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Originally published In Press as doi:10.1074/mcp.M600272-MCP200 on November 22, 2006.
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Molecular & Cellular Proteomics 6:319-332, 2007.
© 2007 by The American Society for Biochemistry and Molecular Biology, Inc.

Research

A Functional Genomics Analysis of the B56 Isoforms of Drosophila Protein Phosphatase 2A*,S

Wei Liu, Adam M. Silverstein{ddagger}, Hongjun Shu§, Bobbie Martinez and Marc C. Mumby

From the Department of Pharmacology, The University of Texas Southwestern Medical Center, Dallas, Texas 75390-9041


   ABSTRACT
TOP
 ABSTRACT
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Members of the B56 family of protein phosphatase 2A (PP2A) regulatory subunits play crucial roles in Drosophila cell survival. Distinct functions of two B56 subunits were investigated using a combination of RNA interference, DNA microarrays, and proteomics. RNA interference-mediated knockdown of the B56-1 subunit (PP2A-B') but not the catalytic (mts) or B56-2 subunit (wdb) of PP2A resulted in increased expression of the apoptotic inducers reaper and sickle. Co-knockdown of B56-1 with reaper, but not with sickle, reduced the apoptosis caused by depletion of the B56 subunits. Two-dimensional gel electrophoresis and mass spectrometry identified proteins modified in cells depleted of PP2A subunits. These included generation of caspase-dependent cleavage products, increases in protein abundance, and covalent modifications. Results suggested that up-regulation of the ribosome-associated protein stubarista can serve as a sensitive marker of apoptosis. Up-regulation of transcripts for multiple glutathione transferases and other proteins suggested that loss of PP2A affected pathways involved in the response to oxidative stress. Knockdown of PP2A elevated basal JNK activity and substantially decreased activation of ERK in response to oxidative stress. The results reveal that the B56-containing isoform of PP2A functions within multiple signaling pathways, including those that regulate expression of reaper and the response to oxidative stress, thus promoting cell survival in Drosophila.

Protein phosphatase 2A (PP2A)1 is a major class of serine/threonine phosphatase that plays a myriad of roles in cellular signaling. The term PP2A refers to a group of enzymes composed of a common catalytic subunit, which forms oligomeric complexes with a large and diverse group of proteins (1–3). The most common forms of PP2A contain a core dimer, composed of the catalytic subunit and a scaffold protein termed the A subunit. The scaffold subunit mediates formation of heterotrimeric holoenzymes by binding to additional regulatory subunits. There are four characterized families of PP2A regulatory subunits that target PP2A to specific substrates and signaling complexes. An important class of regulatory subunit is encoded by the PPP2R5 gene family, which has also been referred to as B56, B', and PR61. The mammalian B56 gene family contains five members, each of which is processed into multiple splice variants. Members of the B56 family target PP2A to a variety of cellular processes including transformation (4, 5), Wnt signaling (6–8), cell polarization (9), and circadian rhythm (10).

PP2A plays both positive and negative roles in pathways that regulate cell survival and apoptosis. Induction of apoptosis by the second messenger ceramide, a membrane sphingolipid, correlates with stimulation of a fraction of PP2A associated with mitochondria in HL60 cells and the murine interleukin-3-dependent cell line NSF/N1.H7 (11). The function of PP2A in ceramide-induced apoptosis involves inhibition of the antiapoptotic activity of Bcl2 by dephosphorylation of serine 70. The ceramide-enhanced dephosphorylation of this site is associated with recruitment of PP2A to Bcl2 via interaction with the B56{alpha} regulatory subunit (12). PP2A also plays a positive role in the apoptosis caused by withdrawal of survival factors from interleukin-3-dependent FL5.12 cells (a murine pro-B-cell lymphoid cell line). In this case, PP2A associates with the Bad proapoptotic protein and dephosphorylates serine 112. Dephosphorylation of this site leads to subsequent dephosphorylation of additional sites, dissociation of 14-3-3 proteins, and unmasking of the apoptotic activity of Bad (13, 14). PP2A also plays a positive role in tumor necrosis factor {alpha}-induced apoptosis of rat IEC-6 cells through a process that involves both Bcl-2 and Bad (15).

In addition to roles in promoting apoptosis in response to specific stimuli, PP2A plays a more fundamental role in promoting cell survival. Silencing of PP2A by RNA interference results in apoptosis of multiple cell types. Knockdown of PP2A in the Schneider 2 Drosophila cell line results in an apoptotic response mediated by the combined loss of the two Drosophila B56 subunits (16, 17). RNAi-mediated knockdown of PP2A also causes apoptosis in the rat PC12 neuronal cell line (18). The ability of PP2A to prevent apoptosis in PC12 cells requires multiple classes of regulatory subunits including the B56 family. The catalytic and scaffold subunits as well as members of the PPP2R2 (B), PPP2R3 (PR72), and PPP2R5 (B56) regulatory subunits of human PP2A have all been identified as prosurvival proteins in an RNAi screen (19). The ability of PP2A to play either positive or negative roles in apoptosis is likely to depend on individual holoenzymes and on the organism, cell type, and apoptotic stimulus.

A deficiency in understanding the functions of PP2A in cellular regulatory processes, including those that regulate cell survival and apoptosis, is a lack of information about the substrates and signaling pathways that are targeted by different forms of the enzyme. In mammalian cells, the problem is complicated by a large number of regulatory subunit isoforms and splice variants. The simplified repertoire of PP2A regulatory subunits in Drosophila melanogaster provides a simpler system in which individual isoforms can be investigated. Previous studies showed that the Drosophila B56 subunits (B56-1/PP2A-B' and B56-2/widerborst) are required for cell survival and prevent programmed cell death (16, 17). To gain insights into the molecular functions of these subunits, we used a combination of RNA interference, DNA microarrays, and proteomics to identify proteins and signaling pathways targeted by PP2A and the B56 family of regulatory subunits.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Culture and RNA Interference—
Schneider S2 cells were maintained in Drosophila serum-free medium (Invitrogen) and seeded at 1 x 106 cells/well in 6-well culture plates for RNAi experiments as described previously (16). Twenty-four hours later, 15 µg of dsRNA was added to the culture medium. Double-stranded RNA corresponding to enhanced green fluorescent protein (EGFP) and the catalytic, B56-1, B56-2, and PR72 subunits of Drosophila PP2A were prepared as described previously (16). For knockdown of reaper and sickle mRNA, regions of 500–800 bp were amplified by PCR using gene-specific primers from cDNA prepared from S2 cells using similar methods. Cells were harvested 3–5 days after addition of dsRNA. In experiments where the B56-1 and -2 subunits were knocked down simultaneously, twice the normal amount of EGFP dsRNA was used in the control treatments. In some experiments the cells were incubated with Z-VAD-fmk (30 µM final concentration) during the dsRNA treatment to block apoptosis. In some two-dimensional gel electrophoresis experiments, cells that had not been treated with dsRNA were incubated with actinomycin D (20 nM) for 24 h to induce apoptosis.

