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. 2011 Jun;188(2):359–367. doi: 10.1534/genetics.110.124222

A Link Between Impaired Purine Nucleotide Synthesis and Apoptosis in Drosophila melanogaster

Catherine Holland 1,1, David B Lipsett 1,2, Denise V Clark 1,3
Editor: M Wolfner
PMCID: PMC3122316  PMID: 21441212

Abstract

The biosynthetic pathways and multiple functions of purine nucleotides are well known. However, the pathways that respond to alterations in purine nucleotide synthesis in vivo in an animal model organism have not been identified. We examined the effects of inhibiting purine de novo synthesis in vivo and in cultured cells of Drosophila melanogaster. The purine de novo synthesis gene ade2 encodes phosphoribosylformylglycinamidine synthase (EC 6.3.5.3). An ade2 deletion, generated by P-element transposon excision, causes lethality in early pupal development, with darkening, or necrosis, of leg and wing imaginal disc tissue upon disc eversion. Together with analysis of a previously isolated weaker allele, ade24, and an allele of the Prat gene, which encodes an enzyme for the first step in the pathway, we determined that the lethal arrest and imaginal disc phenotypes involve apoptosis. A transgene expressing the baculovirus caspase inhibitor p35, which suppresses apoptosis caused by other stresses such as DNA damage, suppresses both the imaginal disc tissue darkening and the pupal lethality of all three purine de novo synthesis mutants. Furthermore, we showed the presence of apoptosis at the cellular level in both ade2 and Prat mutants by detecting TUNEL-positive nuclei in wing imaginal discs. Purine de novo synthesis inhibition was also examined in tissue culture by ade2 RNA interference followed by analysis of genome-wide changes in transcript levels. Among the upregulated genes was HtrA2, which encodes an apoptosis effector and is thus a candidate for initiating apoptosis in response to purine depletion.


THE pathway for de novo synthesis of purine nucleotides is almost universal, with exceptions among the intracellular parasites that obtain purines from their host (Chaudhary et al. 2004). Thus, although all organisms have pathways for uptake and salvage of purine nucleotides, the de novo synthesis pathway is essential. In multicellular organisms, the enzymes for the ten step pathway for synthesis of inosine monophosphate (IMP) are encoded by six genes, where three genes encode polypeptides with multifunctional properties (Figure 1) (Henikoff 1987). IMP is the common precursor for further synthesis to adenosine monophosphate (AMP) and guanosine monophosphate (GMP).

Figure 1.—

Figure 1.—

Purine de novo synthesis pathway genes in D. melanogaster. Pathway intermediates: PRPP, 5-phosphoribosyl-1-pyrophosphate; PRA, 5-phosphoribosyl-1-amine; GAR, 5-phosphoribosylglycinamide; FGAR, 5-phosphoribosyl N-formylglycinamide; FGAM, 5-phosphoribosyl N-formylglycinamidine; AIR, 5-phosphoribsylaminoimidazole; CAIR, 5-phosphoribosyl-5-aminoimidazole carboxylate; SAICAR, 5-phosphoribosyl 4-(N-succinocarboxamide)-5-aminoimidazole; SAMP, adenylosuccinate; XMP, xanthosine monophosphate. Pathway enzymes: 1—PRAT, phosphoribosylamidotransferase; 2—GARS, GAR synthetase; 3—GART, GAR transformylase; 4—FGARAT, FGAR amidotransferase; 5—AIRS, AIR synthetase; 6—AIRC, AIR carboxylase; 7—SAICARS, SAICAR synthetase; 8 and 14—ADSL, adenylosuccinate lyase; 9 and 10—ATIC, AICAR transformylase-inosinate cyclohydrolase; 11—IMPDH, IMP dehydrogenase; 12—GMPS, GMP synthetase; 13—SAMP synthetase.

In mammalian tissue culture cell lines, de novo synthesis of purines is more active in proliferating cells than in differentiating cells (Natsumeda et al. 1984; Barankiewicz and Cohen 1987; Ahmed and Weidemann 1994). As a result, this pathway has been a target for the development of anticancer drugs, including purine analogs such as 6-mercaptopurine and folate antimetabolites such as DDATHF and methotrexate (Christopherson et al. 2002). The cellular effects of these drugs likely vary due to their degree of specificity for a particular enzyme. For example, methotrexate targets three enzymes in the purine de novo synthesis pathway (Sant et al. 1992), whereas DDATHF targets a single enzyme (Christopherson et al. 2002). In addition, the effects of inhibitors vary with cell type, leading to death or reversible arrest, depending on the presence or absence of a G1 checkpoint, respectively (Zhang et al. 1998). The inhibition of purine de novo synthesis by drug inhibitors leads to growth arrest, or quiescence, possibly through detection of ribonucleotide levels by p53 rather than by DNA damage (Linke et al. 1996). However, growth arrest can be independent of p53 status (Bronder and Moran 2002). Thus, the mechanism of growth arrest in mammalian cells caused by purine de novo synthesis inhibitors, and its dependence on p53, has not been clearly established.

