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 Previous Article | Table of Contents | Next Article 
Blood, Vol. 91 No. 9 (May 1), 1998:
pp. 3347-3356
Constitutive Expression of the Promyelocytic Leukemia-Associated
Oncogene PML-RAR in TF1 Cells: Isoform-Specific and Retinoic
Acid-Dependent Effects on Growth, bcl-2 Expression, and Apoptosis
By
James L. Slack and
Min Yu
From the Division of Medicine and the Departments of Hematologic
Oncology/Bone Marrow Transplantation, Roswell Park Cancer Institute,
Buffalo, NY.
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ABSTRACT |
Two major isoforms of PML-RAR are associated with
(15;17)-positive acute promyelocytic leukemia (APL); however,
functional differences between these isoforms have been difficult to
define, and the molecular mechanism by which each isoform contributes to the pathogenesis of APL is not fully understood. To address these
issues, the `short' (S) and `long' (L) isoforms of PML-RAR were
constitutively expressed in the factor-dependent human erythroleukemia cell line, TF1. Expression of the L, but not the S, isoform inhibited growth of these cells in the presence of granulocyte-macrophage colony-stimulating factor (GM-CSF). In the absence of GM-CSF, the S
isoform partially protected against apoptosis, while the L isoform
accelerated cell death. Treatment with all-trans retinoic acid
(ATRA) inhibited cell growth and caused apoptosis only in PML-RAR-expressing cells, and these effects of ATRA were more marked in cells expressing the L isoform. ATRA treatment also led to
downregulation of bcl-2 and endogenous RAR in PML-RAR-expressing cells, but had little effect on the level of exogenously expressed PML-RAR. We conclude that (1) subtle differences exist in the biologic activities of the L and S isoforms of PML-RAR, and (2) both
isoforms are capable of transducing an ATRA-mediated signal that leads
to downregulation of bcl-2 and induction of programmed cell death.
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INTRODUCTION |
ACUTE PROMYELOCYTIC LEUKEMIA (APL)
represents a prototype among human leukemias, and indeed among human
cancers in general, in its sensitivity to the differentiating
properties of specific retinoids such as all-trans retinoic
acid (ATRA; reviewed in Fenaux et al1 and
Warrell2). The addition of ATRA to a standard chemotherapeutic regimen of daunorubicin and cytarabine effectively doubles the cure rate in APL, from less than 35% with chemotherapy alone to approximately 70% with the chemotherapy/retinoid
combination.3,4 The central molecular defect in APL is the
disruption of the receptor for retinoic acid, RAR, and its
reciprocal in-frame fusion with one of four partner genes, PML, PLZF,
NPM, or NuMA.5-8 In greater than 99% of APL cases, the
RAR fusion partner is the PML gene on chromosome 15; the resulting
PML-RAR chimeric transcript is present in malignant promyelocytes,
and the PML-RAR fusion protein accumulates to high levels in these
cells.5 In contrast, the reciprocal RAR-PML fusion
transcript is found in only 80% of APL patients.9 Recent
studies in transgenic mice have confirmed that the PML-RAR protein
is integrally involved in the pathogenesis of APL.10-12 The
two unique features of APL, ie, the dramatic response to ATRA and the
expression of a modified retinoic acid receptor (eg, PML-RAR), are
undoubtedly related, but the nature of this relationship is not yet
clear, and the precise molecular mechanisms by which ATRA causes
differentiation of APL blasts remain largely unknown. The delineation
of these mechanisms will clarify whether APL is a fortunate medical
curiosity or whether it will serve as a paradigm for the development of
effective differentiation therapies in other types of human cancers.
There are two major isoforms of PML-RAR that are found in patients
with (15;17)-positive APL.5 As shown in Fig
1, the so-called "Short" (S) isoform
results from a genomic break in intron 3 of the PML gene, while the
"Long" (L) isoform is a consequence of a breakpoint in PML intron
6; the type S isoform of PML-RAR is found in approximately 35% of
adult APL patients, while the L isoform is found in the remaining 55%
to 60%.13 In both cases, and indeed in all cases of
(15;17)-positive APL, the RAR gene is broken in a large intron
separating the A and B domains of RAR.5 A small subset
(approximately 8%) of APL patients have PML breakpoints located within
exon 6, and express a so-called V (variable) PML-RAR
isoform.14 In a recent large retrospective study, a
statistical association was noted between the PML-RAR S isoform and
high presenting white blood cell (WBC) count, high peripheral blast
plus promyelocyte count, and M3v morphology.13 Furthermore,
we have reported a statistical relationship between the S isoform and
secondary cytogenetic abnormalities in APL.15 Despite these
clinical differences between type S and L APL patients, isoform type
per se does not appear to be a significant prognostic factor for
prediction of long-term disease-free survival.13,16 Nevertheless, the statistical associations noted above suggest that
there are differences in the biology of APL blasts expressing either
the type S or type L PML-RAR isoform, and further insight into the
basic pathophysiology of APL could be obtained by definition of such
differences in vitro.
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| Fig 1.
(A) Schematic diagram of the 5 region of the PML gene
showing the two major breakpoint regions (designated by vertical
arrows) involved in formation of the chimeric PML-RAR fusion gene.
PML exons (rectangles) are numbered. (B) Schematic drawing of the PML-RAR `L' and `S' isoforms, created by fusion of the RAR B through F domains with PML exon 6 or PML exon 3, respectively. Motifs
in the PML gene common to each isoform include a Ring finger (Ring)
followed by two B-boxes (B1 and B2) and a coiled-coil domain consisting
of four distinct coils. A putative nuclear localization signal (NLS)
and a serine/proline-rich region (S/P) are located in PML exon 6 and
are not present in the PML-RAR S isoform.