Preparation of Total Protein and RNA—
Total protein and RNA were isolated from dsRNA-treated S2 cells using the TriPure reagent (Roche Diagnostics) following the manufacturer’s instructions. Total protein extracts for analysis by two-dimensional gel electrophoresis were prepared from three wells of S2 cells treated with the same dsRNA. The protein pellet was dried in a vacuum concentrator and dissolved in 100 µl of 2-D rehydration buffer (7 M urea, 2 M thiourea, 4% (w/v) CHAPS, 50 mM DTT, 3% (v/v) Ampholines, 1 mM sodium orthovanadate, 1 µM okadaic acid, and 1x Complete Mini protease mixture (Roche Applied Science)). The concentration of the solubilized protein was determined by Amido Black protein assay (20).

Total RNA was isolated from the same samples by precipitation of the aqueous phase from the phenol-chloroform extraction with isopropanol. The RNA precipitate was washed once in 75% ethanol, dried at room temperature, and resuspended in RNase-free water. Contaminating DNA was removed by treating with DNase I (Ambion). The RNA quality was examined by analysis on a 1% agarose gel to determine the presence of intact ribosomal RNA. The 260/280 ratio was then measured, and only samples with ratios between 1.8 and 2.1 were used for subsequent analyses.

DNA Microarray Analysis—
Preparation of cRNA, hybridization to Drosophila Genome Chip arrays (DrosGenome 1), and scanning were performed according to the manufacturer’s protocol (Affymetrix, Santa Clara, CA). Briefly 5 µg of RNA was converted into cDNA by reverse transcription using the SuperScript Choice cDNA Synthesis kit (Invitrogen). Biotin-labeled cRNA was generated from the double-stranded cDNA using the BioArray High Yield RNA Transcript Labeling kit (Enzo Life Sciences, Farmingdale, NY). After purification with Rneasy spin columns (Qiagen, Valencia, CA), the labeled cRNA was fragmented and hybridized to the arrays in the University of Texas Southwestern Medical Center Microarray Core Facility. The hybridized chips were stained and scanned using a GeneArray scanner (Affymetrix). The probe-level data from 16 arrays were analyzed using GeneSpring® software, version 7.2 (Silicon Genetics). Following import of the Affymetrix CEL files into GeneSpring, background correction, normalization, and summarization were carried out using the Robust Multichip Average with GC content background correction (GC-RMA) algorithm. The expression of each gene was then normalized to its median across all chips. For those knockdown experiments that were replicated (triplicates for EGFP, catalytic subunit, B56-1, and B56-1 and -2 and duplicates for B56-2 alone), the mean signal was used to compare transcript levels in the PP2A dsRNA-treated samples with those from the control EGFP dsRNA-treated samples. Transcripts whose levels were reproducibly changed were identified using the following criteria. 1) The change in normalized mean signal between the EGFP dsRNA-treated and PP2A dsRNA-treated samples was at least 2-fold. 2) The difference between EGFP and PP2A was statistically significant using one-way parametric analysis of variance with a p value cutoff of 0.05 (false positive rate of 5%). The changes in transcript levels are expressed as the -fold change in signal between PP2A dsRNA samples and the EGFP dsRNA samples.

RT-PCR and Real Time Quantitative Reverse Transcription PCR—
RT-PCR to detect knockdown of PP2A subunit mRNA was performed as described previously (16) using the superscript one-step RT-PCR system (Invitrogen). For real time quantitative reverse transcription PCR, 2 µg of total RNA, isolated as described for the microarray experiments, was used in reverse transcription reactions using TaqMan reverse transcription reagents (Applied Biosystems). The PCRs were performed in triplicate using the SYBR Green PCR kit (Applied Biosystems) according to the manufacturer’s instructions. Fluorescence changes during real time PCR were measured with an ABI PRISM 7900 sequence detection system or an ABI 7500 qRT-PCR instrument (Applied Biosystems). All gene-specific primers were designed using Primer Expression software (Applied Biosystems). A primer pair against Drosophila actin was used as a control for all experiments. The following primer pairs were used for qRT-PCR: reaper forward, 5'-CACAGTGGAGATTCCTGGCC; reaper reverse, 5'-TGTACTGGCGCAGGGTTTC; sickle forward, 5'-GTGCAAGGTCCTGAAGCAAT; sickle reverse, 5'-GTGGCCTTTAGTTTGCTGGA; GstD2 forward, 5'-GCCGCACGGTCATCATG; GstD2 reverse, 5'-TGGTGTTCAGTAGCTTCTTGTTCAG; CG5224 forward, 5'-CCCCTGTCGTGCTGTTCTG; CG5224 reverse, 5'-TTACGTTGACCAGTCGCAAGTC; GstE6 forward, 5'-TTGCCTATTTGGTCTCGAAATATG; GstE6 reverse, 5'-ACAGCCCGCTTGAGAGGAT; GstE7 forward, 5'-ACCAGTTCGTGCCGTCAAAT; GstE7 reverse, 5'-CCGAGTGTTTACCTCCACGAAT; actin forward, 5-CGCGAAAAGATGACTCAGATTATG; actin reverse, 5'-CCGCTTGGATGGCAACAT. The amount of each target mRNA relative to actin mRNA was calculated using the comparative CT ({Delta}CT) method. The difference in the amounts of target mRNA between the EGFP dsRNA-treated and PP2A subunit dsRNA-treated cells was calculated as described in Applied Biosystems User Bulletin 2.

Two-dimensional Gel Electrophoresis—
Extracted S2 cell protein (400 µg) in 225 µl of 2-D rehydration buffer was used to rehydrate 11-cm immobilized pH 4–7 gradient gels (pH 4–7 Immobiline DryStrips, Amersham Biosciences) overnight following the manufacturer’s protocol. After rehydration, the strips were placed in a Multiphor II system (Amersham Biosciences) and focused according to the following program: 10 V-h at 500 V, 500 V-h at 300 V, 17,500 V-h at 3500 V, 25,000 V-h at 3500 V, and 17,000 V-h at 300 V. The focused strips were equilibrated for 10 min in DTT equilibration solution (50 mM Tris-HCl, pH 8.8, 38% glycerol, 6 M urea, 3% SDS, 5 mg/ml dithiothreitol) followed by 10 min in iodoacetamide equilibration solution (same as above but with 45 mg/ml iodoacetamide replacing the DTT). The equilibrated strips were placed on top of 12.5% IPG+1 Criterion gels (Bio-Rad), and the focused proteins were resolved by SDS-polyacrylamide gel electrophoresis. For some experiments, high resolution 2-D gel electrophoresis was carried out using the same methods but with Immobiline IPG strips that were 18 cm long and second dimension SDS gels that were 20 cm long. Following electrophoresis the gels were stained with SYPRO Ruby dye (Molecular Probes), and images were acquired with a Typhoon imaging system (Amersham Biosciences). The 2-D gel images were analyzed with Ettan Progenesis software (Amersham Biosciences). After spot detection, background subtraction, and normalization to total spot intensity, gel images from EGFP control and PP2A dsRNA-treated samples were warped to align matched protein spots on different gels. The software was then used to identify and quantitate spots whose normalized intensities were different in control and treated samples as well as spots that were uniquely present (non-matched) in individual samples. The 2-D gel analysis was carried out in four to five separate experiments, and those spots whose intensities were altered by at lease 2-fold in three or more experiments were considered to be reproducibly affected by loss of PP2A subunits.

