TOXICOLOGICAL SCIENCES 74, 85–92 (2003)
DOI: 10.1093/toxsci/kfg115
Copyright © 2003 by the Society of Toxicology
7,12-Dimethylbenz[a]anthracene-Induced Bone Marrow Toxicity
Is p53-Dependent
Todd J. Page,* Scott O’Brien,* Karrie Holston,* Peter S. MacWilliams,* Colin R. Jefcoate,† ,‡ and Charles J. Czuprynski* ,‡ ,1
*Department of Pathobiological Sciences, †Department of Pharmacology, and ‡Molecular and Environmental Toxicology Center,
University of Wisconsin, Madison, Wisconsin 53706
Received December 17, 2002; accepted April 10, 2003
munotoxicity of these compounds are not completely understood. It has been demonstrated that metabolic activation by
the cytochrome P450 1B1 (CYP1B1) enzyme is required for
their immunotoxic effect (Heidel et al., 1999, 2000; Mann et
al., 1999; White et al., 1985). The role of the arylhydrocarbon
receptor (AhR) in the immunotoxic effects of PAHs is a source
of debate. When the AhR binds PAHs, it translocates to the
nucleus and induces transcription of a variety of Ah-responsive
genes, including P450 cytochromes such as CYP1B1. However, bone marrow stromal cells express a high basal level of
CYP1B1 and are able to metabolize PAHs in the absence of a
functional AhR (Heidel et al., 2000). In addition, 2,3,7,8tetrachlorodibenzo-p-dioxin, or TCDD (i.e., dioxin), binds the
AhR with a greater affinity than PAHs but is not metabolized
and has different effects on the immune system than do the
PAHs (Laupeze et al., 2002; Okey et al., 1984a,b).
PAHs are both carcinogenic and immunotoxic, although the
doses required for these two effects may differ (White et al.,
1985). The mechanisms of carcinogenicity and the mutagenic
properties of PAHs have been studied in greater detail than
their effects on the immune system. As is the case for their
immunotoxic effects, PAHs must be metabolically activated by
the cytochrome P450 family of monooxygenases to be carcinogenic (Badawi et al., 1996; Cavalieri and Rogan, 1992).
These activated metabolites then form adducts with DNA and
cause mutations. Mutations in key regulatory genes, in addition
to changes in gene expression mediated by binding of the AhR,
are thought to be the major mechanisms responsible for the
carcinogenic effects of PAHs (White et al., 1994). The fact that
these two disparate effects of PAHs are interrelated may provide some clues as to the mechanism of immunotoxicity for
these compounds.
The p53 tumor-suppressor protein is responsible for regulating cellular responses to DNA damage and progression
through the cell cycle (Hickman et al., 2002; Sharpless and
DePinho, 2002). The importance of p53 in maintaining a
normal pattern of cellular proliferation is illustrated by p53
being one of the most frequently mutated genes found in tumor
biopsies and transformed cell lines (Gupta et al., 1993). In
addition, p53 plays a crucial role as a “gatekeeper” of genomic
Polycyclic aromatic hydrocarbons (PAHs) are known immunotoxins and carcinogens. Our laboratory and others have demonstrated that metabolism of these compounds by CYP1B1 is required for carcinogenicity and immunotoxicity to occur.
Previously, our laboratory reported significantly decreased bone
marrow cellularity in mice following 7,12-dimethlybenz[a]anthracene (DMBA) administration. In addition, we have observed that
DMBA causes apoptosis via activation of both caspase-8 and -9 in
pre-B cells co-cultured with bone marrow stromal cells in vitro. In
this study, we investigated the importance of the p53 protein in the
bone marrow response to DMBA. Through the use of p53 gene
knockout mice, we demonstrated that the effect of DMBA on bone
marrow cellularity is p53-dependent. In addition, apoptosis of
primary cultures of progenitor B cells cultured with bone marrow
stromal cells and DMBA is also p53-dependent. The results of this
study provide evidence for the importance of p53 in the signaling
pathways by which PAHs cause immunotoxicity.
Key Words: PAH; DMBA; apoptosis; B cell; bone marrow; p53.