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The central paradox in APL is the apparent dual function of the
PML-RAR fusion protein. In the absence of ATRA, it can inhibit differentiation of myeloid cells17-20 and, either alone or
in concert with other genetic abnormalities, cause APL; however, in the
presence of ATRA, PML-RAR actually appears to facilitate myeloid
differentiation,17 and thus seemingly contributes to the
cure of the disease it is believed to cause. Two recent reports suggest
a possible explanation of this paradox. Both Yoshida et
al21 and Raelson et al22 have shown that the
PML-RAR protein is rapidly and specifically degraded in response to
pharmacological concentrations of ATRA. The loss of PML-RAR protein
presumably allows an intrinsic differentiation program to proceed, a
program which may or may not be dependent on ATRA. Despite its
attractiveness, this model does not fully explain the apparent
hypersensitivity of PML-RAR-expressing cells to ATRA. For example,
only blasts from PML-RAR-positive APL patients are uniformly ATRA
responsive23: blasts from APL patients with t(11;17) do not
appear to respond to ATRA, despite a block at the promyelocyte stage of
differentiation, and no other subtype of AML is as consistently
sensitive to ATRA as is APL. It is not known if the failure of other
subtypes of AML to respond to ATRA represents their `lack' of
PML-RAR, or whether their ATRA resistance simply reflects different
inherent mechanisms (or stages) of differentiation arrest. If, in the
presence of ATRA, PML-RAR is truly a pro-differentiative or
pro-apoptotic molecule, then its over-expression in hematopoietic cells
should promote differentiation and/or apoptosis in an
ATRA-dependent manner.
In the current study, our goals were the following: (1) to define
biologic differences, if any, between cells expressing the type S and
type L PML-RAR isoforms in vitro; and (2) to develop an in vitro
model to evaluate the function of the PML-RAR molecule, in
particular its ability to transduce an ATRA-mediated differentiative or
apoptotic signal. We show that TF1 erythroleukemia cells constitutively expressing PML-RAR, but not isogenic control cells, are
hypersensitive to ATRA-induced apoptosis, an effect that is
particularly pronounced in cells expressing the type L PML-RAR
isoform. Further, we show that ATRA downregulates bcl-2 expression in a
PML-RAR-dependent manner. These results suggest that gene
expression programs important in the regulation of differentiation
and/or apoptosis in APL cells are directly regulated by ATRA
through the PML-RAR protein.
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MATERIALS AND METHODS |
Chemicals and reagents.
Except as noted, chemicals and other routine laboratory reagents were
purchased from Sigma (St Louis, MO). ATRA (tretinoin) was dissolved in
absolute ethanol at 10 3 mol/L and stored at  20°C.
Fresh solutions were made weekly. Camptothecin was dissolved in
dimethyl sulfoxide (DMSO) at 10 mmol/L and stored at
20°C. Final concentrations used were 106 mol/L
(ATRA) and 0.15 µmol/L (camptothecin).
Plasmid construction.
The cDNAs for the S and L isoforms of PML-RAR24 were
kind gifts from P. Chambon (Université Louis Pasteur). Using
standard recombinant DNA procedures, each cDNA was subcloned into the
mammalian expression vector pCI-neo (Promega Corp, Madison, WI), which
contains the cytomegalovirus immediate-early promoter/enhancer, as well as the neomycin phosphotransferase gene. The integrity of the plasmids
was confirmed by extensive restriction enzyme digestion and ultimately
by visualization of correctly sized proteins in cell lines.
Cell culture and transfection.
HL-60, KG1, and NB425 cells were cultured at 37°C in a
humidified 5% CO2 incubator in RPMI 1640 supplemented with
10% fetal calf serum, 2 mmol/L L-glutamine, and
penicillin/streptomycin. TF1 cells26 were cultured in the
same medium plus 5 ng/mL granulocyte-macrophage colony-stimulating
factor (GM-CSF) (complete medium). For transfections, 10 × 106 TF1 cells were washed, resuspended at 2 × 107 cells/mL in serum-free RPMI 1640, and incubated with 25 µg of linearized plasmid DNA at room temperature for 5 minutes. Cells were transfected by electroporation (300 V, 950 µF), incubated on ice
for 15 minutes, and transferred to prewarmed complete medium. After 48 hours, geneticin (Life Technologies, Gaithersburg, MD) was added to a
final concentration of 800 µg/mL (active). At this concentration,
essentially 100% of mock-transfected TF1 cells died within 3 weeks.
Populations were screened for expression of PML-RAR using reverse
transcriptase-polymerase chain reaction (RT-PCR) as
described,15 and were then subjected to limiting dilution
cloning. The clonal lines were also screened for PML-RAR mRNA
expression using RT-PCR and subsequently for PML-RAR protein expression by Western blot.
Cell proliferation assays.
Cell growth and viability were quantitated using a colorimetric assay
that detects cleavage of the tetrazolium salt WST-1 to formazan by
mitochondrial dehydrogenases present in viable cells (Boehringer
Mannheim, Indianapolis, IN). The amount of water-soluble formazan dye
correlates directly with the number of metabolically active cells and
was quantified by measuring absorbance at 450 nm using a Dynatech 5000 microplate reader (Dynatech Laboratories, Chantilly, VA). In some
experiments, proliferation was assessed using standard 3H
thymidine assays; excellent correlation was observed between the
colorimetric and 3H thymidine assays.