Two genetic disorders in the purine de novo synthesis pathway have been associated with the ADSL (Sivendran et al. 2004) and ATIC (Marie et al. 2004) genes (Figure 1). In both cases, individuals are short-lived and suffer from profound effects on neurological development. Mutations in the other genes in the purine de novo synthesis pathway have not been recovered in humans.

Since the cellular and developmental effects of specifically inhibiting purine de novo synthesis are not well characterized and would be informative for understanding the mechanisms of drug inhibitors in vivo, we are exploring these effects in Drosophila melanogaster. Purine de novo synthesis gene mutations cause arrest in development from pupal stages to the adult stage with a variety of defects in wing, leg, eye, and bristle development (Tiong et al. 1989; Tiong and Nash 1990; Clark 1994; Ji and Clark 2006) as well as reduced fertility and life span (Malmanche and Clark 2004). Here, we focus on identifying factors that respond to depletion of purine de novo synthesis during development and in a cell line. We examine mutations in Prat and ade2, which encode enzymes for the first and fourth steps in the pathway, respectively (Figure 1). Both the pupal lethal arrest of mutants and the necrosis in imaginal tissue can be suppressed by an apoptotic caspase inhibitor. Consistent with these results, an in situ cell death assay of wing imaginal discs from newly formed pupae shows apoptotic nuclei. To identify genes responding to purine de novo synthesis depletion, we knocked down ade2 expression in a Drosophila Schneider cell line and performed a gene expression microarray analysis. We identified several genes with up- or downregulated transcripts, including HtrA2, a serine protease associated with apoptosis in Drosophila (Challa et al. 2007) and humans (Vande Walle et al. 2008).

MATERIALS AND METHODS

Drosophila culture:

D. melanogaster strains were obtained from the Bloomington Drosophila Stock Center, with the exception of ade24/SM1 (from David Nash), flies carrying Df(3R)dsx43 (from Bruce Baker), and Prat12A19/TM6B (from this lab). Flies were cultured on standard cornmeal–molasses–agar medium at 25° and 60% relative humidity. Schneider cell strain #6 (S2) was obtained from the Drosophila Genomics Resource Center and was cultured in Invitrogen Schneider cell medium supplemented with 10% fetal bovine serum at 25°. Cell viability was measured by exclusion of trypan blue, and metabolic activity was measured using the tetrazolium salt WST-1 (Roche Applied Science).

Fly crosses:

P-element insertion P{EPgy2}EY05576 (Bellen et al. 2004) was used to generate deletions in ade2 (Voelker et al. 1984). Virgin y w67c23; P{EPgy2}EY05576 females were crossed to w1118; al b c sp/CyO; ry506 Dr P{Δ2-3}99B/TM6B, Tb males. Dysgenic F1 w1118; P{EPgy2}EY05576/al b c sp; ry506 Dr P{Δ2-3}99B/+ males were crossed to w; al b c sp virgins to identify both P-element excision and P-element-induced male recombination events (Chen et al. 1998) by exchange of the flanking markers al and b. White-eyed F2 males of genotype w; ade2*/al b c sp, where ade2* represents a possible excision allele, were crossed to w; Df(2L)TE35BC-31, b pr pk cn sp/CyO, b* to balance ade2*, since the b marker on CyO, b* allowed distinction between the two second chromosomes. The resulting w; ade2*/CyO, b* strains were then screened for absence of ade2* homozygous adults. Lethal mutations were balanced over the green fluorescent protein (GFP)-marked CyO, P{GAL4-Kr.C}DC3, P{UAS-GFP.S65T}DC (CyO, Kr-GFP) (Casso et al. 2000) to allow identification of mutant homozygotes. A viable excision allele, ade21–5, was also kept. Mutations were mapped to the ade2 region by complementation testcrosses with w; Df(2L)ED343/CyO, Kr-GFP (Ryder et al. 2004). Deletion breakpoints were mapped by PCR and sequencing using primers flanking the P-element insertion site (Figure 2 and Table S1). A UAS-ade2 cDNA transgene was constructed by inserting the 4.2-kb XbaI partial digest fragment of cDNA GM01721 (Rubin et al. 2000) into the XbaI site of pUASP (Rorth 1998). Transformants in strain w1118 were generated by Genetic Services. cDNA expression was driven by P{tubP-GAL4}LL7 (Lee and Luo 1999).

