![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
|
|
|
|
|
||
2
*
Department of Biochemistry and Winship Cancer Center, Emory University, Atlanta, GA 30322;
Department of Pharmacology and Toxicology, Morehouse School of Medicine, Atlanta, GA 30310;
**
Division of Environmental and Occupational Health Sciences, School of Public Health, Emory University, Atlanta, GA 30322;
Graduate Program in Nutritional Health Sciences, Emory University, Atlanta, GA 30322
2To whom correspondence should be addressed.
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSTime course of development...DISCUSSIONREFERENCES |
|---|
KEY WORDS: • NAD(P)H:quinone reductase • glutathione S-transferase • butyrate • apoptosis • human colon carcinoma cells
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSTime course of development...DISCUSSIONREFERENCES |
|---|
). While there are
several mechanisms that could contribute to this association, a
well-characterized defense mechanism involves the induction of
detoxification enzymes, including members of the glutathione
S-transferase family and NAD(P)H:quinone reductase (quinone
reductase) (Coles and Ketterer 1990
, Hayes et al. 1991
, Joseph and Jaiswal 1994
, Lin et al. 1994
). These enzymes are generally referred to as "Phase 2"
enzymes because they catalyze conversion of mutagenic metabolites or
their precursors to compounds that are less reactive and/or more
readily excreted. This detoxification function is in contrast to
"Phase 1" enzymes, such as cytochrome P-450s, which bioactivate
foreign compounds to DNA-reactive metabolites and contribute to
carcinogenesis (Hashimoto and Degawa 1995
, Joseph and Jaiswal 1994
, Lin et al. 1994
).
More than 40 compounds were identified from dietary sources that
function as Phase 2 enzyme inducers (Fukushima et al. 1997
, Prestera et al. 1993
, Steinmetz and Potter 1991
, Talalay et al. 1988
,
Wattenberg 1992
). Many of these compounds were tested in
animal models and found to increase Phase 2 enzyme activities and
protect against cancer (Wattenberg 1992
). For instance,
diallyl sulfide (Haber-Mignard et al. 1996
,
Sumiyoshi and Wargovich 1990
), dithiolthiones
(Rao et al. 1991
), benzyl selenocyanate (Reddy et al. 1987
) and butylated hydroxyanisole (Reddy and Maeura 1984
) increase glutathione S-transferase
and/or quinone reductase activity, inhibit DNA adduct formation and
inhibit the incidence and multiplicity of colon carcinomas in rats and
mice. Taken together with the epidemiologic data, these studies suggest
that specific chemicals in the human diet protect against cancer by
increasing the activities of detoxification enzymes, thereby decreasing
the concentrations of genotoxic compounds that would otherwise give
rise to carcinogenic mutations.
Phase 2 enzyme inducers encompass a broad range of chemical structures
but generally share the property of directly affecting the cellular
glutathione pool, either by being substrates of glutathione
S-transferases or directly reacting with glutathione
(Spencer et al. 1991
). Mechanistic studies show that the
increase in enzyme activities can be controlled at the transcriptional
level signaled by depletion or oxidation of the glutathione pool
(Bergelson et al. 1994
, Daniel 1993
,
Galter et al. 1994
). This mechanism may be of particular
importance in tumorigenesis because depletion or oxidation of the
glutathione pool was also linked to apoptosis in several systems
(Abello et al. 1994
, Buttke and Sandstrom 1994
, Fernandes and Cotter 1994
, Potten 1992
, Slater et al. 1995
). Accumulating data
indicate that failure of normal apoptosis is an important mechanism in
tumor development (Evan and Littlewood 1998
). Agents
that stimulate apoptosis in precancerous cells therefore might be
cancer preventive by enhancing apoptosis in tumorigenic cell
populations.
The common feature of glutathione depletion/oxidation in both Phase 2 enzyme induction and activation of apoptosis led us to examine whether Phase 2 enzyme inducers could activate apoptosis over the same concentration range that increases enzyme activity. For this study, we used a moderately differentiated human colon carcinoma cell line (HT29) as a model system and representative inducers (benzyl isothiocyanate, allyl sulfide, dimethyl fumarate, butylated hydroxyanisole) at concentrations that increase Phase 2 enzyme activities as represented by quinone reductase and glutathione S-transferase activities. Results show that these inducers activated apoptosis over an 8 to 24 h time course, as indicated by cell detachment from the dishes, nuclear condensation and caspase activation. Pretreatment of cells with a differentiating agent, sodium butyrate, increased both the extent of Phase 2 enzyme induction and activation of cell death. Thus, the results show that the induction of apoptosis is concomitant with Phase 2 enzyme induction in this cell line and suggest that this induction of apoptosis may contribute to in vivo anticancer effects of these compounds.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSTime course of development...DISCUSSIONREFERENCES |
|---|
Allyl sulfide, benzyl isothiocyanate, butylated hydroxyanisole, dimethyl fumarate, dimethyl sulfoxide (DMSO3 ), sodium butyrate and Sudan I were purchased from Aldrich Chemical Co., Inc. (Milwaukee, WI). Cell culture medium and serum were purchased from Life Technologies, Inc. (Grand Island, NY). In situ cell death detection kit, fluorescein, was from Boehringer Mannheim Co. (Indianapolis, IN). Propidium iodide was from Sigma Chemical Co., (St. Louis, MO). DEVD-AMC was from Peptide International, Inc. (Louisville, KY).
Cell culture and incubations.
HT29 cells obtained from American Type Culture Collection were grown in McCoy's medium supplemented with 10% fetal bovine serum, in a humidified incubator at 37°C with an atmosphere of 95% air, 5% CO2. In our standard induction protocol, HT29 cells were plated at 0.5 x 106 cells per 60 mm plate and grown for 80 h to late logarithmic phase (70–80% confluence). Test compounds in DMSO (0.2%, vol/vol) were added directly to the medium and the cells were incubated for up to 24 h. Controls received a change of medium containing 0.2% DMSO, which had no detectable effect on cell differentiation or enzyme expression.