Polycyclic aromatic hydrocarbons (PAHs) are ubiquitous
environmental contaminants, generated by the incomplete
combustion of organic materials, that have been demonstrated
to be both carcinogenic and immunotoxic (Collins et al., 1998).
Humans are exposed to these compounds from a variety of
sources, including incinerators and various other industrial
processes. One of the main routes of exposure is through the
diet (Phillips, 1999). Charbroiling and smoking of meats can
generate high levels of PAHs, and PAHs can be found in fruits
and vegetables grown in areas of high industrial pollution
(Kazerouni et al., 2001). Smoking also accounts for a large
amount of human exposures (Rodgman et al., 2000).
7,12-Dimethylbenz[a]anthracene (DMBA) and benzo[a]pyrene (B[a]P) are the prototypical and best characterized
PAHs. These compounds can have significant effects on both
humoral and cell-mediated immunity (Burchiel et al., 1990;
Ward et al., 1984). The mechanisms responsible for the im1
To whom correspondence should be addressed at Department of Pathobiological Sciences, University of Wisconsin-Madison, 2015 Linden Drive,
Madison, WI 53719. Fax: 608-262-8102. E-mail: [email protected].
wisc.edu.
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integrity. If DNA has been damaged in such a way that it
cannot be effectively repaired, p53 will initiate apoptosis to
prevent the damaged cell from proliferating. Several DNAdamaging agents (e.g., cisplatin, etoposide) have been demonstrated to induce p53 and activate apoptosis in a p53-dependent
manner (Karpinich et al., 2002; Siemer et al., 1999).
The B-cell lineage constitutes the main component of the
humoral immune system. In the bone marrow, there is a specific differentiation pathway for B cells that begins with the
common lymphoid progenitor and ends with immature IgM
expressing B cells, which then leave the bone marrow to
populate the spleen and lymph nodes (Hardy and Hayakawa,
2001). B-cell development is driven by the interaction of the
progenitor B cells with bone marrow stromal cells that secrete
cytokines and express adhesion molecules that drive the differentiation of the progenitor B cells. Disruption of this interaction via exogenous toxicants could have profound effects on
B-cell lymphopoiesis and can result in the impairment of
antibody production directed against pathogens. Previously,
our laboratory demonstrated that DMBA causes apoptosis in
pre-B cells when the latter are co-cultured with bone marrow
stromal cells that metabolize DMBA via CYP1B1 (Heidel et
al., 1999, 2000; Page et al., 2002). Our hypothesis is that
DMBA disrupts hematopoiesis in the bone marrow and affects
normal B-cell development by targeting discreet cell populations along the B-cell lineage continuum and further, that this
disruption is dependent on the presence of functional p53.
The goal of this study was to determine whether the effects
of DMBA on the murine bone marrow are dependent on
activation of p53. To accomplish this goal, we utilized homozygous null p53 mice that were treated with a single bolus
injection of DMBA. We also investigated the importance of
p53 in the progenitor B cell in their response to stromal cells
and DMBA in vitro. We present evidence that the effects of
DMBA on bone marrow cells in vivo and progenitor B cells in
vitro are largely dependent on the presence of functional p53
protein.
MATERIALS AND METHODS
Animals and treatments. A breeding pair of B6.129S-2Trp53 null mice
were purchased from The Jackson Laboratory (Bar Harbor, ME). Homozygous
mice (p53 ⫺/⫺) were produced by breeding heterozygous mice (p53 ⫹/–), and the
litters were genotyped by PCR. Littermates that were genotyped as p53 ⫹/⫹
were used as wild-type (WT) control mice in our experiments. The animals
were housed at the Association for Assessment and Accreditation of Laboratory Animal Care International certified animal care facility of the University
of Wisconsin-Madison School of Veterinary Medicine. All animal use and care
was conducted in accordance with the NIH Guide for the Care and Use of
Laboratory Animals.