Western blotting.
Total cell extract was prepared by lysing washed, precooled cells in
ice-cold RIPA buffer (1× phosphate-buffered saline [PBS], 1%
NP-40, 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulfate) containing the following protease inhibitors added immediately before
cell lysis: pefabloc (1.0 mg/mL); EDTA (1 mmol/L); leupeptin (25 µg/mL); pepstatin (10 µg/mL); and aprotinin (2.5 µg/mL). The protease inhibitors were purchased from Boehringer Mannheim. The lysate
was incubated on ice for 20 minutes, then centifuged at 15,000g
for 30 minutes at 4°C. The supernatant (total cell lysate) was
recovered and stored at 20°C. Protein concentration was determined using the BioRad protein assay kit (BioRad, Hercules, CA) according to
the manufacturer's instructions. Equal amounts of total cell protein
were electrophoresed using standard procedures and transferred to
Hybond-ECL nitrocellulose (Amersham, Arlington Heights, IL). Membranes
were blocked in tris-buffered saline containing 5% dried milk and
0.1% Tween 20, then incubated with primary and (horseradish peroxidase-conjugated) secondary antibodies. Detection of
immunoreactive bands was performed using enhanced chemiluminescence and
exposure to Hyperfilm-ECL (Amersham).
Antibodies.
The following antibodies were purchased from Santa Cruz Biotechnology
(Santa Cruz, CA) and used at dilutions suggested by the supplier or at
empiric dilutions determined by experimentation: rabbit polyclonal
anti-RAR (cat. no. sc-551); mouse monoclonal anti-Bcl-2 (cat. no.
sc-509); rabbit polyclonal anti-MEK2 (cat. no. sc-524); goat polyclonal
anti-actin (cat. no. sc-1616); rabbit polyclonal anti-Bax (cat. no.
sc-930); and goat polyclonal anti-Bcl-xS/L (cat. no.
sc-634-G).
Apoptosis assays.
Two different flow cytometric assays were used to detect and quantitate
apoptosis. Details of the TUNEL assay (see Fig 4) have been
published.27 Briefly, cells were resuspended at 2 × 106 cells/mL and fixed for at least 18 hours at 4°C in
PBS, pH 7.1, containing 5% ultrapure formaldehyde (Polysciences Inc,
Warrington, PA). For end-labeling 3 DNA strand breaks, cells
(1 × 106) were washed in 1× TdT buffer (200 mmol/L
sodium cacodylate, 25 mmol/L Tris-HCl, 250 µg/mL bovine serum albumin
fraction V) and subsequently incubated for 30 minutes at 37°C in 1×
TdT buffer plus 2.5 mmol/L CoCl2, 0.0125 nmol
fluorescein-12-dUTP, and 12.5 U terminal deoxynucleotidyl transferase
(TdT; all purchased from Boehringer Mannheim). After washing, samples
were analyzed on a FACScan flow cytometer (Becton Dickinson, San Jose,
CA) equipped with a 488-nm argon excitation laser. Fluorescein-12-UTP
was detected using a 530/30-nm bandpass filter (FL1 channel). In the
second assay (see Fig 5), apoptosis was quantitated by measuring
membrane redistribution of phosphatidylserine, as detected by binding
to fluorescein isothiocyanate (FITC)-conjugated Annexin-V
antibody.28 Briefly, 1 × 106 cells were
washed with PBS and incubated for 10 minutes in 200 µL binding buffer
containing FITC-conjugated Annexin-V antibody (10 µL of 20 µg/mL
stock) and propidium iodide (10 µL of 50 µg/mL stock). Reagents for
use in this assay were purchased as a kit (ApoAlert Annexin V apoptosis
kit; Clontech Laboratories, Palo Alto, CA). Cells were analyzed by flow
cytometry at 488 nm as discussed above.
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| Fig 4.
(A) Growth and viability in the absence of GM-CSF. TF1
parental cells, TF1-neo clones (n = 4), TF1-S clones (n = 9),
and TF1-L clones (n = 8) were washed extensively and replated at
2.5 × 105 cells/mL in medium lacking GM-CSF. Daily
determinations of viable cell number were performed as in Fig 3. The
final plotted value is the average for all clones tested. The day 0 value was set at 100%, and represents a colorimetric determination
performed 1 hour after plating. (B) Apoptosis determination using a
TUNEL assay (see Materials and Methods) after 72 hours of culture in the presence (left panels) or absence (right panels) of GM-CSF. Top
panels: TF1-neo; middle panels: TF1-S; bottom panels: TF1-L. Viable
cells are represented in the left lower quadrant of each panel, early
(live) apoptotic cells are in the right lower quadrant, and late-stage
apoptotic (dead) cells are displayed in the right upper quadrant. The
number in each quadrant represents the percent of events in that
quadrant.
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| Fig 5.
Apoptosis in response to ATRA or camptothecin. TF1-neo
cells (top panels) and TF1 cells expressing the type L (middle panels) or S (lower panels) PML-RAR isoforms were plated at 1 × 105 cells/mL in complete medium (+GM-CSF) and cultured
with vehicle alone (left panels), ATRA (106 mol/L, 96 hours; middle panels), or camptothecin (0.15 mol/L, 16 hours; right
panels). Apoptosis was evaluated by flow cytometry after staining cells
with an FITC-conjugated Annexin-V antibody (1 µ/mL) and propidium
iodide (2.5 µg/mL) as discussed in Materials and Methods. Black dots,
viable cells; blue dots, early stage (live) apoptotic cells; red dots,
late stage (dead) apoptotic cells. The percentages of events in each
quadrant are shown.
