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Institute for Science and Technology in Medicine (A.B., D.J.S., S.J.C., J.E.B., P.R.H., S.H., R.N.C., W.E.F.) Medical Research Unit, Keele University, North Staffordshire Hospital, Stoke-on-Trent ST4 7QB, United Kingdom; and School of Life Sciences (M.M.-M., G.T.W.), Keele University, Keele ST5 5BG, United Kingdom
Address all correspondence and requests for reprints to: Dr. W. E. Farrell, Institute for Science and Technology in Medicine, School of Postgraduate Medicine, Keele University, North Staffordshire Hospital, Stoke-on-Trent ST4 7QB, United Kingdom. E-mail: w.e.farrell{at}keele.ac.uk.
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ABSTRACT |
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2.4 fold) in active caspases in response to bromocriptine challenge in cells expressing C22orf3 relative to those harboring an empty vector control. The pituitary tumor derivation and its role in apoptosis of this gene led us to assign the acronym PTAG to this gene and its protein product. The ability of cells, showing reduced expression of PTAG, to evade or show a blunted apoptotic response may underlie oncogenic transformation in both the pituitary and other tumor types. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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, and MEG3, have been identified as being inappropriately expressed in pituitary tumors (10, 11, 12). Overexpression of pituitary tumor transforming gene and loss of expression of the growth-regulatory genes, GADD45 and MEG3, are found in the majority of pituitary tumors; however, mechanisms directly responsible for their dysregulation are yet to be defined (11, 12, 13, 14).
Recent studies have highlighted epigenetic mechanisms associated with gene silencing. Inappropriate methylation of CpG islands of key cell cycle control and growth-regulatory genes has been demonstrated for pituitary tumors that include CDKN2A/p16 (15, 16, 17, 18, 19), RB1 (20), DAP kinase (death associated protein kinase) (21) and GADD45 (22), which is associated with gene silencing. Furthermore, the association of methylation with gene silencing provides a detectable epigenetic mark for the isolation and characterization of novel genes implicated in tumorigenesis (23).
Several techniques have been described that characterize DNA methylation status including methylation-sensitive PCR (24), sodium bisulfite sequencing (25), and methylation-sensitive Southern blotting (26). A limitation of these techniques is that they require some prior knowledge of the DNA sequence being studied. However, other techniques have been described that identify novel CpG islands that are differentially methylated in cancer; these techniques include restriction landmark genomic scanning (27), methylation-sensitive representational difference analysis (28), methylation-sensitive arbitrarily primed PCR (MsAP-PCR) (29, 30, 31), CpG island microarrays (32, 33), and methyl-CpG binding domain chromatography (34). In this study we used MsAP-PCR to isolate and identify CpG islands that are differentially methylated in pituitary tumors relative to normal pituitary tissue. Because gene-associated CpG islands frequently encompass or extend into transcribed regions, subsequent sequence analysis allowed us to identify and characterize their role in this tumor type.
In this study, we report the isolation and characterization of a novel chromosome 22 CpG island-associated gene (C22orf3) that is not expressed in a significant proportion of pituitary tumors. Functional characterization of this gene, through induced expression in the pituitary tumor cell line AtT20, shows it to be a novel pituitary-derived proapoptotic gene, which we have termed pituitary tumor apoptosis gene (PTAG).
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RESULTS |
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shows representative examples of differential PCR amplicons generated by MsAP-PCR in separate pooled samples of pituitary adenomas relative to postmortem normal pituitaries. In some cases, differential amplicons were generated that were specific to a particular pituitary tumor subtype (Fig. 1A), whereas others were common to both nonfunctional adenomas and somatotrophinomas (Fig. 1B).
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200 bp, GC content >50%, CpG/GpC >0.5) (35). DNA sequence homology analysis of the CpG islands using the BLAST program revealed that nine were localized to the 5'-regions of known or novel genes and extended into their coding region. The size of each PCR amplicon, percentage of GC nucleotides, CpG/GpC ratio, and sequence homology results are summarized in Table 1. Of the nine transcript-associated CpG islands, four were isolated from the pool of nonfunctional tumors, three were isolated from the somatotrophinoma pool, and two were derived from both tumor subtypes.