Figure 2.—

Figure 2.—

P-element-induced deletion allele ade21–6. Map of region containing ade2 based on Flybase FB2010_06 (June 25, 2010). Thick bars represent exons, with shading indicating untranslated regions and solid indicating coding sequence. Thin lines represent introns. Downward triangle indicates site of P-element insertion P{EPgy2}EY05576 (Bellen et al. 2004). Horizontal line under the map shows the extent of the ade21–6 deletion. Arrows show the position of primers used to identify and sequence the allele (Figure S1).

For examining p35 suppression, we used chromosome 2- and 3-linked insertions of P{UAS-p35.H} (Zhou et al. 1997) or UAS-p35. For ade21–6 suppression, non-Tubby progeny pupae were scored from three crosses: (1) ade21–6 ; +/CyO:TM6B siblings, where CyO:TM6B is T(2;3)TSTL, CyO: TM6B, Tb; (2) ade21–6; +/CyO:TM6B crossed to ade21–6; UAS-p35/CyO:TM6B; and (3) ade21–6; UAS-p35/CyO:TM6B siblings. For ade24 (Tiong et al. 1989) suppression (the ade24 chromosome carried P{neoFRT}40A, which is not relevant in this study), non-Tubby pupae were scored from two crosses: (1) ade24; +/CyO:TM6B crossed with Df(2L)ED343; UAS-p35/CyO:TM6B and (2) ade24; +/CyO:TM6B crossed with Df(2L)ED343; +/CyO:TM6B. For Prat12A19 suppression, non-Tubby pupae were scored from two crosses: (1) +; Prat12A19e11/TM6B crossed with +; Df(3R)dsx43 e11/TM6B and (2) UAS-p35; Prat12A19 e11/CyO:TM6B crossed with +; Df(3R)dsx43 e11/TM6B, where Df(3R)dsx43 deletes Prat and flanking genes (Clark 1994). Non-Tubby stage P1 white prepupae (Bainbridge and Bownes 1981) were also used for detection of p35 transcripts by RT-PCR. For examining suppression of ade21–6 by p535A-1–4 (Rong et al. 2002), progeny of ade21–6; p535A-1–4/CyO:TM6B siblings and ade21–6; +/CyO:TM6B siblings were compared. Crosses were set up in parallel with 5 males and either 10 or 20 virgin females in clear polycarbonate vials to allow scoring of undisturbed pupae on sides of vials. Parents were transferred every 24 hr. Time after egg deposition (AED) was measured in days since the transfer. For tests of suppression of necrosis and pupal lethality of the three purine mutants, each group of data was examined by chi-square analysis, and a P-value was determined (Preacher 2001). For TUNEL assays and real-time quantitative RT-PCR assays, ade21–5and ade21–6 homozygous stage P1 white prepupae and third instar wandering larvae were identified in stocks carrying the CyO, Kr-GFP balancer chromosome, while Prat12A19 e11/Df(3R)dsxR43 e11 individuals were generated as described above.

TUNEL assays:

TUNEL assays followed Kim et al. (2007) with a 100 mm sodium citrate permeabilization step added (Vicente-Crespo et al. 2008). Following TUNEL dUTP-FITC labeling (Roche Applied Science), Draq5 nuclear staining (Molecular Probes) was done at 10 μm in PBS with 0.1% Triton-X100 for 30 min. Positive controls for TUNEL labeling were generated by a 1- to 2-hr, 37° heat shock 4–24 hr before dissection (Perez-Garijo et al. 2004). The ade2 experiment was performed on ade21–6 and ade21–5 discs (with and without heat shock) in parallel, and the Prat experiment was performed with Prat12A19e11/Df(3R)dsx43 e11 and e11 discs (with and without heat shock) in parallel. Discs were mounted in 70% glycerol in 1× PBS and imaged using a Leica TCS-SP2 confocal microscope. Z-series of images were collected using the average intensity setting, and channels were merged using ImageJ (Abramoff et al. 2004).

