with human PLC{delta}1 fused to enhanced green fluorescence protein (EGFP), at ~0.28 µg DNA/cm2, in the same FuGene6 transfection. These expression vectors have been previously described (1).

siRNA transfection efficiency using the Fugene6 or RNAiMax reagents was found to be nearly 100% as assessed using a fluorescently labeled, short double-stranded RNA (Block-It, AlexaFluor Red; Invitrogen). On the other hand, plasmid transfection efficiency using either Fugene6 or FugeneHD (Roche Applied Science) rarely exceeded 25% whether using plasmids encoding the PLC-{delta}1 EGFP fusion protein or EGFP itself.

Cell Proliferation and Viability Assays—To estimate the rate of growth in the presence or absence of {delta}1-siRNA, C6 glioma were plated at densities of 1 x 103 to 1.5 x 105 cells/2-cm2 well and allowed to grow for several days; cell numbers were determined every 24 h with a hemocytometer, and growth rate constants were estimated with the following equation, GR = (ln(nf/ni)/t), where ni = initial cell number, nf = final cell number, and t = time (h). Trypan blue (Invitrogen) was used to determine whether cell membrane integrity was compromised as a result of siRNA treatment. Viability was also assessed using XTT (Biological Industries), which is metabolized to the colored formazin product in the mitochondria (32). During these procedures, C6 glioma cells were cultured in an equivalent medium lacking phenol red. XTT (1 mg/ml) was prepared in serum-free medium containing phenazine methylsulfate (1.53 mg/ml), which stimulates mitochondrial metabolism of XTT (32). 100 µl of XTT/phenazine methylsulfate reagent was transferred to each well containing 400 µl of medium and incubated for 1 h. The conditioned medium from each well was then collected, and the absorbance was measured at 475 nm.

SDS-PAGE and Western Blotting—Each 35-mm dish of monolayer cells was washed twice, with 2 ml each of warm PBS-CaMg. Soluble cellular proteins were then extracted with 0.5 ml of ice-cold extraction buffer (200 mM NaCl, 0.2% Nonidet P-40, 20 mM Tris, pH 8, 1 mM dithiothreitol, 1 mM MgCl2, 1 mM EGTA, and 1% mammalian anti-protease mixture (Sigma)) and incubated for 5 min at 4 °C. The cells were gently scraped up with a rubber policeman and transferred to 1.7-ml Eppendorf tubes. Using this method, the nuclei remained intact, and little of the cellular DNA was extruded. These samples were then subjected to centrifugation at 1000 x g for 4 min at 4 °C, and the supernatant fluids were transferred to new tubes. A portion of each sample was removed for determination of protein concentration. Protein concentration was determined via Bradford assay (Bio-Rad) according to the manufacturer's protocol. An equal volume of acetone at -20 °C was then added to the remaining samples, which were then incubated at -20 °C for at least 30 min. Following centrifugation at 12,000 x g for 5 min, the pellets were washed once with -20 °C acetone/water (1:1, v/v) and dried under vacuum. The dried samples were dissolved in SDS sample buffer to a concentration of 1-2 µg of protein/µl. Samples containing equal concentrations of total protein were separated in an 8% or 10% SDS-polyacrylamide gel and transferred to polyvinylidene difluoride membrane (Bio-Rad) using a Trans Blot semidry transfer apparatus (Bio-Rad) at 14 V for 1.5 h. The blot was then blocked with Tris-buffered saline (TBS) containing 5% nonfat dry milk and 0.1% Tween 20 for 60 min at room temperature. The membrane was incubated in solution containing anti-PLC{delta}1 S-11-2 monoclonal antibody (Upstate Biotechnology, Inc.) or anti-PLC{delta}1 polyclonal (Santa Cruz Biotechnology, Inc., Santa Cruz, CA), anti-cyclin E polyclonal, anti-cyclin A monoclonal, or anti-β-tubulin polyclonal antibody (Invitrogen). The membrane was incubated with secondary antibody solution containing either goat anti-mouse or anti-rabbit IgG (H + L)-horseradish peroxidase conjugate (Bio-Rad). Enhanced chemiluminescence (ECL Plus; Amersham Biosciences/GE HealthCare) was used to detect the binding of the secondary antibody following the manufacturer's protocol. The membranes were imaged using a CCD camera (Eastman Kodak Co.).