For studies of cell differentiation by sodium butyrate treatment, HT29 cells were seeded at 0.5 x 106 cells into 60 mm plates and grown in standard medium for 72 h. Medium was then changed to McCoy's containing 5 mmol/L sodium butyrate, for an additional 72 h. Differentiation was assessed by alkaline phosphatase activity which increased greater than 10-fold by 3 d. Cells were incubated with inducing compounds, added directly to the medium, for up to 24 h as indicated. Cell viability was determined as the percentage of cells that excluded 0.2% (wt/vol) trypan blue.
Enzyme activity determinations.
After exposure to inducing agents, cells were collected from the plates by scraping, frozen in liquid nitrogen and stored at -80°C. For quinone reductase and glutathione S-transferase activity assays, thawed cell suspensions, in 1 mmol/L sucrose, were centrifuged at 9,000 x g for 15 min at 4°C. To the supernatant, 0.2 vol of 0.1 mol/L of CaCl2/0.25 mol/L sucrose was added. The mixture was incubated 15 min on ice and then centrifuged at 15,000 x g for 20 min. The supernatants were used as cytosolic fractions for enzyme assays.
Quinone reductase activity was assayed as the rate of reduction of
2,6-dichloroindophenol (40 µmol/L) by NADH (200 µmol/L) at pH 7.0
measured spectrophotometrically at 600 nm in the presence and absence
of 10 µmol/L of dicoumarol. The dicoumarol-sensitive portion of
the activity was taken as a measure of the quinone reductase activity
(Benson et al. 1980
). Glutathione
S-transferase activity was measured spectrophotometrically
at 340 nm with 1-chloro-2,4-dinitrobenzene (1 mmol/L) and glutathione
(1 mmol/L) as substrates (Habig and Jakoby 1981
).
Protein concentrations were determined using the Bradford method
(Bradford 1976
) with bovine serum albumin as standard.
Propidium iodide staining.
Cells collected from either the medium or the plates were centrifuged
at 100 x g for 10 min. For fluorescence microscopy,
pellets were resuspended in 1 ml of phosphate-buffered saline (PBS)
and a 100 µL aliquot was centrifuged onto microscope slides in a
microcentrifuge at 150 x g for 10 min. Cells adhering
to microscope slides were fixed in 80% methanol for 20 min, rinsed
three times with PBS, and incubated with 50 µL of propidium iodide
(75 µmol/L) for 5 min. Photomicrographs were made with a rhodamine
filter set on an inverted Zeiss fluorescent microscope. For flow
cytometry, cells were fixed and stained with propidium iodide in
Eppendorf tubes, washed with PBS and analyzed with a
Becton-Dickinson FACScan station. The doublet discrimination module
was activated to exclude the cell aggregates and the red fluorescence
(FL-3A) was recorded as an indication of DNA content. Apoptotic cells
lost their fragmented DNA during sample preparation and had a sub-G1
distribution on the FL-3A plot (McConkey et al. 1989
,
Nicolletti et al. 1991
).
DNA fragmentation assay.
After treatment, medium was removed, and cells remaining on the culture
plate were rinsed twice with PBS and lysed by addition of 2 mL of
ice-cold lysis buffer containing 5 mmol/L of Tris, 20 mmol/L of
EDTA and 0.5% (vol/vol) of Triton X-100, pH 8.0. Lysate was
transferred to centrifuge tubes and left on ice for 15 min, then
centrifuged at 27,000 x g for 20 min to separate
intact chromatin (pellet) from fragmented DNA (supernatant)
(McConkey et al. 1989
). Pellet and supernatant DNA
concentrations were measured by a diphenylamine reaction (Burton 1968
).
Caspase activity measurement.
The group II caspase activity was measured with a fluorogenic substrate
DEVD-AMC as described (Stridh et al. 1998
). Cells
were collected from the plates and briefly centrifuged in
microcentrifuge tubes. The pellets were brought up in 100 µL of PBS,
and 50 µL was transferred directly to single wells of a 96-well plate
pre-cooled on dry ice. The remaining volume was used for protein
determinations. The reaction was started by adding 50 µL of
pre-warmed buffer containing 100 mmol/L of HEPES, 10% sucrose
(wt/vol), 0.1% CHAPS (wt/vol), 5 mmol/L of dithiothreitol,
10-6:1 (vol/vol) Nonidet P-40 and 50 µmol/L of
DEVD-AMC. The kinetics of the fluorescence change within the next
45 min were recorded on a Packard fluorescent plate reader. As
described for the enzyme assay, data were normalized with the measured
protein amount. Data from each experiment were calculated as percentage
of control, combined and statistically analyzed as described for the
enzyme assay.
Data analysis.
Experiments were designed to minimize variability in quinone reductase
and glutathione S-transferase activity due to cell density
by establishing a constant density protocol and calculating data as the
percentage of control activity within each. Tests for statistically
significant differences (
= 0.05) were performed by one-way
ANOVA and Dunnett's multiple range tests, with all treatments compared
to the control. Significant differences are reported for values where
variances were equal.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSTime course of development...DISCUSSIONREFERENCES |
|---|
To determine basal activities of quinone reductase and glutathione
S-transferase in normally proliferating HT29 cells, cultures
were maintained for 80 h (~70% confluence). Measured activities
were 1.8 ± 0.4 and 0.4 ± 0.1 µmol ·
min-1 · mg protein-1, respectively,
and were not significantly changed during the 72–160 h culture periods
of individual experiments or following changes in control medium.