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) were unmethylated in both pools of tumors and normal pituitaries, whereas four CpG islands (3, 6, 8 and 9; Table 1) showed tumor-specific methylation relative to normal pituitaries. To favor detection of frequent epigenetic change, sodium bisulfite sequencing was restricted to five molecules across each of the islands analyzed; however, we are aware that this does not represent a saturated analysis of individual islands. Thus, sequencing an increase number of molecules from each adenoma pool may well identify islands methylated as an infrequent event. Table 1 shows the number of methylated molecules (of five sequenced) in the somatotrophinoma and nonfunctional pools of tumors for each of the nine CpG islands. The CpG island associated with the novel gene C22orf3 (see Fig. 2) showed dense methylation; however, this was confined to two of five molecules in each pool of tumors. These findings, derived from pooled adenoma specimens, suggested that not all of the individual adenomas harbored this epigenetic change. However, because the CpG island associated with C22orf3 (PTAG) demonstrated the highest density and frequency of methylation relative to the other CpG islands isolated, we examined the role of this gene and its protein product in greater detail.
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qRT-PCR was performed on 38 individual pituitary adenomas that make up the major pituitary tumor subtypes. Expression of PTAG was significantly reduced relative to normal pituitary in 30 of 38 (79%) pituitary tumors. Subdivision of tumors based on subtype showed that all seven corticotrophinomas and six prolactinomas showed significantly reduced PTAG expression relative to normal pituitary. For the somatotrophinomas and nonfunctional tumors, qRT-PCR showed reduced levels of PTAG expression in eight of 11 (73%) and nine of 14 (64%) tumors, respectively. Figure 3 shows the expression of PTAG in tumors from each subtype, relative to postmortem normal pituitary.
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). In normal pituitaries and those tumors that expressed PTAG, infrequent methylation was confined to single CpG dinucleotides. Figure 4 shows representative examples of molecules sequenced from tumors and normal pituitary tissue. The low frequency of methylation-associated loss of PTAG expression suggested that aberrations other than or in addition to this epigenetic mechanism may account for the loss of PTAG transcript in pituitary adenomas.
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Functional Characterization of PTAG
Sequence homology analysis, using DNA and protein databases, identified PTAG as a novel chromosome 22 gene the protein product of which has no obvious functional domains except for a putative ubiquitin-associated domain and a c-myb DNA binding domain. To determine functional characteristics, stable transfectants of AtT20 cells were generated using an inducible expression vector harboring the complete cDNA sequence for human PTAG. Before transfection we determined the expression status of the murine homolog of PTAG (GenBank accession no. NM177370) in this cell line by RT-PCR analysis. AtT20 cells failed to express this gene relative to normal mouse pituitary, thus providing a suitable model system for functional analysis (Fig. 5A). Induced expression of PTAG was confirmed by RT-PCR analysis in response to a range of isopropyl-ß-D-thiogalactopyranoside (IPTG) concentrations (Fig. 5B). Growth curve analysis over 12 d in the presence of inducing agent (IPTG) showed no discernable difference in doubling time or cell viability as assessed by trypan blue exclusion relative to control cells harboring the empty expression vector (Fig. 5C).
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AtT20 cells were treated with a range of drug doses for 72 h, before acridine orange staining and assessment of morphological changes characteristic of apoptosis (Fig. 6A). There was a dose-dependent increase in the number of cells showing apoptotic morphology, irrespective of PTAG expression status. However, PTAG-expressing cells showed an enhanced apoptotic response to bromocriptine challenge. At the highest drug dose (40 µM) there was a 3-fold increase (P = 0.001) in the number of apoptotic cells relative to control cells harboring an empty vector.