RNA interference in tissue culture cells:

To control for possible off-target effects using double-stranded RNA (dsRNA) (Ma et al. 2006), two non-overlapping templates were prepared using the ade2 cDNA LD23935 (Rubin et al. 2000). The ade2 dsRNA-1 template was a 741-bp SacI–HindIII fragment cloned into both pSPT18 and pSPT19 vectors (Roche Applied Science) to allow synthesis of complementary single-stranded RNA (ssRNA). The ade2 dsRNA-2 template was made by PCR using the T7 primer and 5′-GGGGATCCAAATCGTAGTCGTTGAAGG-3′ to produce a 745-bp product and then digested with EcoRI and BamHI and cloned into pSPT18 and pSPT19. A nonspecific dsRNA was made from a Chlamydomonas lhcb cDNA 313-bp BglII–PstI fragment (GenBank accession AY171229) kindly provided by D. Durnford. Complementary ssRNAs were made using linearized templates and a T7 polymerase transcription kit (Ambion), followed by annealing. Treatment of Drosophila S2 cells with dsRNA was done in triplicate for 4 days at 25° with 30 μg dsRNA/ml of cells at an initial density of 1 × 106 (Maiato et al. 2003).

Microarray hybridization:

RNA was extracted from S2 cells using Trizol (Invitrogen). Each replicate represents cells treated and grown in a separate dish. Three replicates of each of the ade2 dsRNA-1 and lhcb dsRNA treatments and two replicates of the ade2 dsRNA-2 treatment underwent RNA labeling, hybridization using the Agilent 1-color protocol, microarray scanning, and data normalization at the Laboratory for Advanced Genome Analysis at the Vancouver Prostate Centre, Vancouver, Canada, as follows. RNA was linearly amplified and labeled with Cy3, and labeled cRNA quantity and specific activity were assessed with the NanoDrop ND-1000 (Thermo Scientific, Wilmington, DE). Labeled cRNA (1.65 μg) was fragmented for 30 min and applied to Agilent Drosophila microarrays in the 4 × 44,000 format (product G2519F), hybridized for 17 hr at 65° in the Agilent hybridization oven, and washed with Agilent wash buffers. Scanning was done with the Agilent DNA Microarray Scanner at a 5-μm resolution. Agilent’s Feature Extraction 9.1 software was used for background subtraction and quality control measurements. The intensities per spot had their local background subtracted. A step of spatial detrending was done using the negative control spots across the whole array and subtracting a surface fit from the data. The data were loaded into GeneSpring 7.3, and raw values <5 were set to 5. The data were normalized to the median, so that intensities were divided by the median of all intensities. Normalized and raw data were deposited in the GEO database at NCBI (accession GSE24123). A t-test P-value for significance of each value above background was calculated. For comparison among microarrays, mean values for each treatment were used in a t-test comparing expression levels between treatments, and a fold-change value for the corresponding spot was calculated. Normalized values were flagged as A (absent) or P (present or marginal), and these flags were used for filtering the data into different classes for production of gene lists.

Western blots:

A polyclonal anti-Ade2 antiserum (Alpha Diagnostic International) was raised in rats against a recombinant Ade2 polypeptide. A 776-bp BamHI–XhoI fragment from the 3′-end of the LD23935 cDNA (Rubin et al. 2000) was inserted into pQE30 (Qiagen), producing a 219-amino-acid 6X-His protein. The antiserum was affinity-purified against 6XHis-Ade2 as previously described (Gu et al. 1994; Clark and Macafee 2000). Western blot detection was done using the Aurora blocking reagent and protocol (MP Biomedicals) with alkaline phosphatase (AP)-conjugated donkey anti-rat IgG (Jackson Immunoresearch) secondary antibody. α-Tubulin was detected using mouse antibody DM1A (Sigma) and AP-conjugated goat anti-mouse IgG (Sigma). AP was detected using CDP-Star (New England Biolabs). Images were captured on X-ray film and digitally scanned.

Quantitative RT-PCR:

ade2 mRNA levels were checked in the RNA samples used for microarrays by quantitative reverse transcription PCR (qRT-PCR) using a Superscript III Platinum SYBR green two-step kit with ROX and the Applied Biosystems 7900HT instrument. ade2 RNA levels were normalized to ribosomal protein 49 (rp49) levels.

For qRT-PCR analysis of genes showing differential expression in the microarray, primers were designed using either AUTOPRIME version 1.0 (Wrobel et al. 2004) or Primer3 (Rozen and Skaletsky 2000) if the gene lacked introns (Table S1). RNA was treated with RQ1 DNAse (Promega). cDNA synthesis and qRT-PCR were done using a Superscript III Platinum Two-Step qRT-PCR kit with SYBR Green (Invitrogen) and a Rotor-Gene 6000 (Corbett Life Science). Amplification conditions were 95° for 2 min and then 40 cycles of 95° for 15 sec, 60° for 30 sec, and 72° for 30 sec, followed by ramping from 72° to 95° for melt curve analysis. Each reaction was done in triplicate along with a negative control produced without reverse transcriptase. Rotor-Gene data were exported to LinReg Version 12.5 (Ruijter et al. 2009) to determine primer efficiency and CT values and then to REST Version 2.0.13 (Pfaffl et al. 2002) to calculate fold changes in RNA levels between ade2 RNA interference (RNAi) and the lhcb control. The RNA for the gene of interest was normalized to rp49 RNA levels in each sample.