Following 24-h exposure to inducing compounds (benzyl isothiocyanate,
dimethyl fumarate, allyl sulfide, butylated hydroxyanisole),
statistically significant increases (P < 0.05) in
quinone reductase (30–82%) and glutathione S-transferase
activities (36 to 86%) over time-matched controls were found for
benzyl isothiocyanate, allyl sulfide and butylated hydroxyanisole)
(Table 1
).Benzyl isothiocyanate was the most potent of these compounds, followed
by allyl sulfide, dimethylfumarate and butylated hydroxyanisol. The
extents of induction were comparable to those reported in animal
studies (Sparins et al. 1982
, Vos et al. 1988
).
|
Under conditions giving enzyme induction, the same concentrations of
chemicals caused a significant (P < 0.05) increase in
number of cells that detached from culture plates by 20 h for
benzyl isothiocyanate (18-fold), allyl sulfide (5-fold) and dimethyl
fumarate (4-fold) (Fig. 1
A),with a relative response in the sequence benzyl isothiocyanate > allyl sulfide > dimethyl fumarate. For comparison, Sudan I, a
compound that induces both Phase 1 and Phase 2 enzymes was also
studied. The results showed that this compound also increased cell
detachment at a concentration that increased the activities of
glutathione S-transferase and quinone reductase. The number
of detached cells from control plates treated with 0.2% of DMSO was
typically in the range of 1–2%. Detachment of cells is a common
feature of apoptosis in tissue culture that is thought to parallel the
separation of apoptotic cells that occurs during apoptosis in vivo
(Wyllie et al. 1980
). To verify that the released cells
had not undergone necrotic lysis, cells were examined with 0.2% of
trypan blue, and greater than 90% of the detached cells was found to
exclude the dye.
|
We examined cells by fluorescence microscopy with propidium iodide
staining to determine whether cells treated with benzyl isothiocyanate
had undergone changes in nuclear morphology characteristic of
apoptosis. Control cells were rather uniform and flat with normal
heterochromatin staining (Fig. 2
A).Most of the treated cells remained on the plates and showed some
chromatin condensation and pyknotic and fragmented nuclei at 16 h
(Fig. 2
B). However, >90% of the cells that detached had
condensed and/or fragmented nuclei (P < 0.05, data not
shown). Thus, a fraction of HT29 cells treated with the detoxification
enzyme inducer benzyl isothiocyanate exhibited morphological changes
associated with apoptosis, i.e., loss of adherence, nuclear
condensation and nuclear fragmentation prior to permeability change of
the plasma membrane.
|
|
Time course of development of apoptosis |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSTime course of development...DISCUSSIONREFERENCES |
|---|
B). For
dimethyl fumarate, there was also a progressive increase in detached
cells, but the increase was significant only at 24 h (P
< 0.05 Fig. 1A
). These results show that the development of
apoptosis is relatively slow and probably incomplete even at 24 h. Effect of sodium butyrate pretreatment on benzyl isothiocyanate-induced cell death.
Treatment of HT29 cells with sodium butyrate is known to result in
differentiation which ultimately leads to apoptosis (Heerdt et al. 1994
). To determine whether differentiation by sodium
butyrate affected the response to a Phase 2 enzyme inducer, we examined
cells following 72 h of treatment with sodium butyrate (5 mmol/L)
without and with benzyl isothiocyanate. Sodium butyrate alone showed no
significant increase in the number of detached cells at 3 d of
treatment (Fig. 3
A, 0 h) compared to control cells at 3 or 0 d controls (Fig. 1
A, control). However, there was a progressive, modest
increase in detached cells over the subsequent 24 h which was
similar to that previously reported (Heerdt et al. 1994
). These detached cells had condensed nuclei (Fig. 2
E) and included a higher percentage that stained with
trypan blue. Thus, over the longer time course of treatment with sodium
butyrate, cells became apoptotic and then appeared to undergo a
secondary necrosis in which they lost the integrity of their plasma
membrane.
|
A).
Significant decreases (P < 0.05) were observed in the
number of cells remaining attached to the culture plates at 8; 16 and
24 h were also seen for benzyl isothiocyanate butyrate-treated
cells (Fig. 3
B). When examined by fluorescence microscopy in
the presence of propidium iodide, >90% of cells had morphologic
features of apoptosis, i.e., extensive nuclear condensation and
fragmented nuclei (Fig. 2
F). These results suggest that the
differentiated cells undergo enhanced cell death in response to benzyl
isothiocyanate beyond that of sodium butyrate or benzyl isothiocyanate
alone. DNA fragmentation after treatment with enzyme inducers.
To further examine DNA fragmentation, we used the TUNEL assay with
fluorescence confocal microscopy. The results confirmed that benzyl
isothiocyanate induced DNA fragmentation, and this was dramatically
increased in sodium butyrate-pretreated cells compared to controls
(Fig. 2
C, D, G, H).
Fragmentation of DNA can also be detected by flow cytometry in cells
stained with propidium iodide. With this technique, most cells are in
GoG1 phase and appear as a homogeneous peak of cells with a normal DNA
content. During apoptosis, fragmentation and loss of DNA results in an
appearance of particles (cells and apoptotic bodies) with less
fluorescence than the GoG1 cells. As shown in Figure 4
B, cells treated with sodium butyrate and then with either
benzyl isothiocyanate or dimethyl fumarate had a substantial increase
in this sub-G1 population of cells (from <8% to >30%). Thus, these
results further support the interpretation that apoptosis occurs in
response to the inducing agents.
|
B).Analysis of DNA fragmentation by agarose-gel electrophoresis
stained with ethidium bromide revealed only a weak laddering pattern
(data not shown) comparable to that previously reported for HT29 cells
following butyrate-induced differentiation (Heerdt et al. 1994
). Thus, by multiple criteria, DNA fragmentation occurred
in a pattern consistent with the morphologic changes of apoptosis. Caspase activation after treatment with enzyme inducers.