Mass Spectrometry—
Protein spots of interest were excised from SYPRO Ruby-stained 2-D gels illuminated on a UV light box. Peptides for each spot were generated by in-gel digestion with trypsin as described previously (21). The dried peptides were desalted with 0.1% TFA using ZipTips (Millipore) and eluted with 2,5-dihydroxybenzoic acid matrix solution (Agilent) directly onto MALDI sample plates. MS and MS/MS data were acquired by MALDI-Q-TOF mass spectrometry using a QSTAR Pulsar-I quadrupole time-of-flight tandem mass spectrometer (Applied Biosystems). The MS and MS/MS data were searched against the National Center for Biotechnology Information (NCBI) non-redundant Drosophila protein database using Knexus software (Genomic Solutions, Inc.) as described previously (21). To be considered a positive identification, the mass spectra of at least three different peptides from each spot had to match the database entry for that protein.

Caspase Assays—
Assay for caspase-3-like enzyme activity was performed using the ApoAlert Caspase Colorimetric Assay kit (Clontech) as described previously (16). Caspase activities were normalized for total protein recovered in each sample and expressed as the relative activity in PP2A dsRNA-treated cells relative to control cells treated with EGFP dsRNA. The values are reported as the mean ± S.D. from three independent experiments.

Analysis of Drosophila ERK, JNK, and p38 MAP Kinase Activation—
Lysates from S2 cells treated with dsRNA were assayed for activation of the Drosophila homologs of the ERK (rolled), JNK (basket), and p38 MAP kinases (Mpk2 and p38b) using phosphospecific antibodies. S2 cells were treated with control or PP2A dsRNA for 4 days and then subjected to oxidative or osmotic stress. The dsRNA-treated cells were harvested and incubated for 10 min in medium containing no additions, 2 mM hydrogen peroxide, 0.3 M NaCl, or 0.3 M sorbitol. The treated cells were collected by centrifugation for 30 s at 18,000 x g, lysed in 5% SDS sample buffer (62.5 mM Tris-HCl, pH 6.7, 10% glycerol, 5% SDS, 0.1% bromphenol blue, 1% 2-mercaptoethanol), and heated at 95 °C for 10 min. Protein concentrations were determined by Amido Black assay (20), and 40 µg of protein was resolved by SDS-PAGE. Proteins were transferred to nitrocellulose membranes and immunoblotted with antibodies against MAP kinases and PP2A subunits. Antibodies against the catalytic and B56-1 subunits of PP2A were used as described previously (16). Antibodies that recognize Drosophila MAP kinases were obtained from the following sources: anti-phospho-p44/42 MAP kinase (Thr-202/Tyr-204) rabbit polyclonal, anti-phospho-p38 MAP kinase (Thr-180/Tyr-182) rabbit polyclonal, and anti-phospho-SAPK/JNK (Thr-183/Tyr-185) rabbit polyclonal antibodies were from Cell Signaling Technology; anti-ERK, anti-p38, and anti-JNK1 (FL) rabbit polyclonal antibodies were from Santa Cruz Biotechnology. Each of these antibodies has been shown to cross-react with the Drosophila homologs of these protein kinases (22). Immunoblotting was performed according to the manufacturer’s protocol.


   RESULTS
TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Effects of PP2A Subunit Knockdown on S2 Cell Transcripts—
Genome-wide screens for changes in gene expression were carried out to identify signaling pathways altered by knockdown of the catalytic and B56 subunits of PP2A. Affymetrix Drosophila Genome Array chips representing 13,500 genes were used to identify transcripts that were differentially expressed in S2 cells in which these subunits had been knocked down by RNAi. To detect transcripts whose changes were due to loss of PP2A subunits and not simply to dsRNA, RNA from cells treated with a control dsRNA (derived from the sequence of EGFP) was used as the base line for all comparisons. Genes whose transcript levels were reproducibly altered by PP2A dsRNA treatment were identified as those whose levels changed by a least 2-fold relative to the EGFP dsRNA-treated controls and were statistically significant according to one-way analysis of variance with a p value cutoff of 0.05.

Knockdown of the PP2A catalytic subunit resulted in up-regulation of 30 transcripts and down-regulation of 32 transcripts. The largest group of affected transcripts with a common biological function was composed of six up-regulated genes involved in the insect defense response. Five of these genes (CG5224, GstE3, GstE6, GstE7, and GstE8) encode Drosophila glutathione S-transferases, which play roles in insecticide metabolism and protection from oxidative stress. Another defense response gene, the CG18522 oxidoreductase, and two other oxidoreductases (Cyp6a8 and CG2064) were also up-regulated. Transcripts that were down-regulated included a number of genes involved in signal transduction including pebble (a guanyl-nucleotide exchange factor), Cdc2 (a cell cycle kinase), Toll (a receptor involved in activation of the immune response), Serrate (a ligand for the Notch receptor), and CrebA (a cAMP-dependent protein kinase-regulated transcriptional co-activator). The complete set of transcripts altered by knockdown of the PP2A catalytic subunit can be found in Supplemental Table S1.

Because depletion of the PP2A catalytic subunit induces apoptosis of Drosophila S2 cells, it was important to differentiate transcripts whose expression was altered by the depletion of PP2A from those altered by the induction of apoptosis. RNA interference experiments were consequently carried out in the presence of the caspase inhibitor Z-VAD-fmk. Exposure of S2 cells to Z-VAD-fmk inhibited apoptosis caused by knockdown of the catalytic subunit as well as apoptosis induced by actinomycin D (data not shown). Treatment of Z-VAD-fmk blocked up-regulation of two transcripts (CG14291 and CG13510) and down-regulation of five transcripts (CG3739, CG10026, CG8468, sugarless, and CG6199). The results indicated that most of the changes in transcript levels observed in cells depleted of the PP2A catalytic subunit were either independent of apoptosis or caused by changes that occurred upstream of caspase activation (Supplemental Table S1).