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Cell-cycle analysis.
Cells were stained with propidium iodide (50 µg/mL) in the presence
of RNase A (1 mg/mL) and analyzed by flow cytometry as described.29
Statistical analysis.
The Student's t-test was used to test for statistically
significant differences between groups.
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RESULTS |
Generation of cell lines expressing the short (S) and long (L) isoforms
of PML-RAR.
Complementary DNAs encoding the type S and L isoforms of PML-RAR
(see Fig 1) were subcloned into the mammalian expression vector pCIneo.
These plasmids, as well as the pCIneo vector itself, were transfected
into the human factor-dependent erythroleukemia cell line,
TF126; single-cell clones (confirmed by Southern blotting)
were derived by limiting dilution in the presence of both GM-CSF (5 ng/mL) and geneticin (800 µg/mL). These lines are hereafter referred to as TF1-S (clones expressing the PML-RAR S isoform), TF1-L (clones
expressing the PML-RAR L isoform), and TF1-neo (TF1 clones containing vector only).
Nine randomly chosen clones expressing either the PML-RAR S or L
isoform were expanded and characterized. In addition, 6 TF1-neo clones
were derived and used as controls. Expression of exogenous PML-RAR
and endogenous RAR proteins was demonstrated using a polyclonal
RAR antibody (Fig 2), which detects both
endogenous RAR (E; approximately 55 kD), as well as the hybrid
PML-RAR proteins (Fig 2: L-isoform, approximately 110 kD; S isoform,
approximately 95 kD). The level of expression of endogenous RAR was
similar in all TF1-neo (data not shown), TF1-S, and TF1-L clones (Fig 2). The PML-RAR S isoform was expressed at a higher level than the L
isoform in 8 of the 9 clones (Fig 2). This was not a technical artifact, because similar results were seen with different
electrophoretic and transfer conditions (data not shown); stripping and
reprobing the membranes with two different control antibodies confirmed equivalence of loading and transfer (Fig 2).
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| Fig 2.
Expression of PML-RAR in TF1-S and TF1-L clones. Equal
amounts of total cellular protein from nine independent TF1-S and TF1-L
clones were immunoblotted with an antibody to human RAR, as detailed
in Materials and Methods. Molecular weights are indicated to the left
of the gel in kilodaltons (×103). The polyclonal
RAR antibody detects both endogenous RAR (E, at approximately 55 kD), as well as the fusion PML-RAR proteins (S isoform,
approximately 95 kD; L isoform, approximately 110 kD). The blots were
stripped and reprobed with either MEK-2 or actin antibodies as a
control for differences in loading and transfer.
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Expression of PML-RAR inhibits growth of TF1 cells
and confers responsiveness to the growth inhibitory effects of ATRA.
Growth of TF1-neo, TF1-S, and TF1-L clones was assessed using a
colorimetric cell proliferation assay; at each time point, the values
for independent clones were averaged, and composite growth curves were
constructed as shown in Fig 3A. The growth of TF1-neo clones was
indistinguishable from that of parental TF1 cells (data not shown), and
was also similar to that of the TF1-S clones (Fig
3A). After 96 hours of culture, there were
significantly fewer TF1-L cells compared with either TF1-S or TF1-neo
cells (Fig 3A; P = .003, comparing mean optical density
[OD] value of L clones to neo controls at 96 hours. The
difference between TF1-S and TF1-L viable cell number at 96 hours was
also statistically significant (Fig 3A; P = .004). Thus, in
this system, PML-RAR functioned as an isoform-specific inhibitor of
cell growth.
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| Fig 3.
(A) Growth curves of TF1-neo, TF1-S, and TF1-L clones.
Cells were plated at 7 × 104 cells/mL in 96-well plates
in complete medium (GM-CSF at 5 ng/mL). Viable cell number was
determined daily using a colorimetric assay (see Materials and Methods)
with the day 0 value set at 100%. The results are the averages
(±SEM) of values for 3 TF1-neo, 6 TF1-S, and 5 TF1-L clones. (B)
Growth inhibition in response to ATRA. Cells were plated in 96-well
plates at 5,000 cells/well in 100 µL of complete medium, and cultured
for 96 hours in the presence of ATRA (106 mol/L) or an
equal volume of vehicle control (ethanol, EtOH). Viable cell number at
96 hours was quantitated as in (A). The final plotted value is the
average (±SEM) for 4 TF1-neo, 9 TF1-S, and 8 TF1-L clones.
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ATRA treatment (106 mol/L for 96 hours) of parental TF1
cells (data not shown) or TF1-neo clones caused minimal inhibition of
cell growth (OD 450, 84% ± 15% of control; P = .13; Fig
3B). This was not due to lack of expression of RAR, because
endogenous RAR was abundantly expressed in these cells (eg, see Fig
7; and data not shown). In cells expressing the PML-RAR S isoform,
viable cell number was decreased by ATRA to 47% ± 24% of control
(Fig 3B; P = .015 compared with TF1-neo cells), while in
TF1-L cell lines, viable cell number after ATRA treatment was reduced
to 27% ± 8% of control (Fig 3B; P < .001 compared with
TF1-neo cells). The difference in response to ATRA between clones
expressing the type L and type S PML-RAR isoforms was significant
(P = .04). The effect of ATRA on cell-cycle status was also
examined in representative TF1-neo, TF1-S, and TF1-L clones. There was
a small increase in the percentage of cells in G2 in the TF1-S and
TF1-L clones treated with ATRA for 96 hours (from 4.6% to 7.1% and
5.7% to 9.7%, respectively), which was accompanied by a slight
decrease in the percentage of cells in S-phase in each case (data not
shown). There were no significant changes in cell-cycle parameters in
TF1-neo cells treated with ATRA in a similar fashion (data not shown).
