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From the * Institute for Science and Technology in
Medicine, Keele University, Keele, United Kingdom; and
King's College London, Department of
Haematological Medicine, The Rayne Institute, London, United Kingdom.
| Correspondence to: Professor G. T. Williams, Huxley Building, School of Life Sciences, Keele University, Keele ST5 5BG, United Kingdom (e-mail: g.t.williams{at}biol.keele.ac.uk). |
| Received for publication April 22, 2008; accepted for publication November 5, 2008. |
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Abstract |
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 Top Abstract Materials and MethodsResultsDiscussionReferences |
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0.02% control cells)
to 3 or more apoptotic inducers. These clones could be infected at a high
efficiency (>90%) with a lentiviral vector. In conclusion, we have isolated
clones of nontumorigenic prostate cells (PNT2C2), androgen-sensitive prostate
cancer cells (22Rv1), and androgen-independent, metastatic prostate cancer
cells (PC-3), which are suitable as host cells for functional cloning studies
to address cell death control mechanisms in the prostate during cancer
progression.
Key words: Cell death, prostate cancer, PNT2, 22Rv1, PC-3
A better understanding of the key regulatory molecules that control the life/death decisions of prostate cancer cells would greatly assist in the development of new gene and drug therapies for prostate cancer. In recent years, much progress has been made in identifying the key molecular components of the apoptotic machinery in mammalian cells, but our detailed understanding of the molecular mechanisms governing the higher levels of apoptosis control is still seriously incomplete. Forward genetics approaches, such as functional expression cloning, have provided powerful strategies for the isolation of genes which alone control the critical rate-limiting steps of cell death, at least in cells other than prostate epithelia (Cohen and Kimchi, 2001; Williams and Farzaneh, 2004). Importantly, these methods allow the identification of entirely novel regulators of apoptosis, in addition to known apoptosis-controlling genes (Williams et al, 2006). Such critical regulators of apoptosis are likely to prove attractive targets for the design of novel drug therapies.
Functional expression cloning comprises the following steps: 1) mutation of host cells (eg, by cDNA/siRNA library expression or retroviral insertional mutagenesis), 2) application of a stimulus to induce cell death, and 3) characterization of mutations in surviving, cell death–resistant clones (Williams and Farzaneh, 2004). Before such approaches can be applied to the prostate, it is necessary to obtain prostate cells that meet several criteria. They must grow well in vitro, be monoclonal, be easily infected at high efficiency with viral vectors, be easily cloned, and undergo apoptosis with minimal survival of spontaneously apoptosis-resistant cells. With respect to the first criterion, a diverse range of human prostate epithelial cell lines are available, including normal cell lines that have been immortalized in vitro and clinical isolates from patients with prostate cancer (van Bokhoven et al, 2003). We have selected 5 cell lines, ranging from nontumorigenic prostate cells to those isolated from androgen-independent metastatic tissue, for further study. We report here the development of clones of 3 prostate cell lines that are suitable for the application of functional cloning studies.
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Materials and Methods |
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TopAbstractMaterials and MethodsResultsDiscussionReferences |
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Cell Cloning
Three different cloning media were tested, all of which were supplemented
with fetal bovine serum (20% [vol/vol]), gentamicin (50 µg/mL), and
cell-conditioned routine culture medium (10% [vol/vol], prepared from actively
growing cultures). Base media were: 1) Iscove modified Dulbecco medium
supplemented with L-glutamine (2 mM); 2) Dulbecco modified Eagle
medium–F-12 nutrient mixture (1:1) containing L-glutamine (2
mM), and 3) routine culture medium. A double-layer soft agar method was
employed in 10-cm diameter culture plates, comprising cells in 0.3% (wt/vol)
Difco Noble agar (Lab Manchester, Manchester, United Kingdom) laid over a 0.5%
(wt/vol) Noble agar base; both layers made up in cloning medium. Agar was
overlaid with cloning medium. A range of cell densities (usually 500–50
000 cells per dish) were plated to ensure the production of single,
well-separated colonies. After 3–4 weeks of incubation, the medium
overlay was removed, colonies were picked, and the plugs were transferred to
wells of 96-well culture plates containing fresh cloning medium. When
50%–70% confluent, clones were trypsinized and expanded by serial
transfer to 24-well, 12-well, and 6-well plates, and then T25 and T75 flasks.
Cloning medium was gradually replaced with routine culture medium at the 12-
to 6-well plate stage.