[3H]Thymidine Incorporation Assay—C6 glioma cells were transfected with either 160 nM control C-siRNA or PLC{delta}1-specific siRNA and grown in 24-well plates. 48-72 h post-transfection, cultures were washed with PBS-CaMg and incubated in 0.5 ml of RPMI 1640 (7.5% fetal bovine serum, 1% phosphatidylserine) containing 1 µCi/ml [3H]thymidine for 2.5 h (a measure of the number of cells synthesizing nascent DNA, (33)). Alternatively, cultures were transfected and synchronized to the G1/S boundary as described above and then labeled with [3H]thymidine following their release from G1/S block (rate of thymidine incorporation). In each experiment, a portion of each culture was used to determine cell number. Incorporated [3H]thymidine was precipitated with 500 µl of 10% trichloroacetic acid on ice for 20 min. The precipitate was washed twice with 500 µl of each of 10% trichloroacetic acid. Finally, pellets were dissolved with 200 µl of 0.1 N NaOH for 15 min and transferred to a vial containing 4 ml of scintillation fluid and counted in a liquid scintillation spectrometer.

Epifluorescence Microscopy and Indirect Immunofluorescence—Cell monolayers were rinsed once in PBS-CaMg and then fixed with freshly prepared 3.7% (w/v) formaldehyde solution (Fisher) in PBS for 10 min at room temperature. Samples were then washed three times in TBS for 5 min and permeabilized with 0.5% Nonidet P-40 (Sigma) in TBS for 5 min at room temperature. The detergent solution was replaced with blocking solution (TBS containing 5% goat serum (Pierce)) for 30 min at room temperature and then replaced with primary antibody solution (1:200; rabbit anti-cyclin E in TBS with 1% goat serum) overnight at 4 °C. Samples were washed three times in 1 ml of TBS for 7 min each and then incubated at 37 °C for 1 h in goat anti-rabbit IgG (H + L) conjugate Texas Red (Molecular Probes) diluted 1:3000 in TBS containing 1% goat serum. Each well was then washed three times in TBS for 7 min each. Indirect immunofluorescence of PLC{delta}1 was performed as previously described (1). In some experiments, cells incubated with 4',6-diamidino-2-phenylindole (5 µg/ml) for 5 min to assess the percentage of nuclei that appeared apoptotic. Images were captured with an AxioCam 330mA 12-bit CCD camera (Zeiss) and viewed with Carl Zeiss Axovision 3.1 software. Alternatively, fixed cells were visualized by epifluorescence microscopy (Olympus IMT-2 inverted microscope with a 100-watt mercury arc lamp), and images were taken with a Nikon Plan Fluor x40 oil objective (numerical aperture 1.3) and Olympix AstroCam (LSR). These images were processed and analyzed with Esprit imaging software (LSR). To assess the fraction of cells (scored positive or negative) having nicked DNA as a result of siRNA treatments, an in situ TUNEL Assay Cell Death Detection kit (Roche Applied Science) was used according to the manufacturer's directions.

Reverse Transcription-PCR—Total RNA was extracted from siRNA-treated C6 cultures using an RNeasy extraction kit (Qiagen) as per the manufacturer's instructions. Following purification, 0.1-1 µg of total RNA was first heated to 70 °C in the presence of random hexanucleotide primers. The RNA was then transferred to Ready-to-Go reverse transcription-PCR beads (GE Healthcare) and incubated at 42 °C for 30 min. Following the reverse transcription step, cyclin E primers (5'-GTGAAAAGCGAGGATAGCAG-3';5'-TGTTGTGATGCCATGTAACG-3') or glyceraldehyde-3-phosphate dehydrogenase primers were added, and the reactions were cDNA-amplified (18-26 cycles) in a Gene AMP PCR System 2000 thermocycler (PerkinElmer Life Sciences) with each cycle programmed for 95 °C melting for 0.5 min, 55 °C annealing for 0.5 min, and 72 °C extension for 1 min. The reaction products were separated on a 2% agarose gel that was subsequently stained with SYBR Green I (Molecular Probes, Inc., Eugene, OR). Images were recorded using a Kodak Gel Imager system, and the fluorescent bands were quantified using the Kodak gel analysis software. The cyclin E mRNA levels were normalized to expression of the glyceraldehyde-3-phosphate dehydrogenase amplicon in each sample.

