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Interaction of bracken-fern extract with vitamin C in human

Posted Feb 02 2010 4:39am

Bracken-fern (Pteridium aquilinum) can induce acute and
chronic toxicity in man and farm animals [1–3]. In cattle, chronic
bracken-fern toxicity causes carcinomas in urinary bladder [4]
in bovine presenting enzootic hematuria, which is a relatively
common condition in bracken-infested pasturelands [5]. Epidemiological
studies have related the occurrence of neoplasms
in humans to the consumption of bracken crosiers [2,6] or
milk from cows feeding on bracken [7,8]. It has been also

suggested that the consumption of water contaminated with
bracken-derived substances can be toxic [9].
Long-term exposure to bracken produces ionizing-radiationlike
effects [10,11]. Radiomimetic compounds are associated to
the generation of free radicals [12], which would be the primary
effectors of DNA damage and chromosome aberrations
induced by the plant. Moreover, a high incidence of structural
chromosome aberrations was verified in human peripheral lymphocytes
of bracken-fern consumers [13], in lymphocytes of
animals grazing on bracken-fern [14], and in mice bone marrow
and peritoneal cells [15].
Considering that bracken-fern-induced DNA damage could
be related to free radicals generation, it is reasonable to assume
that antioxidant dietary constituents – such as vitamin C –
could revert such damage. The antioxidative effects of this vitamin
are mediated by the scavenging of reactive oxygen and

nitrogen species [16], and by regeneration of other small
molecule antioxidants, such as -tocopherol and glutathione
(GSH) [17]. Furthermore, vitamin C inhibits the mutagenic
activity of several carcinogenic substances [18]. Notwithstanding
the antioxidant activity, vitamin C is also known as a
pro-oxidant substance in vitro [19]. In this case, its interaction
with transitional metal ions is accompanied by H2O2 production
[17,20]. There are reports that vitamin C heightens the cytotoxic
effects of chemotherapeutic drugs, such as arsenic trioxide,
through free radical generation [21].

A correlation between bracken-fern consumption and oral
cancer has not yet been reported. Nonetheless, since the mouth
is the first body compartment exposed to the plant, and head
and neck cancers are the sixth leading cause of cancer mortality
[22], it is important to study cell lines that can contribute
to the understanding of the genetic damages possibly involved
in the etiology of such neoplasies. Considering the association
between genetic alterations and cancer, and the occurrence of
neoplasies related to bracken intake, the aim of the current study
was to evaluate the effects of vitamin C on oral cavity-derived
cells (oral epithelium – OSCC-3 – and human submandibular
gland, HSG) presenting DNA damage induced by bracken-fern
extract. OSCC cells have been used in carcinogenesis mechanisms,
antitumoral treatments and cytotoxicity studies [23–25].
HSG cells have been employed in tissue engineering, cell differentiation,
apoptosis induction and cytotoxicity experiments
[26–29].
2. Materials and methods
2.1. Plant extract and reagents
Bracken-fern was collected in the savanna (cerrado) of Brasilia and identified
by botanists of the University of Brasilia, Brazil. The aqueous extract
was obtained as described before [15]. Briefly, fronds were washed and oven
dried at 37 â—¦C and ground in a Willey mill. The resulting powder was extracted
with distilled water—31 mg of extract for 1 ml. The extract was filtered and
sterilized through a 0.22- m nitrocellulose membrane and stored in cold and
dark until use. Bracken-fern extracts and vitamin C (ascorbic acid, CEVITON®,
ARISTON) were diluted in culture medium at the determined experimental concentrations.
Hydrogen peroxide (Sigma Chemical Co., St Louis, MO, USA) was
used as oxidizing agent. HSG cells and human oral epithelium cells (OSCC-3)
were a generous gift of Prof. B. J. Baum (National Institute of Health Bethesda,
USA).
2.2. Culture conditions
In order to test the effects of vitamin C on the reversibility of DNA damage
caused by bracken-fern, HSG and OSCC-3 cells were grown at 37 â—¦C, in an
atmosphere of 5% of CO2 in Dulbecco’s modified Eagle’s medium, DMEM,
(GIBCO-BRL), pH 7.4, supplemented with 10% fetal calf serum, 100 U/ml
penicillin and streptomycin (100 g/ml).
For comet and acridine orange/ethidium bromide staining assays, HSG and
OSCC-3 cells (5×104) were seeded in 25 cm2 tissue culture flasks and stabilized
for 24 h. For morphological analysis by light microscopy, 105cells were
grown over microscope coverslips placed in six-well culture plates. All experiments
were run in duplicate. The bracken-fern concentration used in the tests
was based on cell viability after the trypan blue exclusion test. Experimental
groups consisted of cells treated with bracken-fern (670 g/ml for HSG cells
and 1340 g/ml for OSCC cells), alone or in combination with vitamin C (10
and 100 g/ml), cultured for 48 h. Negative controls consisted of untreated cultures.
Hydrogen peroxide (1 mM), added to culture 4 h before cell processing,
was used as positive control. After cultivation, cells were washed, harvested,
re-suspended in phosphate buffer saline (PBS) and used for the tests.
2.3. Comet assay
We used the comet assay described by Singh et al. [30] with some modifications.
Briefly, 30 l of cell suspension (105–106 cells) were mixed with 120 l
of low melting point agarose (0.5% in PBS) and spread over microscopy slides
that had been previously covered with a layer of agarose type II (1.5% in PBS).
The slides were immersed in cold lysing solution (2.5M NaCl, 0.1M EDTA,
0.01M Tris and 1X Triton X-100) for 1 h at 4 â—¦C. The slides were then placed
in a denaturating electrophoresis buffer (300mM NaOH, pH 13, 1mM EDTA,)
and subject to 25V (300 mA) for 20 min. After electrophoresis, the samples
were neutralized in 0.4M Tris buffer (pH 7.5) and stained with 50 l ethidium
bromide (20 g/ml; Sigma).
Approximately 200 cells (100 per replicate slide) per experimental group
were randomly analyzed using a fluorescence microscope (Axioplan-Carl Zeiss)
at 400×, with a 515 to 560-nm exciting filter and a 590-nm barrier filter. The
category of DNA damage was assigned to five classes (0–4) based on the visual
aspect of the comets (nucleoids), considering the extent of DNA migration,
according to the criteria established by Visvardis et al. [31]. Nucleoids with a
bright head and no tail were classified as class 0 (undamaged nucleoids) and
nucleoids with a small head and long diffuse tails, as class 4, i.e. as highly damaged
nucleoids. Nucleoids with intermediate characteristics were classified as
classes 1, 2 or 3. The index of DNA damage (IDD) was estimated quantitatively
by the equation described by Jalonszynski et al. [32];
IDD = (n1 + 2n2 + 3n3 + 4n4)
(Σ/100)
where IDD is the DNA damage in arbitrary units, n1–n4 the number of classes
1–4, and Σ is the total number of scored nucleoids, including class 0. IDD
values ranged from 0 (all undamaged nucleoids) to 400 (all maximally damaged
nucleoids). Arbitrary units were converted to the percentage of damaged
DNA (frequency of DNA damage), with the score of 400 as 100% damaged
DNA.
2.4. Detection of apoptosis and necrosis