At 5–7 weeks of age, male or female p53 ⫺/⫺ and p53 ⫹/⫹ C57Bl/6 mice were
injected ip with either DMBA (50 mg/kg) or vehicle (olive oil). After 48 h, the
mice were euthanized; their femurs, tibias, and sternums were dissected free of
muscle tissue; and the ends were removed with a scalpel. For total cell counts,
the femurs were flushed with RPMI 1640 with 5% FBS, using a 25-gauge
needle. The cells were then sequentially passed through a 25-gauge needle and
a 22-gauge needle to generate single-cell suspensions. The cells were then
centrifuged, and the RBCs were lysed using ACK buffer (150 mM NH 4Cl, 1.0
mM KHCO 3, and 100 mM Na 2EDTA, pH 7.3). The white cells were resuspended in 5 ml of RPMI 1640 and centrifuged again. The cell pellets were then
resuspended in 5 ml of RPMI 1640, and an aliquot was removed for enumeration of viable cells using a hemacytometer and Trypan blue exclusion.
Histopathology and cytology. The sternums were removed and fixed in
ice-cold 4% phosphate-buffered paraformaldehyde, decalcified for 30 min
(Formacal-4, Decal Co., Congers, NY), embedded in paraffin, sectioned at
2-␮M thickness, and stained with H&E. Bone marrow smears were prepared
by cutting a tibia longitudinally, streaking the exposed bone marrow onto a
glass slide with a sable-hair brush, and allowing the cell smears to air dry.
These were subsequently stained with Wright’s stain and examined microscopically, as described previously (Heidel et al., 2000; Page et al., 2002). The
sternum sections and the bone marrow smears were examined microscopically
and scored for cell type by an American College of Veterinary Pathologists
board-certified pathologist (P.S.M.). For the bone marrow differential cell
counts, a total of 500 cells were counted, and the percentage of cells in each
of the following categories was enumerated: erythroid precursors, proliferating
granulocytes (myeloblast, progranulocytes, myelocytes), mature granulocytes
(metamyelocytes, bands, segmented neutrophils), and lymphoid cells. The total
cell number for each category was derived by multiplying the percentage of
each cell type present in the Wright-stained bone marrow smears of the tibia
by the total number of bone marrow cells generated after flushing the marrow
from two femurs per mouse. Abnormalities in cellular development in both the
erythrocytic and granulocytic lineages were identified and recorded.
Primary culture of bone marrow progenitor B lymphocytes. Long-term
bone marrow cultures (LTBMC) were established according to the protocol of
Witte (Whitlock et al., 1984). Briefly, cells were flushed from the mouse
femurs with sterile, ice-cold, RPMI 1640 (5% FBS) (Mediatech, Herndon,
VA), then centrifuged and resuspended in 1 ml RPMI 1640 (5% FBS). An
aliquot of cells was then diluted 1:10 with crystal violet (0.01% w/v crystal
violet, 3% acetic acid) and counted using a hemacytometer. Cell density was
adjusted to 1 ⫻ 10 6 cells/ml, and the cells were cultured in RPMI 1640 with
5% FBS at 37°C. The media were changed each week (75% fresh media in
total volume). After 2 weeks of culture, the media were supplemented with 4
ng/ml of recombinant murine IL-7 (R&D Systems, Minneapolis, MN). By 3
weeks, progenitor B cells were visible in suspension and loosely adherent to
the stromal cells.
The phenotype of the progenitor B cells from the LTBMCs was then verified
using fluorochrome-conjugated primary antibodies specific for pre-B cell surface markers and flow cytometric analysis (Thurmond and Gasiewicz, 2000).
The antibodies used were IgM (clone AF6-78), B220 (clone RA3-6B2), AA4.1
(clone AA4.1), and CD19 (clone 1D3). All antibodies were purchased from
Pharmingen (San Diego, CA). Briefly, the cells were isolated from the LTBMCs by gentle agitation, washed twice with PBS ⫹ 2% BSA, and fixed with
4% paraformaldehyde. The cells were then washed again and stained with the
appropriate antibodies. The cells were then washed twice and stored at 4°C
until FACS analysis was performed. The data were analyzed using Cellquest
Pro software (BD Biosciences, San Jose, CA). After phenotyping an aliquot of
the cells harvested from the LTBMCs, the remainder of the cells were then
used for experiments.