Subcellular Fractionation and Lipid Analysis—Nuclei were purified as previously described (1). Following siRNA treatment for 24 h, cultures were synchronized to the G1/S boundary and labeled with 10 µCi/ml [3H]myoinositol for at least 24 h. In some cases, cultures were released from G1/S block for 3 h prior to lipid extraction. Labeled cultures were rinsed with ice-cold PBS-CaMg and then treated briefly with PBS containing 1 mM EDTA to release them from the plastic dishes. The released cells were then subjected to centrifugation at 600 x g for 5 min at 4 °C. Cells were promptly resuspended in 500 µl of prechilled hypotonic resuspension buffer (RSB; 10 mM NaCl, 1.5 mM MgCl2, 10 mM Tris-HCl, pH 7.4) on ice for 7 min. Swollen cells were then transferred to a Dounce homogenizer and lysed by 20 strokes of the glass pestle. Nuclei and debris were then layered onto a sucrose cushion (320 mM sucrose, 7.7 mM MgCl2, 2.1 mM EGTA, and 0.1 mM phenylmethylsulfonyl fluoride) and centrifuged at 300 x g for 3 min at 4 °C. The pellet was washed twice with 0.5 ml of RSB and resuspended in 750 µl of methanol, 0.1 M HCl (v/v, 1:1), placed in silicanized borosilicate glass tubes, and mixed vigorously for 30 s. The lipids were subsequently extracted as previously described (1). Lipid extracts were applied to prescored Linear KD silica plates (Whatman) that had been pretreated with 40% methanol, 1% potassium oxalate, and 1 mM EGTA in water and heat-activated. The solvent system used was chloroform/methanol/water/concentrated ammonium hydroxide (v/v/v/v; 60:47:11.3:2) (1). Areas corresponding to migration of phosphoinositide, phosphatidylinositol 4-phosphate, and PI(4,5) P2 standards were scraped into vials containing 100 µl of a mixture of methanol and 10% Nonidet P-40 in water (v/v, 1:1). 4 ml of scintillation fluid (EcoLite) was added to each vial and mixed, and the vials were counted in a liquid scintillation spectrometer. Counts/min values were normalized to total lipid phase-extractable phosphorus.

Statistical Analysis—All statistical analyses were preformed in GraphPad Prism. To determine the significance of the differences between mean values, one-way analysis of variance with Newman-Keul's post-test or Student's t test was used where appropriate (***, p < 0.001; **, p <0.01; *, p < 0.05). Fisher's exact test was used to analyze the frequency data obtained from indirect immunofluorescence images.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
siRNA-mediated Knockdown of PLC{delta}1 in Rat C6 Glioma Inhibits Proliferation—RNA interference is a sequence-specific post-transcriptional gene silencing mechanism that suppresses synthesis of a specific protein by degrading the mRNA encoding a target protein (30, 31, 34). We identified a candidate site within the rat PLC{delta}1 mRNA coding sequence (residues 152-172) and designed a specific RNA duplex ({delta}1-siRNA). In addition, we purchased three unique siRNA duplexes predicted to suppress expression of rat PLC{delta}1.


Figure 1
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  		FIGURE 1.
siRNA-mediated suppression of PLC{delta}1. A, Western blot analysis 24-72 h post-transfection with {delta}1-siRNA ({delta}1), compared with untreated (C) or cultures treated with FuGene6 alone (F). Lanes were normalized to cell number. B, representative growth curve. Cell number was measured with a hemocytometer. The data points are plotted as average of triplicate determinations, and the bars represent S.D. Suppression of PLC{delta}1 ({delta}1-siRNA) significantly reduced proliferation compared with controls untreated (Control) or transfected control siRNA (C-siRNA). Inset, growth rates were determined based on seven independent proliferation experiments and are calculated as fraction of the control untreated growth rate; ***, p < 0.001. C, Western blot analysis 48 h post-transfection with a control siRNA (Cntrl siRNA) and three commercially available siRNAs (siRNA1 to -3) predicted to suppress expression of PLC{delta}1. Lanes were normalized to total protein. D, representative phase-contrast images 48 h post-transfection with a control siRNA and three different siRNA.

 


Figure 2
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  		FIGURE 2.
Treatment of cultures with {delta}1-siRNA is not cytotoxic. A, to determine if transfection of cultures with siRNA1 resulted in cytotoxicity, cultures (n = 7) were exposed to trypan blue. No significant changes in the fraction of trypan blue-positive cells occurred between 24 and 96 h post-transfection. B, mitochondrial viability assay (n = 8). Absorbance/1 x 104 cells at 475 nM was determined as described under "Experimental Procedures." No significant changes occurred. C, cells were also stained with antibodies that recognized nicked DNA, a marker of apoptosis. Cells were scored as positive or negative, and the fraction of positive was determined.