Induction of Cell Death
Chemical inducers of apoptosis (doxorubicin, etoposide, okadaic acid, and
sodium butyrate) were purchased from Sigma-Aldrich, whereas the agonistic
anti-Fas monoclonal antibody IPO4 was from Dr S. Sidorenko (University of
Washington, Seattle, Washington). Cells were routinely plated in maintenance
medium at a density of 1 x 104 cells/cm2 either in
96-well culture plates (for screening of chemical inducers of cell death) or
in 35-mm diameter culture dishes (for clone characterization). After
incubation for 24 hours, chemical agents (or vehicle for controls) were added
and cells were cultured for a further 72 hours. For ultraviolet (UV)
irradiation, cells were plated in 35-mm diameter culture dishes as above.
After 24 hours of incubation, cells were irradiated with the indicated doses
of UV at 254 nm; doses were verified using a UV (shortwave) intensity meter.
Controls were mock irradiated. Immediately after exposure, the medium was
changed, and cells were cultured for 72 hours.
Determination of Androgen Sensitivity
In experiments with the synthetic androgen R1881 (PerkinElmer LAS Ltd,
Beaconsfield, United Kingdom), cells were plated in Phenol red–free
RPMI-1640 medium supplemented with L-glutamine (2 mM), dextran
charcoal-stripped fetal bovine serum (10% [vol/vol]; Perbio Science UK), and
gentamicin (50 µg/mL; prfR-10dcss) in 96-well culture plates (at 1 x
104 cells/cm2). After 2 hours, R1881 (or vehicle for
controls) was added, and cells were incubated for the indicated time, with
medium changes every 3–4 days. In experiments with the antiandrogen
Casodex (AstraZeneca, Macclesfield, United Kingdom), cells were washed with
prfR-10dcss and replated in 6-well plates (at 1 x 104
cells/cm2) in prfR-10dcss supplemented with testosterone (10 nM)
and either Casodex or vehicle (controls). Cells were incubated for the
indicated times, with medium changes every 3–4 days.
Culture Growth Assays
Growth was assessed by a tetrazolium salt (MTS) assay or by direct
microscopic counting. The MTS assay comprised CellTiter 96 AQueous One
Solution Cell Proliferation Assay (Promega UK, Southampton, United Kingdom),
which was used according to the manufacturer's instructions. Medium blanks
containing the drugs under test were run with each assay, and the absorbance
readings at 490 nm (A490) were subtracted from the appropriate test readings.
For direct microscopic counting, adherent cells were collected by
trypsinization, combined with nonadherent cells, stained with nigrosin blue
(0.1% [wt/vol]), and counted using a hemocytometer. Cells that excluded the
dye were considered to be viable.
Clonogenic Growth Assays
Long-term survival of cells was determined by colony-forming assays. For
initial studies with parental cells, cells were harvested by trypsinization,
combined with nonadherent cells, and washed. Cells were resuspended in fresh
medium (0.6 mL/cm2 culture area), and proportions (usually
2.5–10.0 µL for control cells and 10–100 µL for treated
cells) were plated in 96-well culture plates in 0.2 mL final volume of culture
medium. After 1–3 weeks of incubation (cell line dependent), colonies
consisting of 10 cells or more were counted using a phase-contrast light
microscope under low-power magnification; only wells containing well-separated
colonies were scored. The number of colonies formed per milliliter of cell
suspension was calculated and averaged over the dilution range. Modified
assays of increased sensitivity were used for studies with cell clones. Thus,
harvested cells (adherent plus nonadherent) were microscopically counted after
nigrosin blue staining and, based on the viable cell count in control cell
suspensions, control and treated cell suspensions were diluted to give 1.6
x 104 cells/mL. Serial dilutions were prepared, and 0.1 mL
was plated into wells of a 96-well plate: for control suspensions,
20–200 cells were plated; for treated suspensions, 500–50 000
cells were plated. The number of colonies formed per nominal 1000 cells was
calculated and averaged over the dilution range.
Assessment of Apoptosis
Apoptosis was monitored in trypsinized cells by assessment of caspase
activation using a commercial kit (CaspaTag fluorescein caspase activity kit;
Intergen, Purchase, New York), which contains a carboxyfluorescein derivative
of benzyloxycarbonyl valylalanylaspartic acid fluoromethyl ketone, according
to the manufacturer's instructions. Cells also were stained with Hoechst 33342
to visualize cell nuclei. The proportion of apoptotic cells was determined by
fluorescence microscopy and direct counting.