DNA fragmentation into oligonucleosomal lengths is thought to be
mediated by a process activated by a proteolytic cascade involving
caspases (Thornberry and Lazebnik 1998
). To determine
whether caspases were activated as a result of treatment with Phase 2
enzyme inducers, we used a Caspase-3 substrate DEVD-AMC that is
cleaved to a fluorescent product by Caspase-3 and other caspases with
similar substrate cleavage sequences. Results showed that
Caspase-3-like activity increased about 2-fold (P < 0.05) in cell extracts following treatment of cells with benzyl
isothiocyanate or dimethyl fumarate (Fig. 5
A).There was greater than 20-fold increase in activity following
pretreatment of cells with sodium butyrate (P < 0.05;
Fig. 5
B). Thus, the results show that caspase activation
occurred in a pattern that is consistent with the DNA fragmentation and
the morphologic evidence of apoptosis following treatment with either
benzylisothiocyanate or dimethyl fumarate.
|
To further test whether induction of apoptosis is a general response to
detoxification enzyme inducers, we used a spectrum of compounds at
concentrations known to result in increased activity of quinone
reductase and glutathione S-transferase (Table 1
;
Bergelson et al. 1994
, Daniel 1993
,
Galter et al. 1994
, Prestera et al. 1993
). Allyl sulfide resulted in no significant increase in
caspase activity compared to control cells, but in nearly a 5-fold
activation in sodium butyrate-pretreated cells (P
< 0.01; Fig. 6
).Butylated hydroxyanisole resulted in significant increases in both
untreated and sodium butyrate-pretreated cells (Fig. 6)
. Both of
these inducers resulted in activation at levels similar to benzyl
isothiocyanate and dimethyl fumarate (Fig. 6)
. In addition, Sudan I,
which is thought to induce detoxification enzyme activity through
interaction with the xenobiotic response element, also induced similar
activation of caspase 3 (Fig. 6)
. Thus, the results show that induction
of apoptosis, as defined by cell morphology and caspase activation, is
a common response to agents that induce increased activity of the
detoxification enzymes quinone reductase and glutathione
S-transferase and that differentiation with sodium butyrate
results in a parallel increase in both the increase in enzyme activity
and induction of apoptosis.
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSTime course of development...DISCUSSIONREFERENCES |
|---|
). More recent studies indicate that these features can be
largely explained by activation of a caspase proteolytic cascade
(Samali et al. 1999
, Thornberry and Lazebnik 1998
).
Apoptosis increases in response to exposures to cytotoxic agents
(Thompson 1995
). Potten (1992)
speculated
that spontaneous apoptosis is actually the removal of damaged cells
that contain random genetic defects due to interactions with
DNA-damaging compounds. Failure of apoptosis in cells with specific
mutations may therefore contribute to oncogenesis. Removal of damaged
cells could be a particularly useful defense mechanism within the
gastrointestinal tract because of its contact with dietary mutagens
(Ohgaki et al. 1991
).
In the human colon, epithelial cells form a single layer, with the
proliferative cells arranged in invaginated crypts and differentiated
cells on the luminal surface (Potten 1992
). Migration of
cells from the base of the crypt to the surface is estimated to require
3 to 8 d, while stem cell cycle times are ~36 h. This rate of
proliferation slows in association with differentiation, and the cells
are ultimately sloughed into the lumen. Detoxification enzyme
expression is low in normal crypt cells and increases in the
differentiated surface cells (Hayes et al. 1989
,
Ranganathan and Tew 1991
). Thus, there is an increase in
detoxification enzyme expression that parallels decreased proliferation
rate and progression of cells in terminal differentiation.
Induction of the detoxification enzymes quinone reductase and
glutathione S-transferase is a well-characterized
defense mechanism against carcinogens (Coles and Ketterer 1990
, Prestera et al. 1993
). In principle,
elevation of these enzymes can reduce carcinogenesis due to enhanced
removal of reactive electrophiles. Indeed, in vitro and in vivo
research supports the interpretation that inducibility of these enzymes
may be an important determinant of cancer risk (Hayes et al. 1991
, Joseph and Jaiswal 1994
, Lin et al. 1994
). Foods that contain compounds that induce detoxification
enzymes include members of several vegetable families, such as
Cruciferae (broccoli, Brussels sprouts, cabbage, kale, cauliflower),
Leguminosae (green beans), Umbelliferae (carrots, celery),
Zingerberaceae (ginger), Liliaceae (asparagus, green onions, leeks),
Compositae (leaf lettuce) and Chenopodiaceae (spinach)
(Prochaska et al. 1992
).
For compounds in foods to be chemopreventive, consumption of the
relevant foods must be sufficient to attain the cellular concentrations
of the inducers needed for enzyme induction. Although this cannot be
readily predicted because of the multiple issues of bioavailability,
distribution and clearance rates, available evidence indicates that
relevant concentrations are likely to be achieved under normal biologic
conditions. For instance, the study of Prochaska et al. (1992)
reported potency values for quinone reductase induction
of greater than 10,000 units/g for broccoli, where one unit was defined
as the amount of inducer required to double the quinone reductase
activity of Hepa 1c1c7 cells growing in 150 µL wells. Mean portion
size for broccoli is about 85 g (Block et al. 1992
)
so that a serving would contain about 850,000 units. If the relevant
compounds are 100% absorbed and uniformly distributed into the entire
50 L of body water, one would need 50 L/0.00015 L, or 333,333 units for
a 2-fold induction. Thus, the amount in a normal serving of broccoli
would appear to be sufficient to cause the induction. Since the liver
and intestinal epithelium are exposed to higher concentrations than
that achieved by uniform delivery throughout the body, at least these
tissues should be exposed to concentrations sufficient for induction.
This interpretation is supported by data from Hecht (1995
, Hecht 1999
), who estimated
that isothiocyanates, initially released from cruciferous vegetables as
glucosinolates and then hydrolyzed by myrosinase, represent about
0.02% of the vegetable mass. Assuming an average molecular mass for
isothiocyanates of 170 and a portion size of 100 g, this would
indicate that about 20 mg of isothiocyanates, or 100 µmol would be
consumed. If 5% remained in the intestinal tract and reached the colon
in 250 mL, the concentration would be similar to the concentration of
benzylisothiocyanate used in the present study (i.e., about 20
µmol/L). Similar calculations can be made for fumarates and allyl
sulfide. Thus, the concentrations of inducers used in the present study
appear relevant to conditions that are achieved in vivo.
The results show that the effects of the inducers are enhanced by
pretreatment with 5 mmol/L of sodium butyrate. Butyric acid is
generated in the human colon from the fermentation of fiber.