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). Irrespective of PTAG expression status there was an approximately 4-fold decrease in the percentage of bromocriptine-induced apoptotic cells pretreated with z-VAD-fmk, relative to cells incubated in the presence of bromocriptine alone. These results suggest that the enhanced proapoptotic response observed in bromocriptine-challenged cells expressing PTAG is most likely mediated through caspase activation.
PTAG-Associated Increase of Caspase Activation
To directly determine PTAG-associated caspase activation in the apoptotic response, a fluorochrome-labeled general caspase inhibitor (fam-VAD-fmk) was used to detect relative levels of active caspases over a 24-h time course (Fig. 7). There was a time-dependent increase in the levels of active caspases in AtT20 cells treated with 40 µM bromocriptine, irrespective of PTAG expression status. However, at all time points PTAG-expressing cells showed an enhanced response. At the 24-h time point PTAG-expressing cells showed a 2.4-fold (P = 0.001) increase in caspase activation relative to control cells.
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). A time-dependent increase in the percentage of TUNEL-positive cells was observed for both PTAG-expressing and control cells. At all time points, PTAG-expressing cells showed higher levels of TUNEL-positive cells, and at 72 h showed a 2.5-fold increase (P = 0.002) relative to control cells.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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To increase the likelihood of identifying gene-specific differential methylation in tumors vs. normal pituitary, pooled tumor cohorts were used, and subsequent methylation status was determined within the pool by bisulfite sequencing. Within each of these pools and in contrast to normal pituitary, bisulfite sequencing showed tumor-specific differential methylation of PTAG. However, in each adenoma pool (somatotrophinoma and nonfunctional), although methylation of the PTAG-associated CpG island was dense, it was confined, in each case, to two of five molecules, suggesting that not all of the individual adenomas within the pools were methylated.
Several previous studies have employed MsAP-PCR to isolate DNA sequences that are differentially methylated in a range of cancer types including colon, breast, and lung (29, 30, 31). In these studies, confirmation of methylation was determined by methylation-sensitive restriction digestion and Southern blotting. We have extended this approach by using bisulfite sequencing as a rigorous method of determining the methylation status of differentially isolated products. Whereas previous studies have successfully employed MsAP-PCR to identify novel altered methylation profiles associated with oncogenic transformation, correlations between this change and cognate gene expression status have not been described. To determine associations between methylation and expression of PTAG, we used qRT-PCR of cDNA derived from individual pituitary adenomas that made up the major subtypes. Relative to normal pituitary, these studies showed that a significant proportion (79%) of adenomas, irrespective of subtype, failed to express this gene, suggesting that loss of PTAG transcript is a common underlying aberration in pituitary tumors. Sodium bisulfite sequencing of these individual adenomas showed that loss of expression was infrequently associated with methylation of this gene’s CpG island. However, in those adenomas that were methylated, we did not detect expression of PTAG. Although these findings would suggest mechanisms in addition to methylation to be responsible for loss of PTAG, our analysis did not reveal genetic change as determined by either LOH or sequence analysis as an alternate mechanism. In this context, recent studies have described loss or substantially reduced expression of both GADD45 and MEG3 transcripts in a significant proportion of pituitary tumors (11, 12). Although our own studies have now described a significant correlation between methylation of the GADD45 gene’s CpG island and loss of transcript expression (22), mechanisms responsible for loss of MEG3 await further experimental study.
The isolation of PTAG was performed on the basis of discriminating sequences differentially methylated in pituitary tumor DNA relative to normal tissue. The use of pooled tumor DNA for MsAP-PCR greatly increased the probability of identifying sequences that are methylated. Indeed, within the pooled tumors used to isolate this novel gene, our sequencing of individual clones (molecules) confirmed this. In these cases, although methylation was dense, it was confined to two of five molecules. However, the high frequency of loss across individual adenomas, as determined by qRT-PCR, together with infrequent methylation and the absence of mutations or genetic loss, suggests either more subtle changes outside of the regions we have examined or dysfunction in trans-acting factors that regulate expression of this gene.