Knockdown of the B56-1 subunit resulted in up-regulation of 57 transcripts and down-regulation of 36 transcripts. Although there was limited overlap with transcripts altered by knockdown of the catalytic subunit, there were similarities in the classes of transcripts up-regulated (Supplemental Table S5). The largest group of functionally related transcripts up-regulated by knockdown of the B56-1 subunit were eight genes involved in the defense response including the glutathione S-transferases CG5224 and GstD2 (Supplemental Table S2). Knockdown of B56-1 also led to increases in several transcripts that are up-regulated during the Drosophila innate immune response including Attacin-C and Attacin-B, CG16713, reaper, and amphiphysin (23, 24). The most highly up-regulated transcript, CG16713, was increased 45-fold and encodes a Kunitz-type endopeptidase inhibitor regulated by the Toll pathway. Knockdown of B56-1 led to altered transcription of several genes that have been shown, or inferred, to play a role in apoptosis including CG11593, reaper, mub, and Bruce as well as genes involved in autophagy including Atg1and Cecropin B. Effects of knockdown of the B56-2 subunit were relatively modest compared with knockdown of either the catalytic or B56-1 subunits. Knockdown of B56-2 affected the levels of 21 transcripts: 17 were up-regulated and four were down-regulated (Supplemental Table S3). None of the transcripts altered by B56-2 knockdown corresponded to transcripts altered by depletion of either the catalytic or B56-1 subunits (Supplemental Table S5).

A total of 111 transcripts were altered by simultaneous knockdown of both the B56-1 and -2 subunits. Sixty-three transcripts were up-regulated, and 48 were down-regulated (Supplemental Table S4). The up-regulated transcripts included those for 10 genes involved in the defense response including three glutathione S-transferases (GstD2, CG5224, and CG6662), transferrin 1, superoxide dismutase (CG9027), and a heat shock protein (Hsp67Bc). Six genes with oxidoreductase activity (Cyp6d2, CG30069, CG18522, CG2064, Fdxh, and CG18240) were also up-regulated by knockdown of both B56 subunits. The largest group of functionally related transcripts were 12 genes with biological functions in proteolysis. This included down-regulation of nine transcripts (Neu3, CG30090, CG5896, Tequila, CG4804, CG6687, CG14526, CG14527, and CG30086) and up-regulation of three transcripts (skpC, CG2658, and refractory to sigma P). Twenty of the transcripts altered by knockdown of both B56 subunits were affected to a similar extent by knockdown of the catalytic subunit (Table I). A number of the common transcripts had molecular functions indicative of a potential role in oxidative stress including the glutathione S-transferase CG5224, ferrochelatase, and two oxidoreductases (CG18522 and CG2604). Blocking apoptosis with Z-VAD-fmk inhibited the changes in 11 transcripts (10%) altered in response to combined exposure to B56-1 and -2 dsRNA (Supplemental Table S4).


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  		TABLE I Common transcripts altered by knockdown of the catalytic subunit and by double knockdown of B56-1 and B56-2

Genes that did not have a molecular function annotation in the Drosophila genome database are indicated by NA. C Sub, catalytic subunit.

 
Identification of Proteins Altered in Response to Knockdown of PP2A Subunits—
Knockdown of PP2A was likely to affect protein post-translational modifications, which cannot be detected using DNA microarrays. Therefore, a proteomics approach was used to identify changes resulting from the loss of PP2A subunits. Total cell protein was isolated from S2 cells treated with control or PP2A dsRNA and resolved by 2-D electrophoresis. Protein spots whose intensities differed between the control and treated samples were identified using 2-D electrophoresis analysis software. A representative 2-D gel from Drosophila S2 cells is shown in Fig. 1A. Protein spots P2, P3, P7, and P9–P12 were present in control and PP2A dsRNA-treated samples, but their intensities were up-regulated in cells treated with catalytic subunit or B56-1&2 dsRNA. A second set of proteins consisted of spots P1, P4, P5, P6, and P8, which appeared as new spots in PP2A dsRNA-treated cells (Fig. 1, B–D). The pattern of changes in these spots was the same in cells depleted of either the catalytic subunit or both B56 subunits. A summary of the protein changes caused by knockdown of the PP2A catalytic subunit is presented in Table II.


Figure 1
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  		FIG. 1. 2-D gel analysis of proteins from S2 cells treated with dsRNA for the catalytic or both B56-1 and B56-2 subunits of PP2A. S2 cells were exposed to control EGFP (GFP) dsRNA or dsRNA corresponding to the catalytic (C sub) or both B56 subunits (B56-1&2) of Drosophila PP2A for 4 days. Total cell protein was resolved by 2-D gel electrophoresis, stained with SYPRO Ruby, and imaged with a fluorescence scanner. A, the image of protein spots resolved by 2-D gel electrophoresis from cells exposed to catalytic subunit dsRNA. The pH range is shown at the top, and the migration of the indicated molecular mass markers (in kDa) is shown on the right. Arrows indicate the position of selected protein spots (P9–P12) and two reference proteins identified by mass spectrometry (actin5C and hsc70-4). Boxes indicate areas of the gel that are shown in B–D. B, images of the area indicated by box B in A of 2-D gels from cells exposed to control dsRNA, dsRNA for the catalytic subunit, dsRNA for both B56 subunits, or actinomycin D (Act D) in the presence or absence of Z-VAD-fmk (+ZV). The positions of selected spots are indicated by arrows. C, images of the area indicated by box C in A. D, images of the area indicated by box D in A.

 

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  		TABLE II Summary of proteins altered by knockdown of the PP2A catalytic subunit

The listed proteins correspond to the spots identified in Figs. 1 and 2. The table lists the spot number, the gene name, the -fold change in protein (Protein), the -fold change (±S.D.) in transcript (mRNA) levels (determined by microarray analysis), and the Gene Ontology molecular function and biological process (where available). A, spot that appeared; D, spot that disappeared; ND, not determined.

 
In an effort to differentiate protein changes that occurred as a result of apoptosis from those related more directly to effects of PP2A, the 2-D gel experiments were repeated in the presence of the caspase inhibitor Z-VAD-fmk. The protein spot patterns were also compared with those generated following induction of apoptosis with actinomycin D. Exposure of cells to Z-VAD-fmk blocked the changes in spots P1, P4, P5, P6, P8, P9, and P11, and the dsRNA-induced changes in these spots were mimicked by actinomycin D (Fig. 1, B–D). The effects of actinomycin D were also blocked by Z-VAD-fmk (not shown). In contrast, changes in protein spots P2, P3, P7, P10, and P12 were not inhibited by Z-VAD-fmk nor were they induced by treatment with actinomycin D.

Proteins altered by knockdown of the PP2A subunits were identified by mass spectrometry. Spots P4 and P8 were identified as Rho GDP dissociation inhibitor (RhoGDI) and lamin (Table II). The predicted molecular masses of Drosophila RhoGDI and lamin are 23 and 70 kDa. However, their mobilities in the second dimension SDS gel corresponded to proteins with masses of 20 and 28 kDa, respectively. The lower apparent size and the fact that appearance of these spots was dependent on the induction of apoptosis suggested these spots were caspase cleavage products. Spots P5 and P6 were both identified as Drosophila initiation factor 5 (eIF5). The relative mobilities of the eIF5 spots in the second dimension SDS gel (Mr = 18–20,000) were lower than the predicted molecular mass of Drosophila eIF5 (52 kDa). Formation of these spots was blocked by Z-VAD-fmk and induced by actinomycin D suggesting that Drosophila eIF5 is also a caspase substrate. Spot P1 was identified as the Drosophila ribosome-associated protein stubarista (sta). Although the mobility of stubarista in the second dimension was consistent with its predicted molecular mass (30.2 kDa), the observed isoelectric point (pI = 5.5) was distinct from that predicted from the primary sequence (pI = 4.8), suggesting the protein was covalently modified. The appearance of the stubarista spot was blocked by the caspase inhibitor and induced by treatment with actinomycin D suggesting that induction was dependent on apoptosis. No significant differences in the transcript levels for any of these genes were observed in the microarray analyses (Table II).