Propidium iodide staining. The immunophenotyped pre-B cells were removed from the LTBMCs and co-cultured with monolayers of bone marrow
stromal cells (BMS2 cell line) that were at approximately 75% confluence. The
BMS2 cell line was generously provided by Dr. Paul Kincade (Oklahoma
Medical Research Foundation, Oklahoma City, OK). The cultures were incubated with the indicated concentrations of DMBA for 24 h; pre-B cells were
then removed from the BMS2 monolayers by gentle agitation, as described
previously (Page et al., 2002). The detached cells were centrifuged for 5 min
at 1200 rpm and washed once in ice-cold PBS ⫹ 2% bovine serum albumin.
Cells were fixed in 1 ml of 80% ice-cold ethanol for 30 min at –20°C, then
centrifuged for 5 min at 1200 rpm and resuspended in phosphate-citric acid
buffer (0.192 M Na 2HPO 4, 4mM citric acid, pH 7.8) for 5 min at room
DMBA-INDUCED BONE MARROW TOXICITY
87
viability between vehicle- or DMBA-treated p53 null mice
(data not shown).
Histopathological Analysis of Murine Bone Marrow
FIG. 1. There is no change in total live bone marrow cells after DMBA
treatment of p53 null mice. p53 null and WT mice were injected ip with
DMBA (50 mg/kg). The bone marrow cells were harvested 48 h later and
enumerated using a hemacytometer. Live cells were differentiated from dead
cells through Trypan blue exclusion. The data are presented as the mean ⫾
SEM number of live cells present in the two femurs of each mouse (n ⫽ 6).
Data were analyzed using the unpaired t-test. Asterisk indicates data that are
statistically different from vehicle control (oil-treated) mice, p ⬍ 0.05.
temperature. After centrifugation, the cells were resuspended in 0.5 ml of
propidium iodide staining solution (33 ␮g/ml propidium iodide; Sigma-Aldrich), 1 mg/ml RNase A, 0.2% Triton X-100, in PBS) and analyzed in a
Becton/Dickinson FACScan flow cytometer (BD Biosciences, San Jose, CA).
Because cells undergoing DNA fragmentation and apoptosis exhibit weaker
propidium iodide fluorescence than do cells in the G o/G 1 cell cycle, a decrease
in PI fluorescence is indicative of the morphological changes consistent with
apoptosis (Yamaguchi et al., 1997).
RESULTS
Bone Marrow Cell Depletion Does Not Occur in DMBATreated p53 Null Mice
p53 null and WT mice were injected ip with 50 mg/kg
DMBA or vehicle control and euthanized after 48 h. A 48 h
exposure time was chosen because in previous experiments we
have observed significant depletion of bone marrow cellularity
at this time point (Heidel et al., 2000). The bone marrow cells
were isolated and counted, then live cells were discriminated
from dead cells by Trypan blue exclusion. There was a significant decrease in the number of viable cells recovered from the
bone marrow of DMBA-treated WT mice, as compared with
vehicle-treated mice (Fig. 1). Both the total number of cells and
the percentage of viable cells were decreased in DMBA-treated
versus vehicle control WT mice (data not shown). In contrast,
there was no difference in the numbers of bone marrow cells
recovered from DMBA- and vehicle-treated p53 null mice
(Fig. 1), nor was there any difference in bone marrow cell
Wright-stained bone marrow smears from the tibias of WT
mice exhibited abnormal large blast cells that have been characterized previously by extensive cytochemical staining as
being members of the granulocyte lineage (Fig. 2C) (Heidel et
al., 2000). These blast cells were absent from the WT control
bone marrow (Fig. 2D) and both the DMBA-treated and control p53 null mice (Figs. 2A and 2B). DMBA-treated WT mice
also exhibited aberrant multinucleated granulocytes that were
absent in the WT controls and the DMBA-treated p53 null
mice (Figs. 2E and 2F).
Bone marrow sections from the sternums of WT DMBAtreated mice exhibited a marked hypocellularity when compared with both untreated WT marrow and either treated or
untreated p53 null marrow (Fig. 3). These histopathological
observations confirm the decreased numbers of viable cells
recovered from the femurs of DMBA-treated mice (Fig. 1).