RESULTS

Generation of an ade2 deletion by P-element excision:

The P-element insertion P{EPgy2}EY05576 (Bellen et al. 2004) maps 19 bp upstream from the 5′-end of the ade2-RA and ade2-RB transcripts (Tweedie et al. 2009) (Figure 2). Of 30 independent excisions, ade21–6 and ade23–20 showed pupal lethality. Analysis of ade21–6 genomic DNA showed an 856-bp deletion starting from the site of the P-element insertion and extending into the ade2-coding region (Figure S1). A total of 33 bp of P-element inverted repeat sequence remain at the insertion site, and the sequence upstream of the insertion site is intact. This deletion removes the predicted transcription start sites and 393 bp of coding sequence. A UAS-ade2 transgene, driven by ubiquitously expressed tub-GAL4, is sufficient to rescue the pupal lethality of ade21–6, and the rescued adults appear wild-type (Figure S2). Sequence analysis of the viable ade21–5 allele showed that it had a wild-type sequence (Tweedie et al. 2009) and thus was the result of a precise P-element excision (data not shown).

Purine syndrome phenotype is suppressed by expression of baculovirus protein p35:

The purine syndrome phenotype shows pupal lethality or adult escapers with wing and leg phenotypes indicating defects in cell proliferation (Tiong et al. 1989). The most severe ade2 alleles show arrest predominantly at pupal stage P5 with characteristic darkened tissue, or “necrotic” regions, in the everted wing and leg imaginal discs (Ji and Clark 2006). This phenotype is also found for Prat mutants (Clark 1994; Ji and Clark 2006). To determine whether the developmental arrest and necrotic tissues are associated with apoptosis, we tested the ability of the baculovirus protein p35 (Hay et al. 1994) to suppress these phenotypes.

Flies with an ade21–6, ade24, or Prat12A19 mutation and a p35-expressing transgene were compared to the same mutants not carrying the p35 transgene. UAS-p35 is normally used with a GAL4 driver (Phelps and Brand 1998); however, during the process of strain construction, we noted that the UAS-p35 transgene was sufficient to suppress the purine syndrome phenotype alone in the absence of a GAL4 driver (Figure 3). Both the P5 arrest and the necrosis were suppressed in both ade2 mutants. In addition, the degree of suppression with one copy of the UAS-p35 transgene was less than that for two copies. The necrosis and lethal arrest found for the ade24 and Prat12A19 hemizygous mutants were also suppressed by one copy of UAS-p35 inserted on chromosomes 3 and 2, respectively. Therefore, the suppression of the purine syndrome phenotype by UAS-p35 is not specific to a particular ade2 allele; nor is it specific to a particular UAS-p35 transgene insert or purine de novo synthesis gene.

Figure 3.—

Figure 3.—

Suppression of ade2 and Prat mutant phenotypes by baculovirus protein p35. (A) Presence of necrosis at pupal stage P5 (Bainbridge and Bownes 1981) and older, scored at 7 days AED for ade21–6 and 9 days AED for ade24 and Prat12A19. Numbers at the top indicate total pupae scored for each genotype. (B) Representative P5 individuals for the three ade21–6 genotypes scored in A. (C) Proportions surviving to pupal stages P5, P15, and adult. Numbers at top indicate total pupae/adults scored. Chi-square analyses for groups of data for each mutant, for both A and C, gave P-values ≤0.0001.

Since a GAL4 driver was not used to induce UAS-p35 expression, we wanted to verify that the UAS-p35 transgene was expressed in the absence of a GAL4 driver. RT-PCR assays for p35 transcripts were done for ade21–6;UAS-p35 pupae and for ade21–6; UAS-p35 and UAS-p35;Prat12A19/Df(3R)dsx43 stage P1 prepupal wing discs. Results showed that the p35 transgene is indeed transcribed in the absence of a GAL4 driver for both genotypes (Figure S3). Therefore, the developmental arrest and the necrotic phenotype found for the purine syndrome phenotype in ade2 and Prat mutants are dependent on the caspase activity suppressed by p35.