The appearance of the stubarista protein spot appeared to be a sensitive marker for S2 cell apoptosis. Two-dimensional fluorescence difference gel electrophoresis was used to quantitate the appearance of stubarista following knockdown of PP2A subunits (Supplemental Fig. S1). Depletion of either the B56-1 or B56-2 subunit by itself caused a modest increase in stubarista (2.3- and 1.5-fold, respectively). Simultaneous knockdown of both B56 subunits led to a 6.7-fold increase, whereas knockdown of the catalytic subunit led to a 9-fold increase in this form of stubarista. In contrast, knockdown of either the B{alpha} or PR72 subunits of Drosophila PP2A had no effect on the levels of the stubarista spot. The relative abilities of catalytic subunit, B56-1, and combined B56-1 and -2 knockdowns to induce stubarista correlated with their ability to induce apoptosis and caspase activation (see Fig. 4C). The amount of stubarista formed when B56-1 and B56-2 were simultaneously knocked down (6.7-fold) was higher than the sum of each knockdown by itself suggesting that the loss of these subunits acts synergistically to induce apoptosis.


Figure 4
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  		FIG. 4. Blocking induction of reaper mRNA attenuates apoptosis caused by knockdown of the B56 subunits. S2 cells were exposed to control dsRNA (GFP); dsRNA corresponding to the catalytic (C Sub), B56-1, B56-2, or both B56 subunits (B56-1&2) of PP2A; or dsRNA corresponding to reaper or sickle for 3 days. A, total RNA from cells exposed to the dsRNA indicated at the top was analyzed by RT-PCR using primers specific for the catalytic, B56-1, and B56-2 subunits as indicated on the left. B, RT-PCR analysis of total RNA from cells exposed to EGFP control (GFP) or reaper or sickle dsRNA using primers specific for reaper or sickle. RT-PCR with actin-specific primers is shown as a control for RNA recovery. C, caspase activity was assayed in cells exposed to the dsRNAs indicated at the bottom (plus signs). The bar graph displays activity expressed as percentage of the control cells exposed to EGFP dsRNA (GFP) and represents the mean ± S.D. of three separate experiments. The asterisk indicates that the activity from cells treated with B56-1&2 plus reaper dsRNA was significantly different (p < 0.05) from cells treated with B56-1&2 dsRNA alone using Student’s t test.

 
In addition to proteins that appeared as new spots, there were a number of proteins whose abundance was increased following knockdown of PP2A subunits. Three of these spots, P2, P3, and P7, were identified as the proteins derived from the Drosophila CG5224, CG13941, and CG12505 genes. Increased intensity of these protein spots, as well as spot P12, was not blocked by Z-VAD-fmk nor were they increased by actinomycin D treatment (Fig. 1, B–D). CG5224 is a glutathione S-transferase whose mRNA was also increased by knockdown of the catalytic, the B56-1, or both B56 subunits. The mobility of the CG5224 protein on 2-D gels was consistent with its predicted molecular mass (25.1 kDa) and pI (5.6). Increased expression of the CG5224 protein was in the same range as the increase in its mRNA detected by microarray analysis (4.9-fold) or by qRT-PCR (3.8-fold). The up-regulation of the CG13941 and CG12505 proteins also appeared to be due to increased gene transcription as the increases in protein levels were consistent with the increases in their mRNA (Table II). The increased expression of these proteins was independent of apoptosis because neither up-regulation of the proteins or their mRNA was blocked by Z-VAD-fmk.

Proteins from dsRNA-treated S2 cells were analyzed by high resolution 2-D electrophoresis in an effort to identify altered post-translational modifications. Analysis of these gels revealed several additional proteins that were altered by knockdown of either the catalytic subunit or both B56 subunits (Fig. 2). Spot P13 was present in control cells but absent in cells depleted of the PP2A subunits (Fig. 2, A–D). This protein was identified as Drosophila CG1837, which is a putative protein-disulfide isomerase that is 45% identical to the human thioredoxin domain-containing 5 protein (Table II). In cells depleted of the PP2A subunits a new spot (P14) appeared in this region of the gel that was also identified as CG1837. The more acidic pI and reduced mobility of this form of CG1837 were consistent with the introduction of an additional negative charge by phosphorylation. A small amount of the more acidic spot was also observed in EGFP dsRNA-treated cells (Fig. 2, A and B, white arrows). The migration of these two forms of CG1837 was close to the pI and mass predicted from the amino acid sequence, 5.2 and 46.7 kDa, respectively. A second protein with shifted mobility was observed in a different region of the 2-D gel (Fig. 2, E–H). Spots P15 and P16 were both identified as CG4164, which is 59% identical to the human DNAJB11 protein. However, in this case, the protein shifted to a more basic pI with no change in apparent mass. The increased positive charge was not consistent with increased phosphorylation suggesting that some other post-translational modification had occurred. This region of the 2-D gel also showed a protein spot that disappeared (P17) and a protein spot that appeared (P18) in PP2A-depleted cells. These were identified as Cct{gamma} and Rpn11, respectively. A summary of these protein changes is included in Table II.


Figure 2
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  		FIG. 2. High resolution 2-D gel electrophoresis analysis of protein changes following knockdown of PP2A subunits. Two regions (A–D and E–H) of high resolution 2-D gels of S2 cells exposed to control dsRNA (GFP and 2x GFP), dsRNA for the catalytic subunit (C Sub), or dsRNA for both B56 subunits (B56-1&2) of PP2A for 4 days are shown. Black arrows indicate a set of protein spots (P13–P18) that were altered by the PP2A subunit knockdowns and the corresponding protein (in parentheses) identified by mass spectrometry from spots excised from the EGFP and catalytic subunit knockdown cells. In A–D, two unknown proteins (circled) and actin5C are shown as reference spots that did not change. The white arrow indicates a faint spot present in the control gels (A and B) that migrated at the same position as spot P14 in gels from cells treated with catalytic subunit or B56-1&2 dsRNA (C and D). In E–H, a reference protein spot (CG3902) that did not change following exposure to dsRNA for catalytic or B56-1&2 subunits of PP2A is indicated.