Because database searches revealed no obvious functional roles for the PTAG gene, we initially determined the consequences of enforced PTAG expression on proliferation in the mouse corticotroph cell line AtT20. In the absence of a suitable human pituitary cell line we considered AtT20 cells a suitable model system because they do not express the murine homolog of PTAG as determined by RT-PCR analysis, and our previous studies had shown that enforced expression of p16 in this cell line resulted in a G1-mediated growth arrest (37). However, enforced expression of PTAG in these stable transfected cells did not result in either growth arrest or decrease in cell viability. In contradistinction to these finding, enforced expression of PTAG in AtT20 cells significantly augmented the apoptotic response to bromocriptine challenge, suggesting that the function of this gene is linked to programmed cell death as a proapoptotic mediator.
We initially assessed bromocriptine-induced apoptosis by acridine orange staining, which showed that a greater proportion of PTAG-expressing cells displayed apoptotic morphology relative to cells harboring an empty-vector control. In addition, the general caspase inhibitor z-VAD-fmk significantly reduced the number of apoptotic cells, suggesting that apoptosis was through a caspase-mediated pathway. Increased levels of active caspases in PTAG-expressing cells were more directly confirmed in time course experiments using a fluorescent labeled caspase inhibitor that binds irreversibly to activated caspases. To further define the role of PTAG as a proapoptotic mediator, the latter stages of this process, DNA fragmentation, were measured by TUNEL labeling. In response to bromocriptine, PTAG-expressing AtT20 cells were significantly more sensitive to challenge that those harboring an empty vector.
Previous studies have shown that the pituitary cell lines AtT20 and GH3 show decreased proliferation and increased apoptosis associated with increased p53 expression and reduced bcl-2 levels in response to bromocriptine challenge (38, 39, 40); however, it is not clear whether this response is mediated through D2 receptors (41, 42, 43). Our data also show, at least in AtT20 cells, that the absence of the murine homolog of PTAG did not confer complete protection from bromocriptine-induced apoptosis. However, the augmented apoptotic response, seen in these cells through enforced expression of PTAG, would support its role as a mediator rather than an obligate effector in this pathway. The intracellular cascades activated in both receptor-mediated extrinsic and stress-induced intrinsic apoptotic pathways are complex with activation of proximal and subsequent effector caspases (44, 45). In addition, cooperation between these mediators and amplification of the extrinsic pathway through cross-talk with the intrinsic pathway are also apparent (46). Thus, singular loss of a proapoptotic mediator within these pathways may result in a blunted apoptotic response. In this context, in mouse embryo fibroblasts derived from either bad- (47) or bax-deficient (48) mice, singular loss of these characterized proapoptotic genes shows that they do not account exclusively for physiological cell death.
The frequent loss of PTAG expression in pituitary tumors suggests that this is an early change in pituitary tumorigenesis leading to a blunted apoptotic response. Indeed, several studies have suggested a central role for a compromise apoptotic response in tumor evolution and progression (45, 49, 50). In these cases, resistance to apoptosis will allow damaged cells to proliferate, thus accruing additional genetic damage and tumor outgrowth. Whereas the role of this novel proapoptotic gene in other tumor types awaits investigation, its reactivation in tumors of pituitary origin may sensitize these cells to known and perhaps previously untested apoptosis-inducing agents.
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MATERIALS AND METHODS |
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DNA and RNA Preparation
DNA was prepared from frozen pituitary tissue using the Nucleon DNA extraction kit (Anachem, Bedfordshire, UK) as described by the manufacturer. Total RNA was prepared from frozen pituitary material as previously described (53).
Restriction Enzyme Digestion of Genomic DNA
To determine differences in the methylation pattern between tumor and normal pituitary tissue, pooled DNA samples were first subject to cleavage with a panel of methylation-sensitive restriction enzymes.