The WT DMBA-treated murine bone marrow also exhibited
dilated sinusoids with extensive erythrocyte infiltration, which
was not observed in treated/untreated p53 knockout bone marrow (Fig. 3C).
DMBA-Induced Depletion of Specific Cell Populations in the
Bone Marrow Is p53-Dependent
Wright-stained bone marrow smears were prepared from the
tibias of each mouse, and the cells were assigned to the
erythroid, granulocyte, or lymphocyte lineages by a boardcertified veterinary clinical pathologist (P.S.M.). The granulocytic cells were classified as mature (postmitotic and including
metamyelocytes, bands, and segmented neutrophils) or immature cells (mitotic and including myeloblasts, progranulocytes,
and myelocytes). DMBA treatment of WT mice significantly
decreased the number of erythroid precursors in the bone
marrow of WT but not p53 null mice (Fig. 4A). The numbers
of mature granulocytes were also decreased in the bone marrow of DMBA-treated WT mice (Fig. 4C). Although there was
a slight decrease in the numbers of immature granulocytes in
the WT DMBA-treated mice, it was not statistically significant
(Fig. 4B). The large blast cells that were observed in the
DMBA-treated WT mice (Figs. 2C and 2F) were recorded as
immature granulocytes, which might explain the lack of a
significant decrease in the immature granulocytes. In addition,
there was also a p53-dependent significant (p ⬍ 0.05) decrease
in the lymphocyte population in the DMBA-treated WT mice
(Fig. 4D), which is consistent with the results we have previously reported (Heidel et al., 2000).
DMBA Does Not Induce Apoptosis in p53 Null Bone
Marrow B-Cell Cultures
We have previously reported that the murine pre-B cell line
(70Z3) undergoes apoptosis when it is cultured with bone
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FIG. 2. DMBA treatment affects the
cellularity in tibial bone marrow smears
from WT but not p53 null mice. Wrightstained bone marrow smears were prepared from the tibias of p53 null (A, B,
and E) and WT (C, D, and F) mice 48 h
after they were injected ip with 50 mg/kg
of DMBA (A, C, E, and F) or vehicle (B
and D). DMBA-treated WT mice exhibited abnormal granulocyte maturation
characterized by large blast cells of the
granulocyte lineage (C and F, black arrows) and aberrant, multinucleated granulocytes (F, right arrow). As a result of
the myeloid depletion, small lymphocytes appeared to be relatively more
prominent (C, white arrows). The photomicrographs above are representative
examples of mice used to obtain the data
in Figure 1. A, B, C, and D: ⫻250 magnification; E and F: ⫻400 magnification.
marrow stromal cells and DMBA. To determine whether this
process is p53-dependent, bone marrow cells from C57Bl/6
mice were harvested and cultured in vitro with IL-7 to stimulate the production of pre-B cells. The B cells were phenotyped
using surface marker staining and were found to be B220 ⫹,
IgM –, AA4.1 ⫹, and CD19 ⫹ (Fig. 5). This expression pattern is
consistent with the pre-B cell stage of B-cell development
(Thurmond and Gasiewicz, 2000).
These primary pre-B cells were then co-cultured with a
murine bone marrow stromal cell line (BMS2) in the presence
of DMBA and assessed for apoptosis by propidium iodide
staining. We found that DMBA-induced apoptosis in pre-B
cells derived from WT mice but not from p53 null mice (Fig.
6). Interestingly, the WT cells did not exhibit a dose-response
relationship between DMBA concentration and numbers of
apoptotic cells. Perhaps the primary bone marrow pre-B cells
are very sensitive to DMBA, and the doses tested were all
above the threshold needed to achieve a maximal response.