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| Fig 7.
Regulation of RAR and PML-RAR by ATRA. (A) The
indicated cell types were treated with vehicle control (ethanol
[]) or 106 mol/L ATRA (+) for 96 hours; total
cellular protein was obtained, and equal amounts were electrophoresed
and immunoblotted successively with RAR (top panel) and actin
(bottom panel) antibodies. Molecular weight markers are indicated to
the left of the gel (in kilodaltons × 103). The
migration of the PML-RAR L isoform (L), PML-RAR S isoform (S),
and endogenous RAR (E) are indicated. (B) Quantitation of the
results shown in (A) including data from additional independent experiments. The control level of expression of either endogenous RAR or PML-RAR was set at 100% in each case, and the relative level of expression of these proteins in response to ATRA was quantitated by densitometric analysis of Western blots such as those
shown in (A). Values were corrected for differences in loading and
transfer by incorporating the actin control in the calculation.
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The PML-RAR S isoform, but not the L isoform,
partially protects TF1 cells from apoptosis in response to growth
factor withdrawal.
To evaluate the effect of exogenously expressed PML-RAR on cell
number and viability in response to growth factor withdrawal, TF1-neo,
TF1-S, and TF1-L clones were washed and cultured for 96 hours in the
absence of GM-CSF. As shown in Fig 4A, most
TF1-S clones remained relatively viable after 4 days of growth factor deprivation (OD 450, 56% ± 35% of day 0 value; range, 8.5% to 124%), whereas TF1-L clones uniformly died (OD 450, 4% ± 3% of day 0 value; range, 1.8% to 11%; P < .001 compared with S
clones). TF1-neo clones showed an intermediate pattern of cell
viability (OD 450, 15% ± 7% of day 0 value; range, 8.6% to
25%), similar to that of parental TF1 cells (9.1%). The difference in
viable cell number between TF1-L and TF1-neo clones was statistically significant (P < .001). To confirm that the loss of
viability was due to apoptosis, cells were cultured for 72 hours in the presence or absence of GM-CSF and apoptosis was assessed using a flow
cytometric assay based on incorporation of fluorescently labeled dUTP
into DNA strand breaks.27 After 72 hours of growth factor
deprivation, virtually all (95.4%) TF1-L cells had begun to undergo,
or had completed, apoptosis, compared with only 46% of TF1-S cells
(Fig 4B; compare middle and lower panels). The corresponding
percentages of live, nonapoptotic cells after 72 hours of culture in
the absence of GM-CSF were 4.6% for TF1-L and 53.6% for TF1-S (Fig
4B). For TF1-neo cells, the percentage of apoptotic cells after 72 hours of growth factor deprivation was 67.6% (Fig 4B, top right
panel).
ATRA treatment of PML-RAR-expressing, but not control, cells
causes apoptosis.
To evaluate the effects of ATRA on apoptosis in PML-RAR-expressing
and control cells, TF1-neo, TF1-L, and TF1-S clones were treated with
vehicle control (ethanol) or ATRA (106 mol/L) for 96 hours. As a positive control, cells were also treated with the
topoisomerase I inhibitor camptothecin (0.15 mol/L, 16 hours), a known
inducer of apoptosis in hematopoietic cell lines.30 Apoptosis was evaluated by flow cytometry after staining cells with an
FITC-labeled antibody to Annexin-V.28 Under control conditions (Fig 5, left panels) there was a
small but significant degree of spontaneous apoptosis in the TF1-L
cells (20.7% apoptotic) and, to a lesser extent, in the TF1-S cells
(11.3% apoptotic, v 6% spontaneous apoptosis in the TF1-neo
cells). On a relative basis, the TF1-neo cells were most sensitive to
camptothecin (Fig 5; percent apoptotic/dead cells increased from 6.0%
[control] to 33.4% [camptothecin-treated]). For both TF1-L and
TF1-S, the percent apoptotic/dead cells increased by approximately
twofold in response to camptothecin (Fig 5; TF1-L 20.7% to 39.6%;
TF1-S 11.3% to 20.7%). The final percentage of apoptotic cells
was similar for TF1-neo and TF1-L (33.4% and 39.6%, respectively),
but somewhat lower for the TF1-S cells (20.7%). Thus, camptothecin
caused apoptosis in all three cell types, although expression of either
PML-RAR isoform may have afforded a small degree of protection. In
contrast, ATRA caused significant apoptosis only in cells expressing
PML-RAR, and the degree of apoptosis in response to ATRA was much
more marked in TF1-L (73.4% apoptotic) than in TF1-S cells (28.8%
apoptotic; Fig 5, middle panels). Eighty-nine percent of TF1-neo cells
remained nonapoptotic after ATRA treatment, versus 24.9% of TF1-L and
69.2% of TF1-S cells. These results were qualitatively confirmed using DNA laddering assays and different TF1-neo, L, and S clones (data not
shown).