In order to assess the frequency of cell death mechanism (apoptotic or
necrotic), cells were collected, centrifuged and re-suspended in PBS. Twenty
microliters of the cell suspension (105–106 cells) were stained with a fresh
solution of acridine orange (100 g/ml)/ethidium bromide (100 g/ml)—1:1 for
5 min. Cells were then examined using a fluorescence microscope (Olympus;
barrier filter O 530 NM). Apoptosis or necrosis and cell survival were evaluated
by visual analysis according to previously established criteria [33,34].
Acridine orange penetrates into living and dead cells, emitting green fluorescence
as a result of intercalation into double-stranded DNA, and red–orange
fluorescence after binding with single-stranded RNA. Ethidium bromide emits
red fluorescence after intercalation in DNA of cells with altered cell membrane
(at necrosis or at a late stage of apoptosis). Thus, considering that apoptotic
cells show fragmented nucleus and condensed chromatin, four cell stages can
be identified by this assay (Fig. 1): (i) living cells, with uniform green nucleus
(Fig. 1a), (ii) early apoptosis (cell membrane is still preserved but chromatin
condensation and an irregular green nucleus are visible) (Fig. 1b), (iii) late
apoptosis (ethidium bromide penetrates through altered cell membrane and
stains the nuclei in red, while fragmentation or condensation of chromatin is
still observed) (Fig. 1c and d), and (iv) necrosis (uniformly red-stained cell
nuclei) (Fig. 1e).