Concentrations in the range of 5 mmol/L are normally found, and
concentrations of over 20 mmol/L are generated by high-fiber diets
(Cummings et al. 1987
, Kapadia et al. 1995
, Kashtan et al. 1992
). Thus, the
combination of butyrate and inducers used in the present study appears
to replicate conditions that could be achieved by a high-fiber,
high-fruit and -vegetable diet. Importantly, the potentiation of
effects of inducers by pretreatment with butyrate suggests a potential
mechanistic basis for an interactive effect in cancer prevention
between high-fiber foods and foods that contain high concentrations
of inducers.
Activation of apoptosis and induction of detoxification enzymes are
both sensitive to changes in cellular GSH, and measurements show that
substantial changes in GSH concentration and GSH/GSSG redox state occur
in HT29 cells under the butyrate and benzyl isothiocyanate treatment
conditions as used in the present study (Jones et al. 1996
). However, it is not clear whether activation of apoptosis
and enzyme induction are mechanistically linked. Briehl and Miesfeld (1991)
found that androgen withdrawal in rat ventral
prostate induced apoptosis and also elevated glutathione
S-transferase mRNA, suggesting that the two processes could
be mechanistically linked. In a second study (Flomerfelt et al. 1993
), however, they found that the elevated glutathione
S-transferase in steroid-induced apoptosis is not an
essential step in the apoptotic process but is a coincidental response
to a change in cellular redox state. Thus, while the data are
consistent with both processes being associated with changes in
cellular GSH, the data are presently insufficient to conclude that they
are mechanistically linked.
Whether or not apoptosis and detoxification enzyme induction share a common GSH-dependent mechanism, the elicitation of both responses by Phase 2 inducers in butyrate-differentiated cells could be important because this could provide a greater anticarcinogenic potential than achieved by one mechanism alone. Our results show that chemically unrelated compounds that induce detoxification enzymes also induce apoptosis in the colon carcinoma cells over the same concentration ranges that increase enzyme activity. This apoptotic effect was even more pronounced in the cells that had been differentiated with sodium butyrate. Thus, these compounds may provide two different mechanisms to protect against cancer, namely enhanced elimination of chemical carcinogens and enhanced elimination of precancerous cells.
The concept of dietary chemoprevention is usually used in the context
of protecting normal cells from initiating events that introduce
oncogenic mutations. However, substantial literature is available to
show that carcinogenesis represents a progression of cellular changes
(Vogelstein and Kinzler 1994
), and agents that disrupt
this progression at any point can be considered chemopreventive. In the
present case, we are suggesting that stimulation of apoptosis in
precancerous cells could block or delay tumor development. The HT29
cells used in the present study represent an advanced stage of tumor
development. Therefore, we consider it especially significant that
dietary agents induce apoptosis in these cells at concentrations that
may be achieved in the normal diet. This raises the possibility that
some agents may be particularly effective in prevention of colon
carcinoma formation because they can enhance elimination of
precancerous cells. While it remains unknown whether this occurs in
vivo, if it does, it would provide two mechanisms for cancer prevention
by Phase 2 enzyme inducers. Specifically, these compounds could induce
an increase in detoxification activity and activate apoptosis in
precancerous cells. Thus, one may postulate that such compounds in the
human diet may provide a dual protective mechanism against colon
cancer.
While this possibility is speculative, a comparison of anticarcinogenic effects by these two mechanisms is of interest. For protection due to enhanced detoxification of carcinogens, exposure to the inducer would have to occur several hours prior to exposure to the carcinogen in order to allow time for an increase in detoxification enzyme activity. For a long-term chemopreventive strategy, protection could be maintained only if there were a permanent increase in enzyme expression or a continuous exposure to the inducer. In contrast, if these compounds enhance apoptosis, then they could provide a cancer-preventive effect even after exposure to a chemical carcinogen which caused an oncogenic mutation.
There is extensive epidemiologic literature on the relationships
between consumption of fiber (butyrate source), consumption of
vegetables and fruits (sources of Phase 2 inducers) and colon cancer
(Block et al. 1992
, National Research Council 1989
, Steinmetz and Potter 1991
). Block et al. (1992)
found that 20 out of 23 epidemiologic studies on
fruits, vegetable and colon cancer indicated that diets high in fruits
and vegetables were protective (average relative risk of 1.9 for low
consumption). The National Research Council (1989)
cited
several case-control and correlation studies which showed inverse
relationships between the intake of high-fiber foods and colon
cancer risk but noted that these foods (vegetables to a large extent)
are rich sources of other nutritive and nonnutritive constituents which
could be cancer-inhibiting so that the effects could not be
attributed to fiber, per se. Other studies suggest that cruciferous
vegetables have a protective effect against colon cancer (Kune et al. 1987a
, Kune et al. 1987b
,
Miller et al. 1983
, Young and Wolf 1988
).
In light of the present results, it may be useful to more specifically
consider interactive effects of high-fiber diets and foods high in
Phase 2 inducers in colon cancer risk. Alternatively, this potential
interactive effect could be directly studied in animal models of colon
carcinogenesis, and studies of early and late interventions could help
discriminate between a mechanism involving enhanced detoxification of
mutagenic compounds and one depending upon elimination of precancerous
cells.
In conclusion, the present studies show that compounds that induce Phase 2 detoxification enzymes also induce apoptosis over the same concentration range in HT29 cells. The results indicate that these dietary inducers may therefore protect against cancer by two distinct mechanisms, enhanced detoxification of carcinogenic compounds and enhanced elimination of precancerous cells. Induction of apoptosis in precancerous cells may provide protection against cancer development even after chemical-induced mutagenesis and, therefore, may provide the basis for a novel nutritional strategy for cancer prevention. The practical implication is that consumption of diets high in butyrate production (e.g., fiber) and high in Phase 2 inducers (e.g., cruciferous vegetables) may trigger precancerous cells to die by apoptosis. This could "prevent" development of carcinoma by eliminating cells with an incomplete set of cancer-causing mutations. Such a possibility may be particularly useful in at-risk populations. However, this implication is derived solely from an in vitro study with one colon carcinoma cell line. Clearly, additional in vivo and clinical studies are necessary to assess the usefulness of such a diet as an anticancer strategy.
|
FOOTNOTES |
|---|
3 Abbreviations used: CHAPS,
(3-[(3-cholamidopropyl)dimethylammonio]-1-propane sulfonic acid;
DMSO, dimethyl sulfoxide; PBS, phosphate-buffered saline.