ATRA decreases bcl-2 expression in a
PML-RAR-dependent fashion.
bcl-2 is known to protect a variety of cell types, including myeloid
cells, from apoptosis.31 To explore the relationship, if
any, between bcl-2 expression and PML-RAR, bcl-2 levels were measured by immunoblotting in multiple TF1-neo, TF1-S, and TF1-L clones. The basal level of bcl-2 expression in TF1-neo clones was
essentially identical to that seen in parental TF1 cells grown under
the same conditions (data not shown). Basal bcl-2 levels were
approximately twofold higher in TF1-S clones compared with neo controls
(Fig 6A and B). In contrast, basal bcl-2
expression in TF1-L clones was significantly lower than that seen in
TF1-neo cells (average, 34% of the level in TF1-neo clones; Fig 6A and B). When TF1-neo clones were treated with ATRA (106
mol/L, 96 hours), there was no significant change or, in some experiments, a slight increase in the level of bcl-2 protein (Fig 6).
However, identical ATRA treatment of TF1-S and TF1-L clones resulted in
a significant decline in bcl-2 protein levels. In TF1-L cells, there
was a sixfold decrease in bcl-2 protein levels in response to ATRA,
whereas in TF1-S cells, the decline in bcl-2 in response to ATRA was
approximately threefold (from a higher baseline; see Fig 6B and the
representative Western blot in Fig 6A). The regulation of bcl-2 by ATRA
in these cells occurred at a pretranslational level, as documented by
Northern blot analysis (data not shown). ATRA treatment had no effect
on expression of bcl-x or bax in TF1-neo, TF1-S, or TF1-L clones (Fig
6A).
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| Fig 6.
Expression of bcl-2, bax, and bcl-x in response to ATRA
in TF1-neo, TF1-L, and TF1-S cells. (A) Cells were treated with vehicle alone (ethanol []) or ATRA (106 mol/L; [+]) for
96 hours; equal amounts of total cellular protein were electrophoresed
and immunoblotted successively with antibodies to bcl-2, bcl-x, or bax.
The blots were then reprobed with an actin antibody to control for
variations in loading and transfer. A representative Western blot is
shown. (B) Quantitation of data from four independent experiments with
two different TF1-neo, TF1-S, and TF1-L clones. Shown is the average
level of bcl-2 expression (±SEM) under control conditions (EtOH;  )
or after 96 hours of ATRA treatment (ATRA;  ), compared in all cases
to the bcl-2 expression level in TF1-neo clones under control
conditions (value arbitrarily set at 100%). Values were corrected for
differences in loading and transfer by incorporating the actin control
in the calculation.
|
|
Regulation of PML-RAR and endogenous
RAR by ATRA.
It has been suggested22 that ATRA contributes to the cure
of APL by selectively downregulating the expression of PML-RAR, with
little or no effect on endogenous RAR. To address this issue in the
current model system, TF1 parental cells, TF1-neo clones, and TF1-S and
-L clones were treated with ATRA (106 mol/L for 96 hours) and the level of expression of both endogenous RAR and
transfected PML-RAR was assayed by Western blot (Fig 7). Control experiments were performed with
the cell lines KG1 (a non-APL human myeloid cell line) and NB4 (a human
promyelocytic cell line which expresses the PML-RAR L isoform). The
Western blots were quantitated by densitometry, and values for relative PML-RAR and/or endogenous RAR expression were normalized
for actin expression. In KG1 (data not shown), TF1 parental (data not
shown), and TF1-neo cells, treatment with ATRA had a minimal effect on
the level of expression of endogenous RAR (Fig 7A and B). In
contrast, endogenous RAR was consistently downregulated in all
PML-RAR-expressing cell lines (TF1-S, TF1-L, NB4) in response to
ATRA (Fig 7A and B). The extent of this decrease in endogenous RAR
expression was similar in all three PML-RAR-expressing cell types
(TF1-S clones: 17% of control, range 7% to 31% in three experiments;
TF1-L clones: 13% of control, range 3.5% to 22% in four experiments;
NB4 cells: 10% of control, range 2.9% to 17% in two experiments). In
NB4 cells, the endogenous type L PML-RAR protein was also
downregulated by ATRA (average 13% of control level in two
experiments; Fig 7A and B), in agreement with Raelson et
al.22 In TF1-L clones, there was some variability in the level of expression of exogenous PML-RAR in response to ATRA; however, averaging four experiments in which quantitative values were
obtained after controlling for actin expression, the relative level of
PML-RAR (L) expression after ATRA treatment was 63% of control
(range, 36% to 117%; see Fig 7A and B). The relative level of
exogenous PML-RAR (S) expression after ATRA treatment was 88% of
control (range, 76% to 103%; Fig 7A and B).
| |
DISCUSSION |
One of the major aims of this study was to define biologic differences,
if any, between the type S and type L PML-RAR isoforms in vitro. To
achieve this goal, the two PML-RAR isoforms were constitutively
expressed in TF1 cells, a human erythroleukemia cell line that is
resistant to the differentiative and antiproliferative effects of
ATRA.26 It is noteworthy that the type S PML-RAR isoform
was expressed at significantly higher levels that the type L isoform in
8 of the 9 clones examined (Fig 2). This finding is likely due to
differences in the physiologic properties of the two PML-RAR
molecules in TF1 cells because our data show that the PML-RAR type L
isoform, even when expressed at a lower level than the S isoform,
inhibited growth and promoted apoptosis to a much greater degree. Thus,
it is probable, although not experimentally proven, that TF1 cells can
express only a finite amount of the PML-RAR L isoform and remain
viable and clonable. This interpretation is consistent with the
difficulty other investigators have encountered in establishing cell
lines which constitutively express PML-RAR, and with our own
difficulty in constitutively expressing the PML-RAR L isoform in
other hematopoietic cell lines, including HL-60.