Since the transcription factor p53 can be an effector in the apoptotic response to DNA damage (Lu and Abrams 2006), we tested the ability of a null mutation in p53, p535A-1–4 (Rong et al. 2002), to suppress the necrosis in stage P5-arrested pupae. We found no significant difference in the degree of necrosis between ade21–6; p535A-1–4 and ade21–6; + pupae (Figure S4A). The reaper, hid, and grim genes are apoptosis effectors that antagonize the inhibitor of apoptosis protein DIAP1, allowing activation of caspases (Xu et al. 2009), and these genes map to a region uncovered by the deficiency Df(3L)H99 (White et al. 1994). H99 heterozygotes show a reduction in apoptosis in wing imaginal discs of irradiated larvae (Brodsky et al. 2004). Thus, we asked whether the necrosis in ade21–6 stage P5-arrested pupae was suppressed in a H99/+ genetic background. We found no significant difference in the degree of necrosis in this genetic background (Figure S4B).

Apoptosis is detected in ade2 and Prat mutant wing imaginal discs of white prepupae:

To determine if the ade2 and Prat mutant phenotypes are associated with apoptosis at the cellular level, we examined wing imaginal discs for DNA strand breaks using the TUNEL assay. We found that wing discs from white prepupae for both ade21–6 homozygotes (n = 6) and Prat12A19 e11/Df(3R)dsxR43 e11 heterozygotes (n = 6) showed TUNEL-positive nuclei, particularly over the wing pouch region (Figure 4). We also noted that the TUNEL-positive nuclei were smaller and more condensed than the surrounding nuclei, as is found with apoptotic nuclei (Denton et al. 2008).

Figure 4.—

Figure 4.—

Apoptosis as measured by TUNEL assays in wing imaginal discs of ade2 and Prat mutants. Genotypes of wing discs in each row are indicated at left. Sets of three images are, from left to right: Draq5 nuclear stain, TUNEL FITC stain, and an overlay of the two images. For the ade2 and Prat mutant discs, higher resolution images of each red-box region, highlighting a group of TUNEL-positive nuclei, are shown. Scale bar, 50 μm.

Gene expression response to knockdown of ade2 expression in S2 tissue culture cells:

To examine the gene expression response to a reduction in purine de novo nucleotide synthesis, we generated two non-overlapping ade2 dsRNAs (dsRNA-1 and dsRNA-2) to knock down expression of ade2 in Drosophila S2 tissue culture cells and observed the genome-wide changes in transcript abundance using microarray hybridization. We verified that our ade2 dsRNA knockdown by both non-overlapping dsRNAs was affecting expression of ade2 protein by Western blot analysis (Figure S5A). Growth and metabolic activity of ade2 dsRNA-treated cells were reduced, as measured by WST-1 assay (Figure S5B). Changes in transcript abundance were measured relative to a Chlamydomonas lhcb dsRNA. A total of 198 and 210 microarray probes showed significantly different hybridization signals for ade2 dsRNA-1 vs. lhcb and ade2 dsRNA-2 vs. lhcb treatments, respectively, and 41 probes were common to both comparisons. A candidate list was compiled for the 20 genes showing absolute values of log2 fold changes ≥0.9 (Table S2).

We examined expression of the 20 genes for changes in transcript levels by qRT-PCR. One biological replicate was examined for each of the ade2 dsRNA-1, ade2 dsRNA-2, and control lhcb dsRNA treatments. Transcripts from 9 genes (CG6660, CG3649, CG5866, DNAseII, CG41436, CG5773, Pph13, CG14432, and Ank2) could not be detected by qRT-PCR either at all or sufficiently in any sample to reliably measure fold changes. The data for the 11 other genes are shown in Figure 5A. Interpro domain (Hunter et al. 2009) and microarray data are summarized in Table 1. As expected, ade2 showed significantly reduced transcript levels. Five genes (RPII215, CG34439, HtrA2, FKBP59, and CG11436) showed reproducible fold changes in response to ade2 RNAi in comparison to the microarray data. The other 5 genes showed no significant change in expression; however, for the genes CG31002 and CG13283, results were unreliable, primarily due to low transcript abundance and CT values >35.

Figure 5.—

Figure 5.—

Genes that respond to ade2 RNAi in tissue culture cells. (A) Fold changes in transcript levels measured by qRT-PCR in cells treated with ade2 dsRNA-1 and ade2 dsRNA-2 relative to cells treated with control lhcb dsRNA. (B) Fold changes in transcript levels measured by qRT-PCR in whole wandering third instar larvae and stage P1 prepupal wing discs. Error bars show standard errors of fold changes, and asterisks show results where the probability of a type 1 error P(H1) is <0.05, as calculated by the REST program (Pfaffl et al. 2002).

TABLE 1.