 
Knockdown of B56-1 Up-regulates Inducers of Apoptosis—
Transcripts altered by knockdown of PP2A subunits, especially the B56-1 subunit, included several genes with roles in the induction or prevention of apoptosis. In particular, increased expression of the reaper gene was detected in S2 cells depleted of the B56-1 subunit (Supplemental Table S2). reaper is a member of a group of genetically linked and structurally related genes that includes hid, grim, and sickle. The products of these genes induce apoptosis by inactivating the Drosophila inhibitor of apoptosis proteins (25–28). The up-regulation of reaper raised the possibility that increased expression of one or more of these proapoptotic genes could contribute to the apoptosis caused by loss of PP2A. Transcripts for reaper were readily detected, but mRNA for hid, grim, and sickle was below the detection limit of the microarray analysis. Transcripts for grim could not be detected using three different sets of primers, whereas mRNA for reaper, hid, and sickle was detected in control and dsRNA-treated S2 cells using qRT-PCR. The expression of hid was not affected by knockdown of any of the PP2A subunits (not shown). reaper mRNA was increased 4- and 12-fold at 2 and 3 days following addition of dsRNA for the B56-1 subunit or both B56 subunits (Fig. 3A). In contrast, knockdown of either the catalytic or B56-2 subunits had only a modest effect on reaper mRNA. The expression of sickle was also up-regulated in cells depleted of the B56-1 subunit or both B56 subunits (Fig. 3B). As observed with reaper mRNA, depletion of the catalytic or B56-2 subunits caused relatively modest increases in sickle mRNA. The increase in reaper or sickle mRNA levels was not affected when apoptosis was blocked with Z-VAD-fmk indicating that these effects were not due to the induction of apoptosis.


Figure 3
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  		FIG. 3. reaper and sickle mRNAs are up-regulated by knockdown of the B56-1 subunit. S2 cells were exposed to control EGFP dsRNA or dsRNA corresponding to the catalytic (diamonds), B56-1 (triangles), B56-2 (squares), or both B56 (circles) subunits of PP2A for the times indicated. The treatment with dsRNA for both B56 subunits was also carried out in the presence of Z-VAD-fmk (solid circles). Total RNA was extracted, and the amount of mRNA for reaper (A) and sickle (B) was assayed by qRT-PCR. The data are expressed as the -fold change in mRNA levels in cells exposed to PP2A dsRNA relative to cells exposed to control dsRNA.

 
The potential roles of reaper and sickle in mediating the apoptosis induced by knockdown of PP2A were investigated using RNAi. Knockdown of the catalytic subunit, the B56-1 subunit, the B56-2 subunit, or both the B56-1 and -2 subunits was carried out in the presence or absence of dsRNA for reaper or sickle. Depletion of the catalytic subunit or both B56 subunits caused a 2–3-fold increase in caspase activity. Depletion of B56-1 alone caused a modest (1.5-fold) increase in caspase activity (Fig. 4C), whereas depletion of B56-2 had no effect (not shown). The ability of B56-1 RNAi alone to cause modest caspase activation in this study, but not in a previous one (16), was likely due to increased knockdown efficiency. There was no effect of reaper dsRNA on the increased caspase activity caused by knockdown of the PP2A catalytic subunit. Knockdown of reaper (Fig. 4B) resulted in a statistically significant reduction (30%) in caspase activity in cells treated with B56-1 and -2 dsRNA (Fig. 4C). Depletion of reaper also caused a moderate reduction in caspase activity in cells treated with B56-1 dsRNA alone, although the difference did not reach statistical significance. Knockdown of sickle (Fig. 4B) had no effect on caspase activity induced by knockdown of any of the PP2A subunits. The combined knockdown of reaper and sickle inhibited caspase activity to the same extent observed with knockdown of reaper alone (not shown). The ability of reaper dsRNA to partially suppress caspase activation suggested that increased expression of reaper could contribute to apoptosis caused by knockdown of the B56-1 subunit. In contrast, the increase in sickle expression did not appear to play a significant role in this process.

Knockdown of PP2A Subunits Alters Expression of Genes Involved in Oxidative Stress—
The microarray and 2-D gel analyses demonstrated that knockdown of the catalytic and B56 subunits of PP2A increased the mRNA and protein levels of several genes with known or predicted roles in the response to oxidative stress. These results raise the possibility that PP2A regulates pathways utilized during oxidative stress. To test this hypothesis more directly, transcripts altered in S2 cells depleted of PP2A were compared with transcripts altered in adult flies following oxidative stress induced by exposure to paraquat (a free radical generator) or hydrogen peroxide (29). Thirty (48%) of the transcripts altered by knockdown of the PP2A catalytic subunit in S2 cells were also altered by exposure of flies to oxidative stress (bolded text in Supplemental Table S1). Twenty-five of the common transcripts were changed in the same direction. Commonly regulated transcripts included three glutathione S-transferases (GstE6, GstE7, and CG5224) and two oxidoreductases (CG2064 and CG18522). Forty-seven (42%) of the transcripts altered by combined knockdown of the B56-1 and -2 subunits in S2 cells were also altered by exposure of flies to paraquat or hydrogen peroxide (bolded text in Supplemental Table S4). The levels of 43 of these common transcripts were changed in the same direction. This group included two glutathione S-transferase genes (GstD2 and CG5224) and three genes with oxidoreductase activity (CG2064, ferredoxin, and CG18522). The substantial overlap between the changes caused by depletion of PP2A and those caused by oxidative stress supports a role for PP2A in regulating signaling pathways that affect gene transcription in response to this stress.

Up-regulation of glutathione transferases is an evolutionarily conserved response to oxidative stress (30). To further investigate a potential role of PP2A in this response, the effect of PP2A knockdown on GST gene expression was examined. Based on the Gene Ontology annotation of the Drosophila genome, there are 40 genes that are known or inferred to encode proteins with glutathione S-transferase activity. Oligonucleotide probes for 36 of these transcripts are present on the Drosophila Genome Array chips. The levels of 10 of these transcripts were up-regulated by at least 2-fold following knockdown of the catalytic subunit (Fig. 5A), but only changes in CG5224, GstE3, GstE6, and GstE7 reached the statistical criteria necessary for inclusion in Supplemental Table S1. In most cases, knockdown of the B56-1 subunit or both B56 subunits also caused up-regulation of the same GST transcripts (Supplemental Table S5). To confirm the microarray analysis, the levels of CG5224, GstD2, GstE6, and GstE7 mRNA were determined by qRT-PCR (Fig. 5B). Knockdown of the catalytic subunit, the B56-1 subunit, or both B56 subunits caused increases in these transcripts. The greatest effect was observed with GstD2, which was increased by 5- or 8-fold following knockdown of both B56 subunits and the catalytic subunit. The increases in GST mRNA levels determined by qRT-PCR was in generally good agreement with the microarray experiments. As shown in Fig. 1B, up-regulation of CG5224 (spot P2) was also detected at the protein level. The increase in these transcripts supports the hypothesis that PP2A and the B56 regulatory subunits function within pathways that regulate expression of these genes.