There are significant structural differences between the PML-RAR S
and L isoforms which may account for the observed differences in
biological activity reported in the present study. The two major
structural motifs missing in the S isoform are a serine-proline rich
region and a putative nuclear localization signal (NLS), both encoded
by PML exon 6 (see Fig 1). The importance of these and other structural
elements to the subcellular location and function of the PML protein
has been examined in 3T3 cells.32 In that system, a mutant
PML protein which lacked the NLS was localized to the cytoplasm, did
not participate in nuclear body formation, and was functionally
inactive.32 In contrast, a mutant PML protein that lacked
the serine-proline rich region was found in the nucleus in
normal-appearing nuclear bodies, and was fully functional.32 Although some nuclear localization must be
provided by the NLS in the RAR moiety, the PML-RAR S isoform may
nevertheless be less able to enter the nucleus than the L isoform, and
therefore less able to transmit a retinoid-mediated signal; this
hypothesis is supported by experimental immunofluorescence
data,24 and by data presented in the present report (Figs
3B and 5) showing that the S isoform, despite being expressed at a
higher level than the L isoform, was less effective at transducing an
ATRA-mediated growth inhibitory/apoptotic signal. Despite the
differences in structure and in vitro biologic activity between the L
and S PML-RAR isoforms, both molecules readily cause APL in humans.
Furthermore, when cultured in vitro, blasts from patients with type S
and type L APL were reported to differentiate to a similar degree in
response to ATRA.33 However, it is interesting to note that
there are several clinical differences between APL patients who express either the S or L isoforms. For example, type S APL patients have a
higher incidence of secondary chromosomal abnormalities,15 and also present with higher median WBC and peripheral blast
counts.13 However, after correcting for presenting WBC
count, type S and type L APL patients have essentially identical
clinical outcomes.13,16 In summary, the combined clinical
and in vitro data suggest that there are subtle biologic differences
between the L and S isoforms of PML-RAR, but that these differences
do not affect clinical outcome in patients who receive aggressive,
modern treatment with ATRA/anthracycline combination therapy.
In the absence of ATRA (or other retinoids), PML-RAR has been
reported to inhibit myeloid cell differentiation17-20,34
and to suppress or delay apoptosis.17,35,36 These effects
have generally been relieved by the addition of ATRA, and PML-RAR has been reported to specifically enhance retinoic-acid induced differentiation of U937 cells.17 The growth inhibitory and
pro-apoptotic properties of the PML-RAR L isoform reported here,
even in the absence of ATRA, are somewhat difficult to reconcile with a
previous study,35 and with the presumed oncogenic function
of PML-RAR, but are consistent with another previous report which
examined PML-RAR expression in lymphoid cell lines.37 In
contrast to data reported by Rogaia et al,35 we observed no
protection from growth factor withdrawal-induced apoptosis in TF1 cells
expressing the PML-RAR L isoform. However, cells expressing the S
isoform were partially protected from apoptosis induced by growth
factor withdrawal, and underwent less apoptosis (compared with TF1-neo or TF1-L cells) in response to camptothecin; these results confirm that
PML-RAR may have anti-apoptotic activity, but in the present system
this activity is seen primarily with the S isoform. The data reported
here with the S isoform are consistent with those previously reported
by Fu et al,36 who also observed protection from apoptosis
after expression of the PML-RAR S isoform in TF1 cells. Although it
is possible that the difference between isoforms observed in the
present study is due to the consistently higher expression levels of
the S isoform in the TF1 clones, this seems unlikely, because the
presence of the PML-RAR L isoform actually accelerated apoptosis in
response to growth factor withdrawal (Fig 4), and clearly led to
decreased cell growth and viability even under ideal growth conditions
(Fig 3). Thus, it seems unlikely that higher levels of expression of
the L isoform would confer a growth advantage to these cells. The
reasons for the differences between the results reported here and those
of Rogaia et al35 are therefore unknown, but may be due to
differences in experimental conditions (eg, different expression
plasmids and potentially different methods to culture cells and isolate
clonal cell lines).
In the presence of pharmacologic doses of ATRA (106
mol/L), the type L (and to a lesser extent type S) PML-RAR protein
mediated rapid and extensive programmed cell death (Fig 5), which was
also accompanied by signs of erythroid differentiation (manuscript in
preparation). Essentially no apoptosis was observed in
control TF1-neo cells in response to ATRA. It should be emphasized that the degree of apoptosis of TF1-neo and TF1-L clones in response to
camptothecin, a topoisomerase I inhibitor, was similar, if not somewhat
higher for the TF1-neo cells (Fig 5). This result shows that the
apoptosis seen in the TF1-L clones in response to ATRA is not due to a
generalized increase in susceptibility to apoptosis, but rather is
specific for the PML-RAR and RAR ligand ATRA. The combined
results indicate that PML-RAR, particularly the L isoform,
sensitizes cells to ATRA-induced, but not camptothecin-induced, apoptosis; we hypothesize that ATRA binds to the RAR moiety of PML-RAR and modulates the transcriptional activity of this protein, with activation (or repression) of as yet uncharacterized genes. The
apoptotic effect of ATRA is not mediated by the normal, endogenous RAR, because parental TF1 cells and TF1-neo cells express RAR at
high levels and yet do not undergo significant apoptosis in response to
ATRA.