Genes showing changes in transcript levels in response to ade2 RNA interference in S2 cells that were also assayed by real-time qRT-PCR

Gene Interpro domaina Log2 fold changeb P-value
ade2 IPR010073: Phosphoribosylformylglycinamidine synthase −2.56 0.000
RpII215 IPR000722: RNA polymerase, α-subunit −2.01 0.011
CG17612 IPR007087: Zinc finger, C2H2-type −1.09 0.025
CG31002 IPR002213: UDP-glucuronosyl/UDP-glucosyltransferase 0.88 0.044
CG34439 Unknown 0.91 0.004
0.93 0.007
CG13283 IPR000718: Peptidase M13, neprilysin 0.94 0.004
HtrA2 IPR009003: Peptidase cysteine/serine, trypsin-like 1.09 0.001
MESK2 IPR004142: Ndr family (N-myc-downstream-repressed) 1.10 0.009
CG4615 IPR004254: Hly-III-related (hemolysin III-related) 1.13 0.004
FKBP59 IPR001179: Peptidyl-prolyl cis-trans isomerase, FKBP-type 1.35 0.001
CG11436 IPR013026: Tetratricopeptide repeat-containing 2.05 0.000
2.44 0.001
a

Interpro domain (Hunter et al. 2009) was reported in FlyBase (Tweedie et al. 2009). In cases where several domains are reported, one functionally descriptive entry was selected.

b

Log2 fold change absolute values ≥0.9 as detected by microarray analysis. Transcripts showing two values correspond to the values for two different oligonucleotides on the microarray. See Table S2 for details.

Two ade2 RNAi-responsive genes were further explored in vivo. We selected CG11436 because it showed the greatest response to ade2 RNAi and HtrA2 because it is associated with a pro-apoptotic function. For ade21–6 in comparison to ade21–5, in both whole wandering third instar larvae and dissected stage P1 prepupal wing discs, we found a slight reduction in expression of both genes (Figure 5B). Therefore, both CG11436 and HtrA2 are responsive to ade2 loss of function in vivo, but in an opposite direction in comparison to S2 cells.

DISCUSSION

Depletion of purine de novo synthesis causes darkened tissue to develop in pupal imaginal tissue at the P5 stage when imaginal disc eversion has occurred. This darkened tissue, or necrosis, for lack of a better term, is suppressed by the caspase inhibitor p35. Furthermore, p35 expression improves survival through metamorphosis. Thus, it appears that purine depletion at this stage of development causes a caspase-dependent cell death in imaginal discs and perhaps elsewhere. Examination of wing imaginal discs from white prepupae, before disc eversion, shows compact TUNEL-positive nuclei. We do not know whether these nuclei directly correspond to the necrotic tissue seen after disc eversion; however, the TUNEL-positive nuclei cluster in the center of the wing pouch, which becomes the distal portion of the wing after disc eversion, where we see necrosis along with the distal regions of the legs.

The term “necrosis” is being used here to describe the darkening tissue that we observe in early pupal development. It is not being used to classify the phenomenon as the caspase-independent cellular response that is often markedly distinguished from apoptosis (Edinger and Thompson 2004; Harwood et al. 2005), although non-overlapping classification of different types of cell death can be difficult (Broker et al. 2005). Darkening of imaginal tissue has also been referred to as melanization, in an effort to classify tissues undergoing an immune response mediated by hemocytes, known as melanotic masses (Minakhina and Steward 2006). Mutations in genes associated with apoptosis, such as dcp-1, show a gut melanization phenotype that is not typical of an immune response. There is no hemocyte encapsulation and tissue overgrowth typical of melanotic masses (Minakhina and Steward 2006). It is likely that the darkened “necrotic” tissue that we observe is similar to the melanization seen with mutations in genes such as dcp-1.

The folate antimetabolites methotrexate and aminopterin inhibit dihydrofolate reductase (DHFR), which is required for both purine and pyrimidine de novo synthesis in addition to serine, glycine, and methionine synthesis (McGuire 2003). These drugs appear to have quite different effects compared to mutations in purine de novo synthesis genes. For example, methotrexate causes melanotic tumors in larvae, in addition to affecting ovarian, wing, and leg development (Affleck and Walker 2007). Aminopterin causes a notched wing phenotype (Legent et al. 2006) that is distinct from that for ade2 (Tiong et al. 1989) and Prat (Clark 1994) mutants. Interestingly, apoptosis in wing discs caused by aminopterin can be suppressed by overexpression of the nucleotide salvage enzyme deoxyribonucleotide kinase (Legent et al. 2006), despite the multiple roles of DHFR.

Both Prat and Prat2 encode the same enzyme yet are essential genes with distinctive pupal lethal phenotypes. Simultaneous RNAi knockdown of both genes gives a phenotype similar to the ade2 mutant phenotype (Ji and Clark 2006). In wandering third instar larvae, Prat is expressed in imaginal discs, whereas Prat2 is expressed in the fat body (Penney et al. 2008). ade2 is expressed throughout development, including in the larval fat body (Chintapalli et al. 2007). Since we saw strong suppression of the Prat mutant phenotype, we suspect that the suppression is occurring directly by expression of p35 in imaginal discs, rather than through an indirect effect such as suppression of a fat body disorder due to purine depletion.