Figure 5
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  		FIG. 5. Knockdown of PP2A subunits increases mRNA for multiple glutathione S-transferases. A, S2 cells were exposed to control EGFP dsRNA or dsRNA corresponding to the B56-1 (open bars), B56-2 (light bars), both B56 subunits (dark bars), or the catalytic subunit (black bars) of PP2A for 3 days. Transcripts of the indicated glutathione S-transferase genes were analyzed by DNA microarrays. The -fold change in transcript levels in PP2A dsRNA-treated cells relative to cells treated with the control dsRNA is expressed as the means ± S.E. of three separate experiments. B, cells were exposed to B56-1 (open bars), B56-1&2 (light bars), or catalytic subunit dsRNA (dark bars) as described in A. The levels of the indicated glutathione S-transferase mRNAs were determined by qRT-PCR. The data are expressed as -fold change relative to cells exposed to control EGFP dsRNA and are expressed as the mean ± S.D. of three experiments.

 
Knockdown of PP2A Inhibits Activation of ERK by Oxidative Stress—
The pattern of transcriptional changes in dsRNA-treated cells suggested that PP2A modulates signaling pathways involved in the response to oxidative stress. Drosophila p38 (31, 32) and ERK (33) are activated by oxidative stress, and the Drosophila JNK pathway has been implicated in the transcriptional response to oxidative stress (34). To test the role of PP2A in regulating these pathways, activation of the Drosophila MAP kinases was examined in S2 cells depleted of PP2A subunits. Activation of Drosophila ERK (rolled), JNK (basket), and p38 (encoded by the Mpk2 and p38b genes) was monitored with phosphospecific antibodies that recognize conserved phosphopeptides encompassing the activating phosphorylation sites of these kinases.

Preliminary time course experiments indicated that acute activation of S2 cell MAP kinases by hydrogen peroxide, NaCl, or sorbitol was maximal at 10 min (not shown). ERK was strongly activated by hydrogen peroxide but weakly, if at all, by osmotic stress induced by NaCl or sorbitol (Fig. 6). The ability of hydrogen peroxide to induce ERK activation was significantly reduced following knockdown of the PP2A catalytic subunit and, to a lesser extent, by knockdown of both B56 subunits (Fig. 6, lanes 6 and 7). Knockdown of the PR72 subunit of Drosophila PP2A had no effect on ERK activation. The p38 kinase was activated to a much greater extent by NaCl or sorbitol than by hydrogen peroxide. Based on results from three separate experiments, there was no reproducible effect of knocking down the catalytic or B56 subunits of PP2A on basal or stress-induced phosphorylation of p38. JNK was also activated to a much greater extent by NaCl and sorbitol than by hydrogen peroxide. There was a modest, but reproducible, increase in the basal activity of JNK in cells depleted of the catalytic or both B56 subunits (Fig. 6, lanes 2 and 3). There was also a modest effect of PP2A knockdown on JNK activity induced by sorbitol. Immunoblotting with antibodies against the catalytic or B56-1 subunits of PP2A showed that these proteins were reduced to barely detectable levels following dsRNA treatment (Fig. 6, lower panels). As described previously (16, 17) depletion of the catalytic subunit resulted in loss of the B56-1 subunit due to its instability in the absence of PP2A holoenzymes. In summary, depletion of the catalytic or B56 subunits of PP2A in S2 cells significantly decreased activation of ERK during oxidative stress and caused an increase in basal JNK activity.


Figure 6
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  		FIG. 6. Knockdown of PP2A subunits suppresses ERK activation in response to oxidative stress. S2 cells were exposed to control dsRNA (GFP) or dsRNA corresponding to both B56 subunits (B56-1&2), the catalytic subunit (C Sub), or the PR72 subunit of Drosophila PP2A. Three days later, the cells were incubated with normal medium (None, lanes 1–4), medium containing 2 mM hydrogen peroxide (H2O2, lanes 5–8), 0.3 M NaCl (lanes 9–12), or 0.3 M sorbitol (lanes 13–16) for 10 min. The cells were lysed in SDS buffer and subjected to SDS-PAGE, and the resolved proteins were processed for immunoblotting with phosphospecific antibodies for ERK (P-ERK), p38 (P-p38), and JNK (P-JNK) and antibodies that recognize total ERK, total p38, and total JNK antibody. The same samples were also immunoblotted with antibodies against the catalytic (C Sub) and B56-1 subunits of PP2A to verify the knockdowns. The arrows on the right of the lower two panels indicate the mobilities of Drosophila p38 and the PP2A catalytic and B56-1 subunits. The blot probed with the anti-catalytic subunit antibody was the same blot probed previously with anti-phospho-p38. The residual phospho-p38 signal can be observed just above the catalytic subunit.

 

   DISCUSSION
TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The functional genomics approaches used in this study revealed multiple mechanisms by which the B56 isoforms of PP2A promote cell survival, including a novel role in oxidative stress pathways. There was limited overlap (32%) between genes whose transcript levels were altered by loss of the catalytic subunit and those altered by loss of the B56 subunits. This observation is consistent with the expectation that a subset of the transcriptional effects of the PP2A catalytic subunit would be mediated by the B56 regulatory subunits. The most common effect of knocking down PP2A subunits were changes in expression of functionally related transcripts involved in Drosophila defense responses. The substantial overlap (42–48%) between transcripts altered by knockdown of PP2A subunits with transcripts altered in response to oxidative stress (35) revealed that PP2A is likely to modulate signaling pathways that mediate responses to this stress. In particular, transcripts for a number of genes encoding glutathione transferases were up-regulated by knockdown of PP2A subunits.

Up-regulation of glutathione transferases is an evolutionarily conserved response to oxidative stress (30). Glutathione transferases catalyze the conjugation of reduced glutathione to a wide variety of electrophilic compounds, including compounds generated by reactive oxygen species. Members of the Drosophila GstD and GstE classes, as well as GstS1, participate in the defense against oxidative stress by metabolizing lipid peroxidation products (36, 37). In this regard, they are functionally analogous to the mammalian Alpha class of glutathione transferases. Up-regulation of GstD1 in response to oxidative stress is regulated by signaling through the JNK MAP kinase pathway (34). The transcription factors involved in increased expression of glutathione transferases in Drosophila are not known. However, the promoters of the GstE2 and GstE3 genes of the mosquito Anopheles gambiae, which are also up-regulated by oxidative stress, contain putative binding sites for transcription factors that mediate responses to oxidative stress in other species (38). The effects of knocking down the catalytic subunit or the B56 subunits suggest that the B56-containing isoform of PP2A acts as a negative regulator of Drosophila transcriptional pathways activated by oxidative stress.