 
Treatment of C6 glioma cultures with {delta}1-siRNA reduced the levels of PLC{delta}1 by >80% by 72 h compared with control siRNA or untreated cultures (Fig. 1A). This was associated with reduced cell numbers. Under our transfection conditions, the significant changes in the apparent growth rates (Fig. 1B, inset) were well correlated with reduced expression of PLC{delta}1 (see Fig. 1A ({delta}1-siRNA1), and see supplemental Fig. 1A). Commercially available siRNA1 and siRNA3, greatly reduced expression of rat PLC{delta}1, whereas siRNA2 was less effective (Fig. 1C). Suppression of cell proliferation mirrored their effects on PLC{delta}1 levels (Fig. 1D; see supplemental Fig. 1A), supporting the idea that slower growth is a specific effect of PLC{delta}1 suppression. As the knockdown experiments progressed, it became evident that siRNA3 rapidly reduced PLC{delta}1 levels and profoundly and rapidly reduced the fraction of cells synthesizing DNA by 48 h following treatment. In order to provide a sufficient time window for measuring the relevant cell cycle variables, we compared {delta}1-siRNA and siRNA1 in further studies, since these siRNAs took longer to suppress PLC-{delta}1 expression.


Figure 3
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  		FIGURE 3.
Suppression of PLC{delta}1 changes cell cycle distribution. A, cultures were untreated (Control), transfected with control siRNA (C-siRNA), or transfected with siRNA specific for PLC{delta}1 ({delta}1-siRNA). After 72 h, DNA content was determined by FACS analysis. Histogram data were pooled and normalized to G1-DNA content, and analysis revealed a late S phase peak (arrow) in cultures treated with siRNA1. Inset, analysis using the cell cycle algorithm (Cychlred) assigned a significant portion of this peak to G2/M. One-way analysis of variance was performed on the G2/M data; **, p < 0.01 when comparing control and siRNA1 (*, p < 0.05 when comparing C-siRNA and {delta}1-siRNA, not shown), n = 3.

 


Figure 4
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  		FIGURE 4.
Suppression of PLC{delta}1 alters DNA synthesis. A, cultures were pulse-labeled with [3H]thymidine (0.5 µCi/ml). At 72 h post-transfection, there was a significant increase in radiolabel incorporation as a result of PLC{delta}1 knockdown ({delta}1-siRNA) after 2.5 h compared with control untreated (Control) or transfection with a control siRNA (C-siRNA). *, p < 0.05 when comparing {delta}1-siRNA with both control and C-siRNA. B, triplicate cultures were blocked to the G1/S boundary and released in the presence of [3H]thymidine (0.5 µCi/ml). As a result of knockdown ({delta}1-siRNA), the initial rate of DNA synthesis was enhanced compared with control untreated cells.

 
To address the possibility that the apparent decrease in proliferation was due to an increased rate of cell death, we investigated the cytotoxicity of the various siRNA and whether treatment with these reagents induced apoptosis. Trypan blue staining revealed no significant differences between control and {delta}1-siRNA treatments up to 72 h post-transfection (Fig. 2A). The ability of treated C6 cell mitochondria to metabolize XTT, another indicator of cell viability, was also unchanged (Fig. 2B). Similar results were found using the siRNA1 and -3 (see supplemental Fig. 1, B and C). Another possible explanation for the decrease in growth rate was that the fraction of cells undergoing apoptosis had increased. 72 h post-transfection, the fraction of TUNEL-positive cells was similar between control and {delta}1-siRNA (Fig. 2C). Furthermore, there were no signs of nuclear fragmentation (visualized by 4',6-diamidino-2-phenylindole staining) in C6 cells treated with any of the siRNA (see supplemental Fig. 1D). Taken together, these data suggest that neither cytotoxicity nor increased apoptosis accounts for the decrease in cellular proliferation.

Knockdown of PLC{delta}1 Changes Cell Cycle Distribution—Since knockdown of PLC{delta}1 did not appear to enhance cytotoxicity or apoptosis, we suspected that a delay or blockage in a particular stage of the cell cycle could account for our observations. FACS analysis was used to measure the distribution of DNA content within the cell population. At 72 h post-transfection, it was evident that treatment of cultures with {delta}1-siRNA altered cell cycle distribution (Fig. 3) compared with untreated cultures and those transfected with C-siRNA. Although no significant changes were found among cultures in their G1-DNA or S phase DNA content, analysis indicated a significant decrease in the number of G2/M-DNA content cells compared with control conditions (Fig. 3, inset), suggesting a delay in S phase progression.