The expression of the UAS-p35 transgene without a GAL4 driver was sufficient to suppress ade2 and Prat mutants. We presume that expression of the UAS-p35 transgene is either due to activation of the trangene promoter by flanking enhancer elements or due to a basal level of transcription that is simply a property of this particular construction (Zhou et al. 1997). We are not aware of a previous study detecting expression of UAS-p35 alone, as the numerous studies documented in FlyBase that used UAS-p35 all used a GAL4 driver (Tweedie et al. 2009).

We did not see suppression of ade21–6 by a p53 mutation. p53 is a transcription factor that effects apoptosis in response to radiation-induced DNA damage (Brodsky et al. 2004). There is some evidence for a latent response to DNA damage that is caspase-dependent but independent of p53 (Wichmann et al. 2006); however, the factors involved in detecting this response have not been identified. Drosophila Myc appears to be an alternative sensor to DNA damage caused by radiation (Montero et al. 2008). Thus, it is possible that the reduction in purine de novo synthesis is detected through a p53-independent pathway. Mammalian cell culture studies have equivocal findings as to whether p53 is also involved in detection of reduced nucleotide pools to elicit a Go/G1 arrest (Linke et al. 1996; Bronder and Moran 2002).

We also did not see a significant suppression of ade21–6 in heterozygotes for the H99 deficiency of the reaper, hid, and grim gene region (White et al. 1994). The reduced dosage of these effector genes can suppress the apoptosis response to radiation-induced DNA damage (Brodsky et al. 2004). Therefore, our finding suggests that the caspase-dependent response that we observe may be dependent on a different apoptosis effector. Alternatively, the reduced dosage of the apopotic effector genes may simply not have been sufficient to suppress the response to a block in purine de novo synthesis.

Our gene expression analysis of S2 cells with reduced ade2 expression revealed a small set of genes with changes in transcript levels. We likely have several false negatives due to subtle effects, since the cells were cultured in media that was not depleted of purines. However, these conditions are likely to reflect situations in vivo where purines are available from the diet or from cellular turnover. HtrA2 attracted our interest as it has been quite extensively characterized with respect to a role in stress response and apoptosis. HtrA2 encodes a serine protease that is localized to the mitochondrial intermembrane space (Challa et al. 2007). It is released when the outer mitochondrial membrane is permeabilized in response to stress that leads to inhibition of the inhibitor of apoptosis protein DIAP1, as shown in S2 cells. However, others have shown that HtrA2 is not necessary for apoptosis in vivo, as there is a normal apoptotic response to gamma radiation in wing imaginal discs in HtrA2 mutants (Tain et al. 2009). HtrA2 mutants are viable, but they show male sterility and reduced stress resistance and longevity (Yun et al. 2008; Tain et al. 2009). We found a reproducible twofold increase in HtrA2 transcripts in response to ade2 knockdown in S2 cells. However, this increase was not found in ade21–6 mutant third instar larvae or in stage P1 prepupal wing discs; rather, we observed a weak decrease. Therefore, expression of HtrA2 in response to depletion of purine de novo synthesis needs to be explored further.

In conclusion, we have found a link between depletion of purine nucleotide synthesis and apoptosis in vivo. Whether purine nucleotide levels are sensed directly, or indirectly through a cellular stress response, remains to be determined. However, we found no clear evidence for involvement of p53 or the apoptosis effectors reaper, hid, and grim. Further analysis of the roles of other known members of apoptotic pathways, including HtrA2, and the other genes that we found responsive to ade2 knockdown, will be an important next step in the analysis of this response.

Acknowledgments

The authors acknowledge Sharon Gorski, Douglas Freeman, and the British Columbia Genome Sciences Centre for generously providing expertise and facilities for developing gene expression knockdown in tissue culture cells. We also acknowledge Anne Haegert and Robert Bell of the Laboratory for Advanced Genome Analysis at the Vancouver Prostate Centre, Vancouver, Canada, for carrying out the Agilent microarray hybridization and data analysis. We thank Nancy Macafee for anti-ade2 antibody purification. Microscopy was conducted with technical help from Susan Belfry of the Microscopy and Microanalysis Facility at the University of New Brunswick. This work was supported by a McCain Visitorship and Canadian Institutes of Health Research, New Brunswick Health Research Foundation, and New Brunswick Innovation Foundation grants to D.V.C.

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