DISCUSSION
Previous research into the mechanisms of PAH-mediated
bone marrow toxicity has focused on the role of PAH metabolism by the CYP450 enzymes or the binding of PAH and
activated metabolites by the AhR. In our laboratory and others,
it has been demonstrated that PAHs must be metabolized by
CYP1B1 in order to be immunotoxic (Heidel et al., 1999,
2000; Mann et al., 1999; White et al., 1985). In this study, we
have focused on the contributions of p53, one of the key
regulators of cell death/survival after toxic insult. The results
presented here demonstrate the need for p53 protein in DMBAmediated bone marrow toxicity. Through the use of genedisrupted mice, we have observed that the toxic effects of
DMBA on the murine bone marrow and on primary cultures of
pre-B cells are absent in p53 null mice.
We have further characterized the effects of DMBA on the
bone marrow with respect to the cell type. Significantly, we
DMBA-INDUCED BONE MARROW TOXICITY
89
FIG. 3. Histopathological changes
in the bone marrow of DMBA-treated
WT but not p53 null mice. Sternums
were harvested from p53 null (A and B)
and WT (C and D) mice injected ip with
50 mg/kg DMBA (A and C) or vehicle
(B and D). The sternums were fixed, sectioned, and stained with haematoxylin
and eosin. The black arrows (C) denote a
dilated sinusoid with erythrocyte infiltration. The photomicrographs are representative of mice used to obtain the data in
Figure 1. ⫻100 magnification.
FIG. 4. Decreases in differential bone marrow cell counts are dependent
on p53. The mice were injected ip with 50 mg/kg DMBA or vehicle for 48 h:
(A) erythrocytes, (B) immature granulocytes, (C) mature granulocytes, and (D)
lymphocytes. The cell populations are expressed as the mean ⫾ SEM total
number of cells present in the bone marrow. The data represent five mice from
each treatment group and were analyzed using the unpaired t-test. Asterisks
indicate data that are significantly different from vehicle controls, p ⬍ 0.05.
noted a p53-dependent decrease in erythroid precursors after
DMBA treatment. Functional p53 was required for the depletion of all cell lineages examined (i.e., granulocytes, erythroid
precursors, and lymphocytes). This indicates that the majority
of DMBA-induced bone marrow toxicity may be mediated
through a p53-dependent pathway, even in disparate cell types.
To the best of our knowledge, this is the first observation of the
p53 dependence of DMBA-mediated bone marrow toxicity in
vivo.
The p53 protein plays a major role in the regulation of the
apoptotic cascade, so the observation that ablation of its activity protects against DMBA-mediated toxicity is consistent with
its known biological function. It has been very difficult to
detect apoptosis in the murine bone marrow in vivo after
DMBA treatment. One reason for this might be that bone
marrow macrophages phagocytose apoptotic cells very efficiently, so that at any given time there is a very small population of apoptotic cells present. The results of this study, in
conjunction with our previous observations using cell-culture
model systems, support the notion that DMBA causes apoptosis and/or growth arrest in the murine bone marrow.
It has been previously reported that activation of the nuclear
transcription factor NF-␬B protects pre-B cells against PAHinduced apoptosis. Mann et al. reported that PAHs suppress
NF-␬B and that suppression of NF-␬B by specific inhibitors
leads to pre-B cell apoptosis (Mann et al., 2001). NF-␬B
activation is, in general, an anti-apoptotic signal, whereas p53
activation is pro-apoptotic (Beg and Baltimore, 1996; Bertrand
et al., 1998; Levine, 1997). Other investigators have proposed
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FIG. 5. Nonadherent cells harvested from LTBMCs stain positively for
pre-B cell surface markers. Nonadherent cells from LTBMCs were stained
with the indicated fluorochrome-conjugated antibodies specific for (A) IgM,
(B) B220, (C) AA4.1, and (D) CD19 and analyzed for surface marker expression using flow cytometry. The cells were first gated on FSC and SSC for the
properties of viable lymphocytes. 10,000 events were then collected and
analyzed for each sample.
a link between p53 induction and NF-␬B regulation (HolmesMcNary et al., 2001; Webster and Perkins, 1999). In other
systems, the opposing regulation of p53 and NF-␬B has been
proposed as a mechanism of apoptosis induction (HolmesMcNary et al., 2001). These findings and the results of this
study suggest that PAH metabolites might cause apoptosis in
pre-B cells by both inducing p53 and inhibiting the activity of
NF-␬B.