Infection With Lentiviral Vector
Cells were infected at 5 x 104 cells per well of 24-well
plates with vesicular stomatitis virus–enveloped, self-inactivating
lentivirus, which was carrying a spleen focus-forming virus
promoter–driven enhanced green fluorescent protein (EGFP) and prepared
from biotin-labeled BL15 cells as described in detail previously
(Chan et al, 2005;
Nesbeth et al, 2006). Briefly,
streptavidin paramagnetic particle viral concentrates were recovered from
frozen, washed thoroughly, and used to infect prostate cell line clones and
positive control human TE671 cells. The titer of virus was determined by flow
cytometry on TE671 cells to be 3.4 ± 0.3 x 109/mL.
Using these titers, it is calculated that prostate clones were infected at a
TE671 multiplicity of infection of approximately 7. After 6 days in culture
(including 1 subculture onto a slide chamber), cells were fixed in 2%
paraformaldehyde for 30 minutes at room temperature, washed, permeabilized
with 0.2% Triton X-100 in phosphate-buffered saline for 15 minutes, washed,
and stained with 4'6-diamidino-2-phenylindole-2HCl (DAPI). Cells were
observed for EGFP expression and DAPI staining by fluorescence microscopy.
Statistics
All values are means ± SEM. Data were analyzed either by a paired
Student's t test or, for more than 2 groups, by 1-way analysis of
variance. In the latter case, posthoc analysis was either by Bonferroni's
multiple-comparison test or Dunnett's multiple-comparison test (when comparing
multiple groups vs a single group). Homogeneity of variance was checked by
Bartlett's test and, where necessary, data were transformed prior to
analysis.
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Results |
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TopAbstractMaterials and MethodsResultsDiscussionReferences |
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The growth of the various cell lines showed differential sensitivity to etoposide (Figure 1A). PNT2C2 and P4E6 cells were characterized by similar dose-response curves, with 50% growth inhibition occurring at around 10 µg/mL etoposide, and approximately 90% growth inhibition being achieved at the highest etoposide doses studied. Dose-response curves were shifted toward higher etoposide concentrations for LNCaP and for PC-3 cells in particular. For LNCaP cells, growth inhibition was maximal at around 60% at etoposide concentrations of 50–200 µg/mL, whereas for PC-3 cells, 50% growth inhibition was achieved only with the highest concentration (200 µg/mL) of etoposide studied.
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Based on these findings, clonogenic assays were performed at selected etoposide concentrations. Clonogenic activity of PNT2C2 cells was effectively inhibited by 25 µg/mL etoposide, whereas colony formation was detectable for LNCaP and PC-3 cells treated with 200 µg/mL etoposide (Table). The clonogenic activity of 22Rv1 cells was completely abolished by 50 µg/mL etoposide (Table).
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Doxorubicin inhibited the growth and clonogenic activity of all parental
cell lines. With respect to growth, 22Rv1 cells appeared the most sensitive,
followed by LNCaP and P4E6 cells, with PNT2C2 and PC-3 cells exhibiting the
least sensitivity (Figure 1B).
Clonogenic activity of P4E6 and PC-3 cells was abolished by doxorubicin at
0.25 µg/mL, and that of PNT2C2, 22Rv1, and LNCaP by doxorubicin at
0.5–1.0 µg/mL
(Table).
Okadaic acid inhibited culture growth in most cell lines (Figure 1C). Growth inhibition tended to reach a maximum at doses between 50 and 100 nM okadaic acid and appeared more complete for PNT2C2 and PC-3 cells (80%–85% inhibition) than for P4E6 (55% inhibition) and LNCaP (40% inhibition) cells. In contrast, data from clonogenic assays demonstrated that LNCaP and 22Rv1 cells were more sensitive to okadaic acid than either PNT2C2 or PC-3 cells, because LNCaP and 22Rv1 cells pretreated with 20 nM okadaic acid failed to form colonies, whereas PC-3 cells formed colonies at this dose (undetectable at 50 nM okadaic acid), and PNT2C2 cells formed colonies even after pretreatment with 100 nM okadaic acid (Table).