Proteomics experiments identified Drosophila proteins that may be direct substrates of PP2A as well as proteins that are proteolytically cleaved following the induction of apoptosis. RhoGDI, lamin, and eIF5 were identified as putative caspase substrates in Drosophila S2 cells. Both RhoGDI (39, 40) and lamin (41, 42) are cleaved by caspases in mammalian cells. The 2-D gel analysis also identified proteins, including the glutathione transferase CG5224, whose increase in protein levels correlated with an increase in mRNA levels. Neither the induction of the protein or mRNA for these genes is dependent on apoptosis indicating they are transcriptional targets of pathways modulated by the B56 isoforms of PP2A. A third set of changes detected by 2-D gels were a set of proteins that appeared to undergo altered covalent modification. The shifted mobility of the putative protein-disulfide isomerase encoded by the CG1837 gene is consistent with phosphorylation, suggesting that the CG1837 protein could be a substrate of B56-containing forms of PP2A. The 2-D gel shift in the CG4164 protein, which is 59% identical to the human DnaJ protein DNAJB11, is consistent with a covalent modification resulting in the loss of negative charge following knockdown of PP2A. Although the nature of such a covalent modification is unknown, one possibility is carboxymethylation. Several mammalian DnaJ proteins are prenylated and carboxymethylated resulting in the loss of the hydroxyl group at the C terminus (43). A prominent apoptosis-dependent change in protein levels was an increase in the stubarista protein. The increase in stubarista correlated with apoptosis because it was blocked by Z-VAD-fmk. Stubarista is a ribosome-associated protein proposed to play a positive role in translation and cell size as well as a negative role in cell proliferation (44, 45). The 2-D gel data did not provide sufficient information to determine whether the dramatic increase in stubarista protein was due to increased abundance or post-translational modification.

The distinct effects of RNAi-mediated knockdown of the B56-1 and B56-2 subunits indicate that the two proteins are functionally distinct. Depletion of the B56-1 subunit, but not the B56-2 subunit, resulted in dramatic up-regulation of reaper and sickle mRNA. Genetic evidence also supports distinct roles for the two B56 subunits. The Drosophila B56-1 (PP2A-B') protein interacts with a homeodomain transcription factor, and ablation of B56-1 by RNAi results in embryos without salivary glands (46). The B56-2 subunit (widerborst) has been shown to play a role in polarization of planar cells in the wing epithelium during development (9). The B56-2 subunit, but not the B56-1 subunit, stabilizes the Period protein during the Drosophila circadian rhythm (10).

RNAi-mediated knockdown of the B56-1 subunit of Drosophila PP2A caused increased expression of the mRNA for reaper and sickle. The products of these genes induce apoptosis by inactivating the Drosophila inhibitor of apoptosis proteins (25–28), and both genes are transcriptional targets of the Drosophila p53 protein (28, 47–50). The lack of an effect of catalytic subunit knockdown on reaper and sickle is unexpected based on the assumption that the effects of losing B56-1 would reflect a subset of effects caused by loss of the catalytic subunit. Loss of the B56-1 subunit could activate the p53 pathway, whereas loss of the catalytic subunit may cause general inhibition of the transcriptional machinery and blunt the activating effect brought about by loss of B56-1. Although increased expression of reaper was not detected, the ability of reaper knockdown to attenuate the apoptosis caused by knockdown of the B56 subunits is consistent with a previous study (17). Co-knockdown of sickle mRNA had no effect, indicating that sickle does not play a significant role in the apoptosis caused by loss of the B56 subunits. This lack of effect is consistent with previous data showing that sickle acts as an enhancer rather than a determinant of apoptosis (49, 51). Co-knockdown of reaper and sickle had no effect on the extent of caspase activation caused by knockdown of the catalytic subunit, consistent with the lack of effect on expression of these transcripts. This result is distinct from the previous study (17) where partial knockdown of reaper attenuated apoptosis mediated by knockdown of the catalytic subunit. This difference may be due to the extent of catalytic subunit knockdown in each study. Despite this difference, the enhanced expression of reaper and sickle is consistent with the notion that activation of the p53-dependent transcriptional pathway is at least one mechanism involved in the apoptosis caused by knockdown of the B56 subunits of Drosophila PP2A (17).

Knockdown of either the catalytic or B56 subunits of PP2A led to a substantial decrease in the activation of ERK by oxidative stress. Although the mechanisms involved in ERK activation in response to this stress have not been defined, our data indicate that PP2A plays a positive role in this process. The activation of ERK in response to oxidative stress is generally associated with enhanced cell survival (52, 53). A decrease in an ERK-mediated prosurvival activity, due to loss of PP2A, may contribute to the apoptosis caused by knockdown of the catalytic or B56 subunits. A positive function of the B56 forms of PP2A in ERK activation by oxidative stress contrasts with the negative role of the B55/B{alpha} form of PP2A in activation of ERK by insulin (16). The ability of PP2A to play both positive and negative roles in activation of the ERK pathway has also been observed during the development of Drosophila photoreceptors (54). The ability of PP2A to play multiple roles in pathways involved in regulation of Drosophila ERK is likely to be mediated by the unique substrate specificities imparted by the different regulatory subunits.

Increased activity of the Drosophila JNK pathway is associated with transcriptional up-regulation of genes involved in oxidative stress responses (55) and with the induction of apoptosis (56–60). Elevated basal JNK activity may play a role in the transcriptional effects and apoptosis caused by depletion of the PP2A subunits. It is notable that the reaper gene product has been shown to activate JNK via up-regulation of ASK1 protein kinase activity (56). Thus, the increase in JNK activity observed in cells depleted of the B56 subunits may be a function of increased reaper expression. These data suggest that an important action of the B56 forms of PP2A is to regulate stress-sensitive MAP kinase pathways to ensure maintenance of a proper balance between prosurvival and proapoptotic activities.


  FOOTNOTES
 
Received, July 25, 2006, and in revised form, November 14, 2006.

Published, MCP Papers in Press, November 22, 2006, DOI 10.1074/mcp.M600272-MCP200

1 The abbreviations used are: PP2A, protein phosphatase 2A; RNAi, RNA interference; dsRNA, double-stranded RNA; qRT-PCR, real time quantitative RT-PCR; EGFP, enhanced green fluorescent protein; 2-D, two-dimensional; 2-D electrophoresis, two-dimensional polyacrylamide gel electrophoresis; Z-VAD-fmk, benzyloxycarbonyl-Val-Ala-Asp-(OMe)-fluoromethyl ketone; ERK, extracellular signal-regulated kinase; JNK, c-jun N-terminal kinase; MAP, mitogen-activated protein; SAPK, stress-activated protein kinase; RhoGDI, Rho GDP dissociation inhibitor. Back

* This work was supported by National Institutes of Health Grant GM49505. 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

S The on-line version of this article (available at http://www.mcponline.org) contains supplemental material. Back

{ddagger} Present address: Trimeris, Inc., 3500 Paramount Parkway, Morrisville, NC 27560. Back

§ Present address: Advanced Mass Spectrometry Laboratory, University of North Texas Health Science Center, 3500 Camp Bowie Blvd., RES 2-146W, Fort Worth, TX 76107-2699. Back

To whom correspondence should be addressed: Dept. of Pharmacology, University of Texas Southwestern Medical Center, 5323 Harry Hines Blvd., Dallas, TX 75390-9041. Tel.: 214-645-6152; Fax: 214-645-6151; E-mail: marc.mumby{at}utsouthwestern.edu


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