Manuscript received April 7, 1999. Initial review completed May 17, 1999. Revision accepted July 19, 1999.
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSTime course of development...DISCUSSIONREFERENCES |
|---|
1. Abello P. A., Fidler S. A., Buchman T. G. Thiol reducing agents modulate apoptosis in porcine endothelial cells. Shock 1994;2:79-83[Medline]
2.
Benson A. M., Hunkeler M. J., Talalay P. Increase of NAD(P)H:quinone reductase by dietary antioxidants: Possible role in protection against carcinogenesis and toxicity. Proc. Nat. Acad. Sci. USA 1980;77:5216-5220
3. Bergelson S., Pinkus R., Daniel V. Induction of AP-1 (Fos/Jun) by chemical agents mediates activation of glutathione S-transferase and quinone reductase gene expression. Oncogene 1994;9:565-571[Medline]
4. Block G., Dresser C. M., Hartman A. M., Carroll M. D. Nutrient sources in the American diet: quantitative data from the NHANES II survey. I. Vitamins and coenzymes. Am. J. Epidemiol. 1985;112:13-26
5. Block G., Patterson R., Subar A. Fruit, vegetables and cancer prevention: a review of the epidemiological evidence. Nutr Cancer 1992;18:1-29[Medline]
6. Bradford M. M. A rapid and sensitive method for quantification of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 1976;72:248-254[Medline]
7.
Briehl M. M., Miesfeld R. L. Isolation and characterization of transcripts induced by androgen withdrawal and apoptotic cell death in the rat ventral prostate. Mol. Endocrinol. 1991;5:1381-1388
8. Burton K. Determination of DNA concentration with diphenylamine. Meth. Enzymol. 1968;12:163-166
9. Buttke T. M., Sandstrom P. A. Oxidative stress as a mediator of apoptosis. Immun. Today 1994;15:7-10
10. Coles B., Ketterer B. The role of glutathione and glutathione transferases in chemical carcinogenesis. Crit. Rev. Biochem. Molec. Biol. 1990;25:47-70[Medline]
11.
Cummings J. H., Pomare E. W., Branch W. J., Naylor C.P.E., McFarlane G. T. Short-chain fatty acids in human large intestine, portal, hepatic and venous blood. Gut 1987;28:1221-1227
12. Daniel V. Glutathione S-transferases: Gene structure and regulation of expression. Crit. Rev. Biochem. Molec. Biol. 1993;28:173-207[Medline]
13.
Evan G., Littlewood T. A matter of life and cell death. Science 1998;281:1317-1322
14. Fernandes R. S., Cotter T. G. Apoptosis or necrosis: intracellular levels of glutathione influence mode of cell death. Biochem. Pharmacol. 1994;48:675-681[Medline]
15. Flomerfelt F. A., Briehl M. M., Dowd D. R., Dieken E. S., Miesfeld R. L. Elevated glutathione S-transferase gene expression is an early event during steroid-induced apoptosis. J. Cell Physiol. 1993;154:573-581[Medline]
16. Fukushima S., Takada N., Hori T., Wanibuchi H. Cancer prevention by organosulfur compounds from garlic and onion. J. Cell Biochem. (Supplement) 1997;27:100[Medline]
17.
Galter D., Mihm S., Dröge W. Distinct effects of glutathione disulphide on the nuclear transcription factors 
B and the activation protein-1. Eur. J. Biochem. 1994;221:639-648[Medline]
18. Haber-Mignard D., Suschetet M., Berges R., Astorg P., Siess M. H. Inhibition of aflatoxin B1- and N-nitrosodiethylamine-induced liver preneoplastic foci in rats fed naturally occurring allyl sulfides. Nutr. Cancer 1996;25:61-70[Medline]
19. Habig W. H., Jakoby W. B. Assays for differentiation of glutathione S-transferases. Meth. Enzymol. 1981;77:398-405[Medline]
20. Hashimoto Y., Degawa M. Induction of cytochrome P450 isoforms by carcinogenic aromatic amines and carcinogenic susceptibility of rodent animals. Pharmacogenetics (spec) 1995;5:s80-s83
21.
Hayes P. C., Harrison D. J., Bouchier I.A.D., McLellan L. I., Hayes J. D. Cytosolic and microsomal glutathione S-transferase isoenzymes in normal human liver and intestinal epithelium. Gut 1989;30:854-859
22. Hayes J. D., Judah D. J., McLellan L. I., Kerr L. A., Peacock S. D., Neal G. E. Ethoxyquin-induced resistance to aflatoxin B1 in the rat is associated with the expression of a novel alpha-class glutathione S-transferase subunit, Yc2, which possesses high catalytic activity for aflatoxin B1-8,9-epoxide. Biochem. J. 1991;279:385-398
23. Hecht S. S. Chemoprevention by isothiocyanates. J. Cell. Biochem. 1995;22(Suppl):195-209
24. Hecht S. S. Chemoprevention of cancer by isothiocyantes, modifiers of carcinogen metabolism. J. Nutr. 1999;129:768S-774S
25.
Heerdt B. G., Houston M. A., Augenlicht L. H. Potentiation of specific short-chain fatty acids of differentiation and apoptosis in human colonic carcinoma cells. Cancer Res 1994;54:3288-3293
26. Jones D. P., Kirlin W. G., Patten E., DeLong M. J. Redox responses preceding enzyme induction and apoptosis in human colon HT29 cells. The Toxicologist 1996;30:1605
27.
Joseph P., Jaiswal A. K. NAD(P)H:quinone reductase (DT diaphorase) specifically prevents the formation of benzo[a]pyrene quinone-DNA adducts generated by cytochrome P4501A1 and P450 reductase. Proc. Natl. Acad. Sci. USA 1994;91:8413-8417
28. Kapadia S. A., Raimundo A. H., Grimble G. K., Aimer P., Silk D. B. Influence of three different fiber-supplemented enteral diets on bowel function and short-chain fatty acid production. J. Parenter. Ent. Nutr. 1995;19:63-68
29.