Knockdown of PLC{delta}1 Alters S Phase Progression—In addition to FACS analysis, we pulse-labeled cells with [3H]thymidine to measure the number of S phase cells in asynchronous cultures. At 72 h post-transfection, cultures were washed and incubated in normal growth medium with 1 µCi/ml [3H]thymidine for 2.5 h (Fig. 4A). Under these conditions, cultures transfected with {delta}1-siRNA incorporated more [3H]thymidine, indicating that either a greater number of cells synthesizing new DNA were present or that a similar proportion of S phase nuclei were present synthesizing new DNA at a faster rate or both. To determine whether the rate of DNA synthesis was altered, cells were synchronized to the G1/S boundary and released in the presence of 1 µCi/ml [3H]thymidine (Fig. 4B). DNA synthesis was measured in 2-h intervals after release from G1/S phase block. In cultures treated with PLC{delta}1-specific siRNA, DNA synthesis was significantly faster during the first 4 h.


Figure 5
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  		FIGURE 5.
Suppression of PLC{delta}1 alters S phase progression. A, to assess S phase progression, cultures were blocked to the G1/S boundary, as described under "Experimental Procedures," and then released for 3-5 h. These nine histograms are representative of two experiments. B, overlay of +5 h release histograms seen in A; the arrow indicates a delay in S phase progression. C, at 5 h after release, the relative shift of the S phase peaks were determined by calculating the distances migrated of the S phase peak as a fraction of the total distance between the G1 and G2/M peaks. Significance was determined by one-way analysis of variance; *, p < 0.05; ***, p < 0.001. D, fraction of G2/M cells 5 h after release from the G1/S boundary was calculated by estimating the area under the G2/M peak as a fraction of the total number of cells measured.

 


Figure 6
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  		FIGURE 6.
Suppression of PLC{delta}1 up-modulates cyclin E expression. A, asynchronous cultures (Asynch) were treated with C-siRNA (C) or {delta}1-siRNA (K) for the times indicated, whereas synchronous cultures were treated 24 prior to double thymidine synchronization and then released for the times indicated. Lanes were normalized to total protein. B, C6 cell cultures were also treated with C-siRNA or siRNA1, which targets a different region of PLC{delta}1 mRNA, for 72 h. Lanes were normalized to total protein. These samples were immunoblotted for PLC{delta}1, cyclin-E, and the constitutively expressed protein, β-tubulin.

 
S phase progression was also assessed by FACS analysis in synchronized cultures. Cultures blocked to the G1/S boundary were released, and their DNA content was measured 3-5 h later (Fig. 5). Suppression of PLC{delta}1 altered S phase progression. At 3 and 5 h, a greater proportion of cells had progressed into S phase from the G1/S block (Fig. 5A), in agreement with our [3H]thymidine incorporation results (see Fig. 4B, 2-6 h after G1/S release). Cultures transfected with {delta}1-siRNA, however, were significantly delayed in their progression to G2/M (Fig. 5B, arrow). In each histogram, the distance that the S phase peak migrated from the G1 peak was determined and calculated as a fraction of the distance between the G1 and G2/M peaks (Fig. 5C). In cultures treated with {delta}1-siRNA, the S phase peak migrated further compared with controls, yet significantly fewer G2/M cells were observed. An S phase peak was still observed at 5 h (Fig. 5D), supporting the idea of a delay in the completion of S phase and transition into G2/M. This delay, however, is preceded by accelerated DNA synthesis. Taken together, these results may explain the overall reduction in proliferation observed in our asynchronous culture experiments.

Knockdown of PLC{delta}1 Elevates the Levels of Cyclin E—Prior studies have shown that PLCβ and PLC{gamma} are linked to cyclin D and cyclin-dependent kinase 4/6 (Cdk4/6), important regulators of G1 progression (35, 36). One explanation for the changes in DNA synthesis and distribution of S phase cells is that PLC{delta}1 alters factors that regulate progression through this phase. Since both initial onset of DNA synthesis and exit from S phase were affected, we focused on an important regulator, cyclin E (37, 38). Treatment of cells with PLC{delta}1-specific siRNA substantially elevated the levels of cyclin E in both asynchronous (Fig. 6A) and synchronized cultures (Fig. 6B) compared with controls. On the other hand, cyclin A levels remained unaltered (data not shown). Peak expression of cyclin E occurred sooner and persisted throughout the later stages of S phase compared with controls. Moreover, elevated cyclin E levels were observed with other PLC{delta}1-specific siRNA, including siRNA1 (Fig. 6C). These results suggest that higher cyclin E levels are not the result of off-target effects of the siRNAs used here.