DMBA is also carcinogenic, and p53 is often mutated in
transformed cells (Gupta et al., 1993). The requisite role of p53
in bone marrow toxicity suggests that the decreased cellularity
noted in WT mice may reflect the efforts of the immune system
to protect itself from the generation of leukemias and lymphomas following PAH exposure. p53-Induced apoptosis occurs
when the levels of DNA adduct formation caused by DNAdamaging agents (e.g., radiation, toxins, etc.) overwhelm the
DNA repair machinery of the cell. If PAH metabolites form
mutation-causing adducts in the p53 gene, the cells of the
immune system may lose the protective mechanism of eliminating aberrant cells, as suggested by the absence of bone
marrow hypocellularity in the p53 null mice. This could, in
turn, lead to a higher incidence of tumor formation in these
mice after DMBA exposure. Indeed, it has been reported that
FIG. 6. Primary pre-B cells from WT but not p53 null mice undergo
apoptosis when cultured with stromal cells and DMBA in vitro. Primary pre-B
cells were generated from the bone marrow of both WT and p53 knockout
mice, as described in the methods. The pre-B cells were then isolated and
cultured with bone marrow stromal cells (BMS2 cells) for 24 h with the
indicated concentrations of DMBA. Afterward, the pre-B cells were collected,
stained with propidium iodide, and analyzed by flow cytometry, as described
in Materials and Methods. (A) is a representative dot plot showing forward and
side scatter (all cells were gated on R1) and a representative histogram for
propidium iodide-stained WT cells treated with 10 ␮M DMBA in vitro. The
sub G 0 population of cells, denoted as M1, were calculated for each sample and
are reported as the percentage of apoptotic cells in (C). (B) shows the same
plots for p53 null cells treated with 10 ␮M DMBA. In (C), the open bars are
WT pre-B cells, and the shaded bars are p53 null pre-B cells. The data are
expressed as the mean ⫾ SEM percentage of apoptotic cells for WT pre-B cells
(open square) and p53 null pre-B cells (filled square), (n ⫽ 7, wild type; n ⫽
4, p53 null).
DMBA-INDUCED BONE MARROW TOXICITY
p53 null mice have a higher incidence of mammary tumors
after DMBA administration and are more susceptible to teratogenic malformation after ionizing radiation than are WT
mice (Jerry et al., 2000; Kato et al., 2001).
Although it is possible that the toxicity observed in the bone
marrow after DMBA exposure might protect against leukemia
or lymphoma, disruption of B-cell development can have profound deleterious effects on acquired immunity. Previous studies have documented a decrease in B-cell reactivity and IgM
production among workers occupationally exposed to PAHs
(Winker et al., 1997). In addition, it has also been demonstrated that the potent AhR agonist, TCDD, alters B-cell populations (Thurmond and Gasiewicz, 2000). The precise mechanisms underlying these adverse effects on B cells are
unknown. In this study, we demonstrate that the effects of
DMBA on the murine bone marrow are dependent on functional p53. Previous work in our laboratory found that the
interferon-inducible kinase (PKR) is upregulated in pre-B cells
in response to DMBA treatment (Page et al., 2002). This
observation is important because PKR has been shown to
activate p53 (Cuddihy et al., 1999a,b). Coupled together, these
observations point to a possible pathway of DMBA-mediated
bone marrow toxicity. Our working model is that metabolites
of DMBA stimulate bone marrow cells to release a soluble
factor (e.g., cytokine or chemokine) that leads to PKR activation, which in turn activates p53 and causes apoptosis in the
affected cell types.
In conclusion, this study has demonstrated that p53 protein
is necessary for bone marrow depletion of granulocytic, erythroid, and lymphocytic cells after DMBA treatment. In addition, we have demonstrated that pre-B cells derived from the
bone marrow of p53 null mice are also resistant to DMBAmediated apoptosis in vitro. These studies provide new insights
into the molecular mechanisms underlying DMBA-induced
immunotoxicity.
ACKNOWLEDGMENTS
This work was supported by National Institutes of Health grants RO1CA81493 and P30-ES09090. T.J.P. is supported by National Research Service
Award Fellowship F32-ES11073-01.
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