Kashtan H., Stern H. S., Jenkins D. J., Jenkins A. L., Thompson L. U., Hay K., Marcon N., Minkin S., Bruce W. R. Colonic fermentation and markers of colorectal-cancer risk. Am. J. Clin. Nutr. 1992;55:723-728
30. Kune G. S., Kune S. The nutritional causes of colorectal cancer: an introduction to the Melbourne study. Nutr. Cancer 1987;9:1-4
31. Kune S., Kune G. S., Watson L. F. Case-control study of dietary etiological factors: the Melbourne colorectal cancer study. Nutr. Cancer 1987;9:21-42[Medline]
32.
Lin D., Meyer D. J., Ketterer B., Lang N. P., Kadlubar F. F. Effects of human and rat glutathione S-transferases on the covalent DNA binding of the N-acetoxy derivatives of heterocyclic amine carcinogens in vitro: a possible mechanism of organ specificity in their carcinogenesis. Cancer Res 1994;54:4920-4926
33. McConkey D. J., Hartzell P., Nicotera P., Orrenius S. Calcium-activated DNA fragmentation kills immature thymocytes. FASEB J 1989;3:1843-1849[Abstract]
34. Miller A. B., Howe G. R., Jain M., Craib K.J.P., Harrison L. Food items and food groups as risk factors in a case-control study of diet and colo-rectal cancer. Int. J. Cancer 1983;32:155-161[Medline]
35. National Research Council Diet and Health 1989 National Academy Press Washington, DC
36. Nicoletti I., Migliorati G., Pagliacci M. C., Grignani F., Riccardi C. A rapid and simple method for measuring thymocyte apoptosis by propidium iodide staining and flow cytometry. J. Immunol. Methods 1991;139:271-279[Medline]
37. Ohgaki H., Takayama S., Sugimura T. Carcinogenicity of heterocyclic amines in cooked food. Mutat. Res. 1991;259:399-410[Medline]
38. Potten C. S. The significance of spontaneous and induced apoptosis in the gastrointestinal tract of mice. Cancer Metast Rev 1992;11:179-195[Medline]
39. Potter J. D. Colon cancer—do the nutritional epidemiology, the gut physiology and the molecular biology tell the same story?. J. Nutr. 1993;123(Suppl 2):418-423
40.
Prestera T., Holtzclaw W. D., Zhang Y., Talalay P. Chemical and molecular regulation of enzymes that detoxify carcinogens. Proc. Natl. Acad. Sci. USA 1993;90:2965-2969
41.
Prochaska H. J., Santamaria A. B., Talalay P. Rapid detection of inducers of enzymes that protect against carcinogens. Proc. Natl. Acad. Sci. USA 1992;89:2394-2398
42.
Ranganathan S., Tew K. D. Immunohistochemical localization of glutathione S-transferases , µ, and 
in normal tissue and carcinomas from human colon. Carcinogenesis 1991;12:2383-2387
43. Rao C. V., Nayini J., Reddy B. S. Effect of Oltipraz [5-(2-Pyrazinyl)-4-methyl-1,2-dithiol-3-thione] on azomethane-induced biochemical changes related to early colon carcinogenesis in male F344 rats. Proc. Soc. Exp. Biol. Med. 1991;197:77-84[Medline]
44. Reddy B. S., Maeura Y. Dose-response studies of the effect of dietary butylated hydroxyanisole on colon carcinogenesis induced by methylazoxymethanol acetate in female CF1 mice. J. Nat. Cancer Inst. 1984;72:1181-1187
45.
Reddy B. S., Sugie S., Maruyama H., el-Bayoumy K., Marra P. Chemoprevention of colon cancer by dietary organoselenium benzylselenocyanate in F344 rats. Cancer Res 1987;47:5901-5904
46. Samali A., Zhitovosky B., Jones D. P., Nagata S., Orrenius S. . Cell Death Diff. 1999;6:495-496[Medline]
47. Slater A. F., Stefan C., Nobel I., van den Dobbelsteen D. J., Orrenius S. Signalling mechanisms and oxidative stress in apoptosis. Toxicol. Lett. 1995;82–83:149-153
48.
Sparins V. L., Chuan J., Wattenberg L. W. Enhancement of glutathione S-transferase activity of the esophagus by phenols, lactones, and benzyl isothiocyanate. Cancer Res 1982;42:1205-1207
49. Spencer S. R., Xue L., Klenz E. M., Talalay P. The potency of inducers of NAD(P)H:(quinone-acceptor) oxidoreductase parallels their efficiency as substrates for glutathione S-transferases. Biochem. J. 1991;273:711-717
50. Steinmetz K. A., Potter J. D. Vegetables, fruits and cancer. II. Mechanisms. Cancer Causes and Control 1991;2:427-442[Medline]
51. Stridh H., Kimland M., Jones D. P., Orrenius S., Hampton M. B. Cytochrome c release and caspase activation in hydrogen peroxide- and tributyltin-induced apoptosis. FEBS Lett 1998;429:351-355[Medline]
52.
Sumiyoshi H., Wargovich M. J. Chemoprevention of 1,2-dimethylhydrazine-induced colon cancer in mice by naturally occurring organosulfur compounds. Cancer Res 1990;50:5084-5087
53.
Talalay P., DeLong M. J., Prochaska H. J. Identification of a common signal regulating the induction of enzymes that protect against chemical carcinogenesis. Proc. Natl. Acad. Sci. USA 1988;85:8261-8265
54.
Thompson C. B. Apoptosis in the pathogenesis and treatment of disease. Science 1995;267:1456-1462
55.
Thornberry N., Lazebnik Y. Caspases: enemies within. Science 1998;281:1312-1316
56.