To address whether an increase in cyclin E mRNA levels could account for the elevated cyclin E protein levels observed, the C6 cultures were treated with control or PLC{delta}1-specific siRNA1 or -3. Under conditions where PLC{delta}1 levels were suppressed and the protein levels of cyclin E were elevated, no significant changes in the levels of cyclin E transcript were observed when cultures were treated with siRNA1 or -3 (79 ± 26 and 93 ± 21% of control, respectively (mean ± S.D. from two independent experiments performed in triplicate)).

Transient Expression of Human PLC{delta}1 Reduces Levels of Cyclin E in Cells Treated with siRNA and Synchronized to the G1/S Boundary—We co-transfected C6 glioma cells with human PLC{delta}1EGFP and the rat-specific {delta}1-siRNA to determine if we could reverse the observed cyclin E phenotype; {delta}1-siRNA was not predicted to suppress human PLC{delta}1EGFP expression (Fig. 7). Cultures were synchronized to the G1/S boundary during the transfection interval of 72 h and fixed at the G1/S boundary. Most cells expressing detectable levels of PLC{delta}1EGFP had lower levels of cyclin E as measured by indirect immunofluorescence intensity, and many of these lacked cyclin E in the nucleus (Fig. 7, A and B). When the overall intensities were measured and compared, cotransfection with PLC{delta}1EGFP caused a 33% suppression of total cellular indirect immunofluorescence, whereas cotransfection with EGFP caused an apparent fall of 12% that was not statistically significant (Fig. 7C). Analysis of the fractions of cells having high (above mean) intensity levels indicated that coexpression of PLC{delta}1EGFP with rat-specific siRNA directed toward endogenous PLC{delta}1 caused a 2.5-fold decrease in the population frequency in this group, whereas no significant fall was noted with EGFP (Fig. 7D). These results suggest that moderate overexpression of active human PLC{delta}1 counteracts the effects of the {delta}1-siRNA treatment, further demonstrating the specificity of the siRNAs used here and supporting the idea that PLC{delta}1 is an important regulator of cyclin E expression.


Figure 7
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  		FIGURE 7.
Expression of exogenous human PLC{delta}1EGFP suppresses cyclin E expression. C6 cell cultures were cotransfected with either PLC{delta}1EGFP plasmid and {delta}1-siRNA or EGFP and {delta}1-siRNA. 48 h later, the cultures were synchronized to the G1/S boundary (additional 24 h) and fixed, and cyclin E levels were imaged by indirect immunofluorescence (IMF) using a highly specific cyclin E primary antibody and Texas Red-labeled secondary antibody. Two representative epifluorescence images of the cultures, co-transfected with PLC{delta}1EGFP plasmid and {delta}1-siRNA, are shown. Image A was obtained in the Texas Red channel. The circles correspond to cells appearing in the EGFP channel image (B). C, intensities of indirect immunofluorescence of individual cells were binned into transfected and untransfected groups depending on detection of signal in the EGFP channel. This bar graph summarizes the images analyses of 410 and 124 cells, in 14 and 10 separate fields, in cultures cotransfected with PLC{delta}1EGFP plasmid and {delta}1-siRNA or EGFP and {delta}1-siRNA, respectively (***, p < 0.001; n = 2). D, indirect immunofluorescence intensities were stratified, and the frequencies of observing intensity levels above the untransfected mean were analyzed and expressed as the fraction of total cells in each group and the indicated confidence intervals (CI).

 
Suppression of PLC{delta}1 Increases Nuclear PIP2 Levels—We and others have previously reported a marked rise in nuclear PIP2 and PIP in cells synchronized to the G1/S boundary (1). To determine if suppression of PLC{delta}1 alters nuclear phosphoinositides, C6 glioma cells were treated with {delta}1-siRNA. Twenty-four hours later, cultures were labeled with [3H]myoinositol for 24 h and blocked at the G1/S boundary. Suppressing PLC{delta}1 significantly increased levels of nuclear phosphoinositides, particularly PIP2 and PIP, compared with control cultures (Fig. 8A). By contrast, no significant changes in whole cell PIP2 and PIP were evident (Fig. 8B). These data support a role for PLC{delta}1 in the metabolism of nuclear phosphoinositides at the G1/S boundary.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Suppression of endogenous PLC{delta}1 in rat C6 glioma cultures significantly reduced cellular proliferation. FACS analyses and [3H]thymidine labeling suggest that S phase progression and exit were delayed, resulting in a reduced population of G2/M cells, which may account for the overall decrease in growth rate. In an attempt to explain this phenomenon, we investigated whether siRNA-mediated suppression of PLC{delta}1 altered key regulators of S phase progression (37, 38).