Sodium butyrate, a histone deacetylase inhibitor, has been shown to induce apoptosis in several prostate cell lines (including LNCaP) in culture, but not in PC-3 cells (Frønsdal and Saatcioglu, 2005). Indeed, butyrate inhibited culture growth of P4E6, LNCaP, and 22Rv1 cells (Figure 1D). Clonogenic survival was evaluated in LNCaP cells only; respective values at 1, 5, and 10 mM sodium butyrate, relative to untreated controls, were: 1.49% ± 0.23%, 0.07% ± 0.03%, and <0.03% (detection limit).
Fas stimulation using the antibody IPO4 inhibited the growth of P4E6 and PC-3 cells (Figure 1E) but had no effect on the growth of PNT2C2 or 22Rv1 cells (data not shown). P4E6 cells were markedly more sensitive than PC-3 cells, and this was mirrored in the clonogenic survival. Thus, for P4E6 cells (n = 2 experiments), clonogenic survival was 1.65% ± 0.99% control at 0.10 µg/mL IPO4 and fell below the detection limit of the assay (0.26% control) at 0.25 µg/mL IPO4, whereas for PC-3 cells (n = 3 experiments), values were 12.86% ± 1.09% control at 1 µg/mL IPO4 and 10.83% ± 1.85% control at 10 µg/mL IPO4.
For UV irradiation, culture growth was assessed by direct microscopic counting of viable cells. Clonogenic assays were performed as normal. Doses of up to 120 J/m2 inhibited culture growth and clonogenic activity of all cell lines (Figure 2). Growth of all cell lines showed similar sensitivity to UV, as exemplified by PNT2C2 and LNCaP cells (Figure 2A). However, the clonogenic activity of LNCaP cells appeared less markedly affected than growth (Figure 2B). Nevertheless, at a UV dose of 80 J/m2, clonogenic activity of PNT2C2 cells was undetectable (detection limit of 0.03% control), and at a UV dose of 120 J/m2 it was reduced to less than 2% control value in P4E6, 22Rv1, and PC-3 cells and to less than 10% control value in LNCaP cells (Figure 2C).
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PNT2C2, 22Rv1, and PC-3 cells were successfully cloned in soft agar, and the resulting clones were initially screened for clonogenic survival after exposure to a single dose of UV (120 J/m2). For each cell line, clones were obtained with a range of sensitivities to UV (Figure 3).
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Further Characterization of PNT2C2 Clones
Four PNT2C2 clones (clones 2, 5, 9, and 11) displayed acceptable levels of
spontaneous apoptosis resistance (<0.01% control cells) to UV irradiation
(Figure 3A) and were subjected
to further rounds of screening with chemical inducers of apoptosis. An initial
experiment revealed unacceptably high levels of clonogenic survival (>0.3%
control values) for clones 2 and 9 with etoposide (10 µg/mL) and okadaic
acid (100 nM), respectively, so these clones were excluded from further study.
In contrast, in further experiments, clones 5 and 11 displayed reproducibly
acceptable levels of resistance to these 2 chemical agents
(Figure 4). Treatment with
etoposide and okadaic acid for 24 hours induced apoptosis in parental cells
and clones (Figure 4). Compared
with parental cells, a greater proportion of clone 5 and clone 11 cells
displayed caspase activation in response to okadaic acid
(Figure 4).
The growth of clones 5 and 11, as well as parental cells, was unresponsive to treatment with the synthetic androgen R1881 over an incubation period of 43–162 hours, indicating that these cells are androgen insensitive (data not shown).
Further Characterization of 22Rv1 Clones
Clonogenic survival of 22Rv1 clone 8 also was evaluated with etoposide (200
µg/mL), okadaic acid (200 nM), and doxorubicin (1 µg/mL), and was
undetectable (<0.004% control) with each agent, whereas corresponding
values for parental cells were 0.0081% ± 0.0021%, 0.0149% ±
0.0077%, and 0.0404% ± 0.0046% control, respectively. When assessed
after 24 hours of treatment, etoposide and okadaic acid induced apoptosis in
parental and clone 8 cells, whereas doxorubicin produced a clear-cut increase
in the proportion of apoptotic cells in clone 8 cells only
(Figure 5). Furthermore,
okadaic acid induced apoptosis in a greater proportion of clone 8 cells than
in parental cells at this time point
(Figure 5).
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The growth of parental 22Rv1 cells is known to be androgen sensitive (Sramkoski et al, 1999), so it was of interest to determine whether clone 8 retains androgen sensitivity. Growth of this clone was stimulated to an extent similar to parental cells by the synthetic androgen R1881 (Figure 6A). Prolonged treatment with the anti-androgen Casodex inhibited clonogenic survival of both parental and clone 8 cells, with the latter showing a slightly reduced survival compared with parental cells at 20 µM Casodex only (Figure 6B).