Vogelstein B., Kinzler K. W. Colorectal cancer and the intersection between basic and clinical research. Cold Spring Harbor Symp. Quant. Biol. 1994;59:517-521
57. Vos R.M.E., Snoek M. C., van Berkel W.J.H., Müller F. Differential induction of rat hepatic glutathione S-transferase isoenzymes by hexachlorobenzene and benzyl isothiocyanate. Biochem. Pharmacol. 1988;37:1077-1082[Medline]
58.
Wattenberg L. W. Inhibition of carcinogenesis by minor dietary constituents. Cancer Res 1992;52(Suppl.):2085s-2091s
59. Wyllie A. H., Kerr J.F.R., Currie A. R. Cell death: the significance of apoptosis. Int. Rev. Cytol. 1980;68:251-307[Medline]
60. Young T. B., Wolf D. A. Case-control study of proximal and distal colon cancer and diet in Wisconsin. Int. J. Cancer 1988;42:167-175[Medline]
This article has been cited by other articles:
|
|
 |
|
  D. P. Jones Radical-free biology of oxidative stress Am J Physiol Cell Physiol, October 1, 2008; 295(4): C849 - C868. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 S. N. T. Ngo, D. B. Williams, L. Cobiac, and R. J. Head Does Garlic Reduce Risk of Colorectal Cancer? A Systematic Review J. Nutr., October 1, 2007; 137(10): 2264 - 2269. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 C. Steiner, W. H.M. Peters, E. P. Gallagher, P. Magee, I. Rowland, and B. L. Pool-Zobel Genistein protects human mammary epithelial cells from benzo(a)pyrene-7,8-dihydrodiol-9,10-epoxide and 4-hydroxy-2-nonenal genotoxicity by modulating the glutathione/glutathione S-transferase system Carcinogenesis, March 1, 2007; 28(3): 738 - 748. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. J. Aleman, M. P. DeYoung, M. Tress, P. Keating, G. W. Perry, and R. Narayanan Inhibition of Single Minded 2 gene expression mediates tumor-selective apoptosis and differentiation in human colon cancer cells PNAS, September 6, 2005; 102(36): 12765 - 12770. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 R. Hu, V. Hebbar, B.-R. Kim, C. Chen, B. Winnik, B. Buckley, P. Soteropoulos, P. Tolias, R. P. Hart, and A.-N. T. Kong In Vivo Pharmacokinetics and Regulation of Gene Expression Profiles by Isothiocyanate Sulforaphane in the Rat J. Pharmacol. Exp. Ther., July 1, 2004; 310(1): 263 - 271. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J. Cummings, B. T. Ethell, L. Jardine, G. Boyd, J. S. Macpherson, B. Burchell, J. F. Smyth, and D. I. Jodrell Glucuronidation as a Mechanism of Intrinsic Drug Resistance in Human Colon Cancer: Reversal of Resistance by Food Additives Cancer Res., December 1, 2003; 63(23): 8443 - 8450. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
M. N. Ebert, A. Klinder, W. H.M. Peters, A. Schaferhenrich, W. Sendt, J. Scheele, and B. L. Pool-Zobel Expression of glutathione S-transferases (GSTs) in human colon cells and inducibility of GSTM2 by butyrate Carcinogenesis, October 1, 2003; 24(10): 1637 - 1644. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
L. M. Knowles and J. A. Milner Diallyl Disulfide Induces ERK Phosphorylation and Alters Gene Expression Profiles in Human Colon Tumor Cells J. Nutr., September 1, 2003; 133(9): 2901 - 2906. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. H. Fowke, F.-L. Chung, F. Jin, D. Qi, Q. Cai, C. Conaway, J.-R. Cheng, X.-O. Shu, Y.-T. Gao, and W. Zheng Urinary Isothiocyanate Levels, Brassica, and Human Breast Cancer Cancer Res., July 15, 2003; 63(14): 3980 - 3986. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. Seow, J.-M. Yuan, C.-L. Sun, D. Van Den Berg, H.-P. Lee, and M. C. Yu Dietary isothiocyanates, glutathione S-transferase polymorphisms and colorectal cancer risk in the Singapore Chinese Health Study Carcinogenesis, December 1, 2002; 23(12): 2055 - 2061. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 K. C. Nelson, J. S. Armstrong, S. Moriarty, J. Cai, M.-W. H. Wu, P. Sternberg Jr, and D. P. Jones Protection of Retinal Pigment Epithelial Cells from Oxidative Damage by Oltipraz, a Cancer Chemopreventive Agent Invest. Ophthalmol. Vis. Sci., November 1, 2002; 43(11): 3550 - 3554. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K. R.K. Sticha, P. M.J. Kenney, G. Boysen, H. Liang, X. Su, M. Wang, P. Upadhyaya, and S. S. Hecht Effects of benzyl isothiocyanate and phenethyl isothiocyanate on DNA adduct formation by a mixture of benzo[a]pyrene and 4-(methylnitrosamino)-1-(3-pyridyl)-1-butanone in A/J mouse lung Carcinogenesis, September 1, 2002; 23(9): 1433 - 1439. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. S. Hecht, P. Upadhyaya, M. Wang, R. L. Bliss, E. J. McIntee, and P. M.J. Kenney Inhibition of lung tumorigenesis in A/J mice by N-acetyl-S-(N-2-phenethylthiocarbamoyl)-L-cysteine and myo-inositol, individually and in combination Carcinogenesis, September 1, 2002; 23(9): 1455 - 1461. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
C. Bonnesen, I. M. Eggleston, and J. D. Hayes Dietary Indoles and Isothiocyanates That Are Generated from Cruciferous Vegetables Can Both Stimulate Apoptosis and Confer Protection against DNA Damage in Human Colon Cell Lines Cancer Res., August 1, 2001; 61(16): 6120 - 6130. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 R. Yu, S. Mandlekar, and A.-N. T. Kong Molecular Mechanisms of Butylated Hydroxylanisole-Induced Toxicity: Induction of Apoptosis through Direct Release of Cytochrome c Mol. Pharmacol., August 1, 2000; 58(2): 431 - 437. [Abstract] [Full Text]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