Separate DNA pools comprised a cohort of 10 somatotrophinomas, 10 nonfunctional tumors, and five postmortem normal pituitaries, respectively. Specimens were selected from male individuals to avoid confounding effects of X-chromosome inactivation. A total of 20 µg of DNA for each pooled sample were first treated with a panel of methylation-sensitive restriction enzymes. Briefly, restriction digest reactions were performed using SmaI (100 U) at 25 C for 6 h, SacII (100 U) and HpaII (50 U) at 37 C for a further 16 h, and finally BstUI (100 U) at 60 C for 6 h, after which DNA was precipitated with ethanol and resuspended in 70 µl H2O. Samples were then digested using 50 U of the methylation-insensitive enzyme MseI by incubating at 37 C for 6 h, purified by phenol/chloroform extraction, and stored at –20 C. All restriction enzymes were purchased from New England Biolabs (Hertfordshire, UK).
MsAP-PCR
Differentially methylated sequences within tumor samples were identified by MsAP-PCR using arbitrary primers; LI (5'-CGTTCGTATCGACGGCGCGA-3'), BS5 (5'-CTCCCACGCG-3'), and BS13 (5'-CGGGGCGCGA-3') as previously described (29). PCRs were performed using 0.5 µg DNA from the digested pooled tumors, normal pituitaries, and undigested normal pituitary, in a total of 50 µl containing 10 mM Tris-HCl (pH 9.0), 50 mM KCl, 0.1% Triton-X, 1 U Taq DNA polymerase, 200 µM each of dATP, dCTP, dGTP, dTTP, 0.5 M Betaine, 2% (vol/vol) dimethylsulfoxide, and 40 pmol of arbitrary primer. Because the primers used are arbitrary in nature, but biased toward CpG-rich regions, the same primer may be used in both the forward and reverse reaction. Therefore, PCRs were performed using either a single primer or a combination of two primers. In all cases, PCRs were performed using the following conditions: 1 cycle at 96 C for 5 min, 35 cycles at 42 C for 1 min, 72 C for 1 min, 94 C for 0.5 min, and a final extension at 72 C for 5 min. PCR products were resolved on 8% polyacrylamide gels and visualized by silver staining, as previously described (21).
Isolation and Characterization of DNA Fragments
Following PAGE, PCR products appearing to be differentially methylated by MsAP-PCR in pooled tumors vs. normal pituitaries were excised and eluted into sterile H2O. The eluate (2–5 µl) was reamplified using the same primer sets used in the original MsAP-PCR, under the same conditions as described above. Amplified products were resolved on 1% (wt/vol) agarose gels stained with ethidium bromide, and PCR products were excised and purified using the GeneClean II kit (Anachem, Bedfordshire, UK) followed by cloning into the PGem TA vector (Promega, Southampton, UK). Individual colonies were sequenced using a Big Dye Terminator cycle sequencing protocol (PE Applied Biosystems, Cheshire, UK), on an ABI Prism 310 Genetic analyzer, and data were analyzed with Sequencing Analysis V3.0 software (ABI Prism, PE Applied Biosystems). Sequence data were used to determine genomic information including homology to characterized or novel genes and cytogenetic map positions employing the Ensembl database (http://www.ensembl.org). Furthermore, each sequence was analyzed for CpG content and CpG/GpC ratio using the GrailEXP (http://compbio.ornl.gov/grailexp) program to predict conformity to criteria defining CpG islands (35).
Sodium Bisulfite Modification of DNA
Sodium bisulfite conversion before sequencing was carried out as previously described (21). Briefly, 5 µg DNA were denatured by incubating at 37 C for 15 min in the presence of 0.4 M NaOH; 370 µl of freshly prepared sodium bisulfite solution (2.2 M sodium bisulfite, 8 M urea, 10 mM hydroquinone, pH 5.0) was added to the DNA and incubated at 55 C for 4 h. Modified DNA was recovered using the Gene Clean II kit (Anachem), followed by desulfonation of the modified DNA by incubating samples with 0.3 M NaOH at 37 C for 15 min. DNA was precipitated with ethanol and resuspended in 20 µl of TE buffer (pH 8.0).