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Discussion |
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TopAbstractMaterials and MethodsResultsDiscussionReferences |
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Initially, we studied a total of 5 cell lines representative of a broad spectrum of prostate disease, and we evaluated their responses in terms of culture growth and clonogenic activity to a range of treatments that have been shown previously to induce apoptosis in prostate cells. The main aim of this part of the study was to identify agents capable of inducing effective cell death, because 3 of the cell lines (PNT2C2, P4E6, and 22Rv1) were relatively uncharacterized with respect to apoptotic stimuli. As would be predicted from previous work with prostate cell lines, the effects of the various agents were both cell line dependent and treatment dependent. In general, the PNT2C2 cell line showed the highest sensitivity to most chemical agents and UV in terms of both culture growth and clonogenic activity responses, whereas of the metastases-derived cell lines, LNCaP showed a high degree of resistance to UV, and PC-3 showed a high degree of resistance to etoposide. The clonogenic sensitivity of 22Rv1 to the various cell death inducers can be considered intermediate between that of PNT2C2 and LNCaP/PC-3 cells.
Host cells for functional cloning studies should be of homogeneous genetic background and, crucially, be amenable to the efficient and uniform induction of cell death. For these reasons, we attempted to clone the prostate cell lines. Clones were successfully produced from PNT2C2, 22Rv1, and PC-3 cell lines, and an initial screen revealed significant heterogeneity for each cell line in the sensitivity of clones to UV irradiation. Indeed, many prostate cell lines have been shown to comprise a heterogeneous mixture of clones differing, for example, in sensitivity to chemotherapeutic agents and androgens (Wan et al, 2003; Iguchi et al, 2007; Ware et al, 2007). This may simply reflect cell heterogeneity in the original tissue from which the cell line was derived, or it could be due to the accumulation of mutations upon prolonged periods in cell culture, especially if the cell line exhibits some degree of genetic instability. In this regard, periods of nutrient exhaustion and/or toxic metabolite accumulation in cell culture will tend to select for apoptosis-resistant clones.
Further characterization of clones demonstrated at least 1 clone per cell
line with high sensitivity (mean clonogenic survival 0.02% control value)
to several cell death inducers. These comprised PNT2C2 clones 5 and 11 treated
with UV (120 J/m2), etoposide (10 µg/mL), or okadaic acid (100
nM); 22Rv1 clone 8 treated with UV (480 J/m2), etoposide (200
µg/mL), okadaic acid (200 nM), or doxorubicin (1 µg/mL); and PC-3 clones
2 and 3 treated with UV (360 J/m2), doxorubicin (0.25 µg/mL), or
okadaic acid (100 nM). Under such conditions, a false-positive rate of no more
than 1 in 5000 colonies would be anticipated in functional cloning
experiments.
Androgens play a crucial role in regulating cell death/survival in the normal prostate epithelium, whereas anti-androgen therapy is an important treatment for early-stage prostate cancer. Of the 3 cell lines cloned, only 22Rv1 cells are androgen sensitive (Sramkoski et al, 1999), and both 22Rv1 parental cells and the derived clone 8 exhibited growth sensitivity to the synthetic androgen R1881 in the present study. Furthermore, the clonogenic activity of these cells could be inhibited by prolonged treatment with the anti-androgen Casodex. However, the degree of inhibition of clonogenic growth was insufficient to merit its use as a selection agent during functional cloning experiments with 22Rv1 clone 8 as host cells. Rather, Casodex could be employed in a secondary screen to identify cell death–resistant clones of 22Rv1 clone 8 with altered antiandrogen sensitivity.
In summary, we have successfully produced clones of 3 prostate cell lines that are uniformly sensitive to cell death induction by at least 3 different stimuli and can be infected at high efficiency with a lentiviral vector. These clones are therefore suitable as host cells in functional cloning studies to identify critical regulators of cell death in prostate cells. Moreover, the cell lines that we have developed are models of distinct stages of prostate disease, including nontumorigenic cells (PNT2C2), androgen-sensitive prostate cancer cells (22Rv1), and androgen-independent, metastatic prostate cancer cells (PC-3). Future studies with these clones are likely to shed light on cell death control mechanisms in the prostate during cancer progression and the development of resistance to therapeutic agents.
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Acknowledgments |
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Footnotes |
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