2.3. Comet assay
We used the comet assay described by Singh et al. [30] with some modifications.
Briefly, 30 l of cell suspension (105–106 cells) were mixed with 120 l
of low melting point agarose (0.5% in PBS) and spread over microscopy slides
that had been previously covered with a layer of agarose type II (1.5% in PBS).
The slides were immersed in cold lysing solution (2.5M NaCl, 0.1M EDTA,
0.01M Tris and 1X Triton X-100) for 1 h at 4 â—¦C. The slides were then placed
in a denaturating electrophoresis buffer (300mM NaOH, pH 13, 1mM EDTA,)
and subject to 25V (300 mA) for 20 min. After electrophoresis, the samples
were neutralized in 0.4M Tris buffer (pH 7.5) and stained with 50 l ethidium
bromide (20 g/ml; Sigma).
Approximately 200 cells (100 per replicate slide) per experimental group
were randomly analyzed using a fluorescence microscope (Axioplan-Carl Zeiss)
at 400×, with a 515 to 560-nm exciting filter and a 590-nm barrier filter. The
category of DNA damage was assigned to five classes (0–4) based on the visual
aspect of the comets (nucleoids), considering the extent of DNA migration,
according to the criteria established by Visvardis et al. [31]. Nucleoids with a
bright head and no tail were classified as class 0 (undamaged nucleoids) and
nucleoids with a small head and long diffuse tails, as class 4, i.e. as highly damaged
nucleoids. Nucleoids with intermediate characteristics were classified as
classes 1, 2 or 3. The index of DNA damage (IDD) was estimated quantitatively
by the equation described by Jalonszynski et al. [32];
IDD = (n1 + 2n2 + 3n3 + 4n4)
(Σ/100)
where IDD is the DNA damage in arbitrary units, n1–n4 the number of classes
1–4, and Σ is the total number of scored nucleoids, including class 0. IDD
values ranged from 0 (all undamaged nucleoids) to 400 (all maximally damaged
nucleoids). Arbitrary units were converted to the percentage of damaged
DNA (frequency of DNA damage), with the score of 400 as 100% damaged
DNA.
2.4. Detection of apoptosis and necrosis
In order to assess the frequency of cell death mechanism (apoptotic or
necrotic), cells were collected, centrifuged and re-suspended in PBS. Twenty
microliters of the cell suspension (105–106 cells) were stained with a fresh
solution of acridine orange (100 g/ml)/ethidium bromide (100 g/ml)—1:1 for
5 min. Cells were then examined using a fluorescence microscope (Olympus;
barrier filter O 530 NM). Apoptosis or necrosis and cell survival were evaluated
by visual analysis according to previously established criteria [33,34].
Acridine orange penetrates into living and dead cells, emitting green fluorescence
as a result of intercalation into double-stranded DNA, and red–orange
fluorescence after binding with single-stranded RNA. Ethidium bromide emits
red fluorescence after intercalation in DNA of cells with altered cell membrane
(at necrosis or at a late stage of apoptosis). Thus, considering that apoptotic
cells show fragmented nucleus and condensed chromatin, four cell stages can
be identified by this assay (Fig. 1): (i) living cells, with uniform green nucleus
(Fig. 1a), (ii) early apoptosis (cell membrane is still preserved but chromatin
condensation and an irregular green nucleus are visible) (Fig. 1b), (iii) late
apoptosis (ethidium bromide penetrates through altered cell membrane and
stains the nuclei in red, while fragmentation or condensation of chromatin is
still observed) (Fig. 1c and d), and (iv) necrosis (uniformly red-stained cell
nuclei) (Fig. 1e).
2.5. Statistical analysis
The statistical analysis of the acridine orange/ethidium bromide doublestaining
data was based on cells/group after arcsine
√
x transformation
(x = number of death cells). The comet assay data was analyzed considering
two parameters: (i) frequency of cells with damaged DNA; (ii) IDD. Statistical
analysis of the data was based on cells/group after arcsine
√
x transformation
(x = number of cells withDNAdamage) or log x transformation (x = IDD). Trans

 3. Results
Results ofDNAdamage analysis are shown in Fig. 2.Vitamin
C alone did not reduce DNA damage caused by bracken-fern in

HSG and OSCC-3 cells. However, at a higher concentration
(100 g/ml), vitamin C alone induced DNA damage in both cell
lines. Moreover, bracken-fern extract together with vitamin C
showed a synergistic effect on the frequency of HSG cells with
DNA damage.
We have also evaluated if the bracken-fern extract alters cell
morphology (Fig. 3). Fig. 3a illustrates the similar morphology
of control HSG and OSCC-3 cells. Both cell lines show
polygonal form that, depending on the cell-culture confluence,
can be slightly elongated or spherical; cytoplasmic projections
can bind one cell to another. The cell nuclei are spherical or
ovoid, with fine and slightly granular chromatin showing one or
more nucleoli. Cytoplasm has a relatively homogeneous appearance,
presenting some cytoplasmic densities. When cells were
treated with vitamin C (10 g/ml) alone, there were no differential
morphological features compared to the control. On the
other hand, bracken-fern extract-treated cells showed apoptosisrelated
morphological features, such as chromatin condensation
and cytoplasmic volume loss (Fig. 3b), presence of vacuoles
(Fig. 3c) and changes in the membrane symmetry.
The quantification of cell death, distinguishing the necrotic
from the apoptotic cells, was performed by the acridine
orange/ethidium bromide double-staining assay (Fig. 4). Both
bracken-fern extract and vitamin C were cytotoxic to HSG and
OSCC-3 cells, either alone or in combination. According to this
assay (see Section 2), cell death was caused by apoptosis in all
treatments.

 

 

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