Bisulfite Sequencing of CpG Islands
We employed bisulfite sequencing to determine the methylation status of the CpG islands isolated by MsAP-PCR, using primers designed according to criteria previously described (25). Primer sequences are available on request. PCRs were performed with 1–2 µl of modified DNA in 50 µl containing 10 mM Tris-HCl (pH 9.0), 50 mM KCl, 0.1% Triton-X, 1 U Taq DNA polymerase, 200 µM each of dATP, dCTP, dGTP, dTTP, and 20 pmol of each primer. PCR products were resolved by agarose gel electrophoresis, cloned into the PGem TA vector (Promega), and individual molecules were sequenced as described above.
Quantitative RT-PCR Analysis of C22orf3 (PTAG)
We employed qRT-PCR to determine expression status of PTAG in pituitary tumors relative to normal pituitary. Total RNA (5 µg) was used for cDNA synthesis using Superscript II RT according to the manufacturer’s instructions (Invitrogen, Paisley, Scotland, UK). Quantitative PCR was performed using 1 µl cDNA, corresponding to 250 ng initial RNA, and TaqMan MGB probes specific to PTAG (Assay ID: Hs00202161_m1) and 18S rRNA (Assay ID: Hs99999901_s1) as an endogenous control, according to manufacturer’s instructions (Applied Biosystems, Warrington, UK). Quantitation of PTAG gene expression in pituitary tumors relative to normal pituitary was determined using the relative standard curve method, with normal pituitary as the calibrator. Real time fluorescence measurement of PCR samples was performed using an ABI Prism 7000 sequence detection system, and data were analyzed using ABI Prism 7000 SDS software.
Sequence and LOH Analysis of PTAG
The coding sequence of PTAG was sequenced using intronic primers designed to encompass each of the five coding exons (Table 2). Tumor and normal pituitary DNA (200 ng) was subject to PCR amplification as described above, and PCR amplicons were subject to cycle sequencing reactions (as described above) using both sense and antisense PCR primers.
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). Products were resolved on 8% nondenaturing polyacrylamide gels and visualized by silver staining as described (21). LOH was defined in tumor DNA by a reduction in allele intensity of greater than 80% (as assessed by scanning densitometry) or by the absence of one of the PCR-amplified alleles from tumor DNA.
Generation of Stable Transfected Cell Lines
The complete coding sequence of human PTAG (GenBank accession no. NM012265), obtained in a clone from the Mammalian Gene Collection (MGC: 3580, Cambridge, UK), was subcloned into a modified Gateway-compatible inducible vector, pOPRSVI (Stratagene, Cambridge UK). Recombination-mediated transfer was performed using the Gateway cloning system according to the manufacturer’s instructions (Invitrogen), and orientation and sequence integrity of the insert were confirmed by sequence analysis.
The mouse pituitary cell line AtT20 was characterized as a suitable expression model, because these cells do not express endogenous PTAG as assessed by RT-PCR analysis using primers specific to the murine homolog of PTAG (GenBank accession no. NM177370) and porphobilinogen deaminase (PBGD) (GenBank accession no. NM013551) as an internal control (Table 2). The inducible vector containing the complete coding sequence of PTAG (pOP-PTAG) was transfected into AtT20 cells, harboring the constitutive expression vector (pCMVlacI) for the lac repressor (lacI), and stable transfectants generated as previously described (37). Control transfectants were also generated harboring the empty expression vector (pOP-MCS). Cells were cultured as previously described (37), and selection was performed in the presence of 500 µg ml–1 Geneticin and Hygromycin B (Invitrogen) followed by soft agar cloning to isolate single clones. Maximal expression of PTAG was determined in IPTG dose-response experiments and RT-PCR analysis (see Table 2 for primers and PCR conditions) as previously described (37).