In addition, we found that increased expression of cyclin E could be prevented by expression of human PLC{delta}1 in G1/S-blocked cells treated with rat-specific {delta}1-siRNA. On the other hand, cotransfection with EGFP failed to show a similar effect. Taken together, our results demonstrate an important and specific role for this PLC in cell cycle regulation.

We previously demonstrated that PLC{delta}1 accumulates in the nucleus at the G1/S transition and suggested that this protein modulates the levels of nuclear PI(4,5) P2 at this transition (1). Here, we also demonstrate that suppression of PLC{delta}1 leads to a significant increase in the levels of nuclear PIP2 and PIP, supporting the idea that PLC regulates metabolism of nuclear phospholipids at the G1/S boundary. Indeed, cotransfection with active PLC{delta}1 of C6 cells treated with {delta}1-siRNA reverses the increase in cyclin E levels at a point in the cycle where PLC{delta}1 is mainly localized to the nucleus (1).

Although our previous work (1) and results here demonstrate that most of PLC{delta}1 is localized to the nucleus at G1/S, we have yet to directly address whether the changes in cyclin E levels or the effects on cell cycle are due to nuclear PLC{delta}1. Nonetheless, our results are consistent with a role for the nuclear form of this PLC in S phase progression, hydrolysis of nuclear PI(4,5) P2, and modulation of cyclin E expression.


Figure 8
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  		FIGURE 8.
Suppression of PLC{delta}1 elevated nuclear phosphoinositides at the G1/S boundary. A, cultures were transfected with {delta}1-siRNA (open bars) and blocked to the G1/S boundary (black bar) while labeled with [3H]myoinositol. Control cultures (filled bars) were not transfected with siRNA. Nuclei were isolated by osmotic swelling and extracted as described under "Experimental Procedures." The data are representative of two independent experiments; the data are plotted as the average of triplicate determinations, and the bars represent S.D.; **, p < 0.01; *, p < 0.05. Counts/min values were normalized to lipid extractable phosphorus. (Note that the right y axis is 10-fold greater and refers only to phosphatidylinositol (PI), whereas the left y axis refers to both PIP2 and PIP). B, whole cell PIP2 levels as a function of PLC{delta}1 suppression. Cultures were transfected with 160 nM {delta}1-siRNA or untreated while being labeled with [3H]myoinositol for 72 h. Whole cell phosphoinositide levels were also determined as previously described (1), and no significant changes were observed.

 
The simplest explanation for an accelerated entry into S phase and altered DNA synthesis is a perturbation of S phase regulators, such as cyclin E, cyclin A, and their common regulatory kinase Cdk2 (37, 38). Cyclin E levels are regulated by E2F transactivation of transcription (39) and ubiquitin-mediated destruction (40, 41). Our results show that PLC{delta}1 does not modulate cyclin E transcription; therefore, it is likely that the rate of degradation of cyclin E is reduced, leading to an increase in protein level. Interestingly, genetic evidence has linked PLC1, a homolog to PLC{delta}1, to proteolytic degradation of C-type cyclins in yeast (42).

There appear to be two pathways that regulate cyclin E degradation: a phosphorylation-independent mechanism that regulates rapid turnover of a free pool of cyclin E and a phosphorylation-dependent mechanism that appears to contribute to the ubiquination of the more stable cyclin E in complex with Cdk2 (43).

Since Cdk2 associates with both cyclins A and E, suppression of PLC{delta}1 could alter the ratio of Cdk2 bound to cyclins E and A. Thus, increasing cyclin E levels and presumably its complex with Cdk2 might result in less Cdk2 available to bind cyclin A for appropriate S phase exit (44). Overexpression of cyclin E in BK cells, however, was not found to alter the amount of Cdk2 associated with this cyclin (44). Other questions that remain are whether PLC{delta}1 plays a direct role in the degradation of cyclin E, whether it occurs through a phosphorylation-dependent or -independent mechanism, and whether metabolism of nuclear PIP2 is required.

Since we find that suppression of PLC{delta}1 leads to an increase in the levels of nuclear PIP2, these levels could alter transcription and thereby affect cyclin E degradation, albeit indirectly. Previous studies have demonstrated an important role for phosphatidylinositol 5-phosphate-binding proteins that regulate transcription in the nucleus (reviewed by Jones and Divecha (45)). PIP2 and enzymes that metabolize this lipid have been localized to the nuclear matrix, envelope, nucleoli, and nuclear speckles, the latter involved in mRNA splicing and RNA modification (46-49).