ATRA treatment of PML-RAR-expressing, but not control, TF1 cells
caused a significant decrease in expression of bcl-2, but not bax or
bcl-x (Fig 6). The effects of ATRA on bcl-2 levels and apoptosis in
cells expressing PML-RAR and other myeloid cell types have been
investigated by others.38-40 Calabresse et al38 reported that ATRA treatment of fresh, cultured APL blasts resulted in
a rapid decrease in bcl-2 expression, whereas no modulation of bcl-2
was observed in response to ATRA in non-APL cells; however, the fresh
APL cells did not undergo apoptosis in response to ATRA alone.38 Treatment of NB4 cells (a patient-derived
PML-RAR-positive APL cell line) with ATRA also leads to a
significant decrease in bcl-2 expression39 (and our
unpublished observations, November 1996). Although bcl-2 can be
downregulated by retinoids in myeloid cells that do not express
PML-RAR,40,41 the data presented by Calabresse et
al38 and in the current report strongly suggest that
PML-RAR facilitates bcl-2 downregulation by ATRA and suggests that
bcl-2 is a direct molecular target of retinoids in PML-RAR-positive cells. Because bcl-2 can protect cells from chemotherapy-induced apoptosis,42 it is possible, as suggested in other
systems,40,43 that the decrease in bcl-2 levels in response
to ATRA in APL blasts and other PML-RAR-expressing cells sensitizes
these cells to the subsequent apoptotic effects of chemotherapy. Thus,
the ability of PML-RAR to facilitate bcl-2 downregulation in
response to ATRA, with a consequent increased susceptibility to
chemotherapy-induced apoptotic cell death, may explain why ATRA and
other RAR agonist retinoids44,45 are effective clinical
agents in PML-RAR-positive APL, but not in other
(PML-RAR-negative) AML subtypes. If downregulation of bcl-2
expression is critical to the effectiveness of chemotherapeutic and/or differentiative agents in AML and APL, then alternative methods of bcl-2 inhibition, such as the use of antisense
oligodeoxynucleotides,43,46 may offer alternative
strategies of treatment in AML patients or APL patients who are
resistant to ATRA.
The effectiveness of ATRA in APL depends on the presence of the
PML-RAR protein.1,4 One model to explain the success of
ATRA in APL is suggested by recent studies which demonstrate that the
PML-RAR protein is significantly and rapidly downregulated by ATRA,
apparently at a posttranslational level.21,22 Because PML-RAR can inhibit myeloid differentiation,17-20 its
downregulation by ATRA could allow normal differentiation to occur and
lead to clinical complete remission, as seen both in vitro and in vivo. In this model, binding of ATRA to the RAR moiety may lead to a
change in conformation of the PML-RAR molecule that results in its
specific destruction via the proteasome pathway21; by
extension, ATRA binding to PML-RAR would not necessarily directly
activate expression of genes involved in myeloid differentiation or
apoptosis. An alternative model suggests just the opposite, ie, that
PML-RAR, on binding ATRA, becomes an active inducer of myeloid
differentiation and apoptosis, presumably by activating or inhibiting
expression of specific genes or gene programs. Results from the present
study offer support primarily for the latter model. We have confirmed,
as reported by others,22 that the PML-RAR protein is
downregulated by ATRA in NB4 cells, but our results suggest that the
endogenous RAR protein is downregulated to a similar degree (Fig 7);
thus, there was no net change in the ratio of PML-RAR to RAR
after ATRA treatment. Furthermore, we observed little (L-isoform) to
essentially no (S-isoform) downregulation of exogenously expressed
PML-RAR protein in response to ATRA. These results call into
question the generality of a posttranslational pathway for
downregulation of PML-RAR in response to ATRA. Finally, it is
notable that a consistent and striking downregulation of endogenous
RAR in response to ATRA was observed only in cells that expressed
PML-RAR. This result suggests that endogenous RAR is a molecular
target of ATRA in PML-RAR-positive cells.
Overall, our results support a model in which PML-RAR, in the
presence of ATRA, inhibits cell growth, facilitates differentiation, and induces apoptosis. The apoptotic response appears to be general, because PML-RAR can mediate ATRA-induced apoptosis in
lymphoid37 as well as erythroid cell lines (Fig 5).
Identification of target genes that are modulated by ATRA specifically
in PML-RAR-expressing cells will be an important avenue of further
study, and at least one such gene, transglutaminase II, has been
identified.47 Identification and functional
characterization of such genes may lead to a fuller understanding of
APL, and this knowledge may lead to the development of novel strategies
for the treatment of both resistant APL and other AML subtypes.
Ultimately, it is hoped that the knowledge gained in the study of APL,
particularly the mechanisms by which this disease is sensitive to
retinoids, can be translated to other AML subtypes, in which cure rates
continue to be distressingly low.
| |
FOOTNOTES |
Submitted June 6, 1997;
accepted December 15, 1997.
Supported in part by grants from the National Cancer Institute to
Roswell Park Cancer Institute.
Address reprint requests to James L. Slack, MD, Division of Medicine,
Roswell Park Cancer Institute, Elm & Carlton Sts, Buffalo, NY 14263.
The publication costs of this article were defrayed in part by page charge payment. This article must therefore be hereby marked "advertisement" is accordance with 18 U.S.C. section 1734 solely to indicate this fact.
| |
ACKNOWLEDGMENT |
We thank Earl Timm, Carlton Stewart, and Sigrid Stewart for help with
flow cytometric assays and analysis of flow cytometric data.
| |
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Abstract
Introduction
Abstract