Growth Curve Analysis
The effect of induced expression of PTAG was assessed in stable transfected clones (AtT20-PTAG) relative to a clone stably transfected with empty vector (AtT20-MCS). Cells were seeded at a density of 2.5 x 105 per 75 cm2 flask in 5 ml of medium and induced with 5 mM IPTG or PBS alone, and media were replenished at 2-d intervals, as previously described (37). Triplicate individual flasks were sacrificed at each time point (2 d) and cells were counted and their viability determined by trypan blue exclusion. The experiment was repeated three times.
Detection of Apoptosis by Acridine Orange Staining
AtT20-PTAG and AtT20-MCS control cells (2 x 105) were seeded into individual wells of a six-well cell culture plate in 2 ml of medium and induced with 5 mM IPTG for 48 h before bromocriptine challenge. Cells were assayed for apoptotic response as described (54). Briefly, adherent and nonadherent cells were collected and washed in PBS before being pelleted; 10 µl of cell suspension were then mixed with 10 µl acridine orange (50 µg ml–1 in PBS) and analyzed on a wet mount slide using a Leica DMR fluorescent microscope (Leica Corp., Deerfield, IL). The percentage of cells with apoptotic morphology (nuclear and cytoplasmic condensation, nuclear fragmentation, membrane blebbing, and apoptotic body formation) was determined from 200 cells in each of three separate fields, and the mean from three independent experiments was determined. Where indicated, bromocriptine challenge was also performed in the presence of 50 µg ml–1 of the general caspase inhibitor z-VAD-fmk (Flowgen, Leicestershire, UK) before staining.
Measurement of Caspase Activation in Situ by Fluorochrome-Labeled Inhibition of Caspases (FLICA)
Caspase activity in AtT20 cells was measured using the fluorochrome (FAM)-labeled broad spectrum inhibitor of caspases (fam-VAD-fmk). This reporter molecule irreversibly binds to active caspases allowing measurement of caspase activation in situ (55). AtT20-PTAG and AtT20-MCS control cells were seeded and induced with IPTG as described above and were then challenged with 40 µM bromocriptine for 72 h before labeling according to the manufacturer’s instructions (Flowgen). Briefly, a 300-µl aliquot of cells at a density of 1 x 106 ml–1 was labeled using 10 µl of a freshly prepared 30x working dilution of fam-VAD-fmk by incubation at 37 C and 5% CO2 for 15 min, followed by two washes with 2 ml of a 1x working dilution wash buffer. Cells were resuspended in 320 µl PBS and placed on ice before analysis. Fluorescence was measured at an excitation wavelength of 485 nm and emission at 535 nm using an automated fluorescence plate reader (Wallac 1420 Victor, Milton Keynes, UK).
DNA Fragmentation Labeling Assay (TUNEL)
Apoptosis-induced DNA fragmentation was measured using the APO-BRDU kit (BD Biosciences, Oxford, UK). Cells were seeded, induced with IPTG, and challenged with 40 µM bromocriptine as described above, followed by fixation in 1% paraformaldehyde in PBS at 4 C for 30–60 min, postfixed in 70% ethanol and stored at –20 C for 12–18 h before staining. Briefly, fixed cells (1 x 106) were washed twice with PBS before treatment with a TUNEL reaction mix (bromolated deoxyuridine triphosphate, terminal deoxynucleotidyl transferase, and reaction buffer) for 60 min at 37 C, followed by washing and incubation with a fluorescein isothiocyanate-labeled bromodeoxyuridine monoclonal antibody for 30 min at room temperature. TUNEL-positive cells were counted from 200 cells in each of three separate fields for each measurement, using a Leica DMR fluorescent microscope. The mean from three separate experiments was determined.
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FOOTNOTES |
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Abbreviations: FLICA, Fluorochrome-labeled inhibition of caspases; IPTG, isopropyl-ß-D-thiogalactopyranoside; LOH, loss of heterozygosity; MsAP-PCR, methylation-sensitive arbitrarily primed PCR; PBGD, porphobilinogen deaminase; PTAG, pituitary tumor apoptosis gene; qRT-PCR, quantitative RT-PCR.
Received for publication March 2, 2004. Accepted for publication April 16, 2004.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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