Increased nuclear levels of this lipid could also effect chromatin remodeling or histone modifications that alter transcription. In vitro studies have shown that PI(4,5) P2 binds to actin-related proteins that are part of the BAF (Brg- or Brm-associated factors) complex (50), and others demonstrated that PIP2 is sufficient to target the BAF complex to chromatin in vitro (51). PI(4,5) P2 also binds to the C-terminal tails of histones H1 and H3 (52), suggesting a role in histone regulation. The hydrolysis of nuclear PIP2 associated with histones could modulate transcription (45, 52). Consistent with this idea, in vitro studies have also shown that the presence of PI(4,5) P2 counteracts H1-mediated basal transcription by RNA polymerase II (52).

High levels of cyclin E are typically associated with unregulated proliferation (53), and previous work has demonstrated that increased cyclin E leads to an accelerated progression into S phase when degradation is hindered (54-57). Under some circumstances, overexpression of this protein results in both enhanced G1/S transition and delayed exit from S phase (44), comparable with the phenotype we observed here.

Other mammalian PLC isoforms have been shown to promote expression and/or activity of cell cycle regulators. FGF-2-mediated PLC{gamma}1 activation promotes up-regulation and nuclear import of Cdk4 and stimulates nuclear export of the Cdk inhibitor p27kip1 (36). PLCβ1a and PLCβ1b, which both possess functional nuclear import sequences, promote cyclin D3-Cdk4 complex formation and hyperphosphorylation of retinoblastoma protein, a critical regulator of G1/S transition (35). Our study points to a comparable role for PLC{delta}1.

It is possible that generation of local second messengers, such as diacylglycerol, activates protein kinase C (16) and modulates cell cycle progression. Indeed, nucleus-localized PLCβ1 has been shown to regulate IGF-1-stimulated proliferation of Swiss 3T3 fibroblasts and the commitment of MEL cells to proliferate or differentiate through the generation of diacylglycerol and activation of nuclear protein kinase C isoforms (18, 58, 59). In regenerating rat liver nuclei, PLCβ localizes to chromatin that is actively incorporating bromodeoxyuridine, whereas PLC{gamma}1 is associated with interchromatin regions and the nuclear envelope (60). These authors have also suggested that nuclear PLCβ plays a role in DNA synthesis, whereas PLC{gamma}1 is more likely involved at the G2/M transition and nuclear lamin phosphorylation (60). Recent work shows that PLC{gamma}1 is also critical for reassembly of the nuclear envelope from PI(4,5) P2-enriched vesicles.

Our results place PLC{delta}1 at an unidentified control point, somewhere in the S phase. The timing of its peak nuclear localization to the G1/S boundary coincident with peak nuclear PI(4,5) P2 levels (1) suggests that hydrolysis of this phosphoinositide is required for normal S phase progression, possibly through enhanced degradation of cyclin E.


   FOOTNOTES
 
* The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

Formula The on-line version of this article (available at http://www.jbc.org) contains supplemental Fig. 1. Back

1 To whom correspondence should be addressed: Dept. of Anesthesiology, Health Sciences Center Level 4, Rm. 075, SUNY @ Stony Brook, NY 11794-8480. Tel.: 631-444-8178; Fax: 631-444-2907; E-mail: mrebecchi{at}notes.cc.sunysb.edu.

2 The abbreviations used are: PIP2 or PI(4,5) P2, phosphatidylinositol 4,5-bisphosphate; XTT, sodium 3'-[1-[(phenylamine)-carbonyl]-3,4-tetrazolium]-bis(4-methoxy-6-nitro)benzenesulfonic acid hydrate; PLC, phospholipase C; PBS, phosphate-buffered saline; siRNA, small interfering RNA; EGFP, enhanced green fluorescent protein; TBS, Tris-buffered saline; TUNEL, terminal deoxynucleotidyl transferase-mediated nick end labeling; FACS, fluorescence-activated cell sorting; PIP, phosphatidylinositol phosphate. Back


   ACKNOWLEDGMENTS
 
We thank the Stony Brook Immunology Clinic for assistance with flow cytometry analysis and Pooja Kondabolu for technical assistance. We thank Dr. Peter Glass and the Stony Brook University Anesthesiology Department for unwavering support and encouragement.



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