carc$$0202
Carcinogenesis vol.18 no.2 pp.271–277, 1997
Oxidative DNA base damage and its repair in kidneys and livers
of nickel(II)-treated male F344 rats
Kazimierz S.Kasprzak1,5, Pawel Jaruga2,
Tomasz H.Zastawny3, S.Lynn North1, Charles W.Riggs4,
Ryszard Olinski3 and Miral Dizdaroglu2
1Laboratory
of Comparative Carcinogenesis, National Cancer Institute,
FCRDC, Frederick, MD 21702, 2Chemical Science and Technology
Laboratory, National Institute of Standards and Technology, Gaithersburg,
MD 20899, USA, 3Department of Clinial Biochemistry, L.Rydygier Medical
School, Bydgoszcz, Poland and 4Data Management Services, Inc., FCRDC,
Frederick, MD 21702, USA
5To
whom correspondence should be addressed
DNA base damage was assayed using gas chromatography/
mass spectrometry with selected ion monitoring (GC/MSSIM) in renal and hepatic chromatin of male F344 rats up
to 14 days after a single i.p. injection of 90 µmol Ni(II)
acetate/kg body wt. Ten different damaged bases were
quantified. No damage was found in either organ 12 h
after Ni(II) treatment. The damage became significant only
from day 1, with magnitude and persistence depending on
the organ and base. In livers, levels of five DNA base
products were significantly elevated over those in control
rats. They were: 8-oxoguanine (by 46% at day 1 postinjection); 2,6-diamino-4-hydroxy-5-formamidopyrimidine
(by 107% at day 1); 5-(hydroxymethyl)uracil (by 94% at
day 1); 5,6-dihydroxyuracil (by 128% at day 1); and 5hydroxyhydantoin (by 39% in terms of the overall adjusted
means for days 1–14 post-injection). The elevation was
highest at day 1 post-injection followed by a decrease at
later days, except for 5-hydroxyhydantoin. In kidneys,
the levels of only three damaged bases, 8-oxoguanine, 5hydroxyhydantoin and 5,6-dihydroxyuracil were increased
significantly (by 31, 73 and 60%, respectively) and one
base, 8-oxoadenine, was increased by 26%, just below
significance, all in terms of overall adjusted means for days
1–14 post-injection. Hence, unlike those in the liver, the
renal increases persisted for 14 days. The results reveal a
tissue specific response to Ni(II)-mediated oxidative DNA
base damage with apparently faster DNA repair in liver
than in kidney, the main target of Ni(II) carcinogenicity.
Introduction
Our previous investigations revealed that intraperitoneal (i.p.)
injection of a soluble Ni(II) acetate to Fischer rats initiated
renal tumors promotable by oral treatment with sodium barbital,
a multi-tissue tumor promoter (1). Likewise, Ni(II) acetate
injected i.p. to pregnant Fischer rats initiated renal tumors in
*Abbreviations: BSTFA, bis(trimethylsilyl)trifluoroacetamide; GC/MS-SIM,
gas chromatography/mass spectrometry with selected ion monitoring; 5OHMe-Ura, 5-(hydroxymethyl)uracil; 5-OH-5-Me-Hyd, 5-hydroxy-5-methylhydantoin; 5-OH-Ura, 5-hydroxyuracil; 5-OH-Cyt, 5-hydroxycytosine; 5,6diOH-Ura, 5,6-dihydroxyuracil; 5-OH-Hyd, 5-hydroxyhydantoin; 8-oxo-Ade,
7,8-dihydro-8-oxoadenine; FapyAde, 4,6-diamino-5-formamidopyrimidine; 8boxo-Gua, 7,8-dihydro-8-oxoguanine; FapyGua, 2,6-diamino-4-hydroxy-5-formamidopyrimidine.
© Oxford University Press
the offspring (2). In both cases the treatment with sodium
barbital started at least 2 weeks after the nickel injection, i.e.
at a time when the nickel dose was essentially excreted (3,4).
We also found that Ni(II) treatment like the above resulted
in oxidative DNA base damage in the rat kidneys and livers
1–2 days after the injection (1,2,5). Because some of the
damaged bases had been already known to be promutagenic,
we speculated that their generation might be involved in the
mechanisms of carcinogenesis by nickel and, presumably, other
transition metals (6). However, in spite of causing damage in
both the kidney and liver, Ni(II) acetate initiated carcinogenesis
only in the kidney (1). The reasons for this difference remained
unclear. Some of them would, perhaps, be related to different
spectra of the damage and different capacity of repair in these
organs. To answer these questions, the present study was
designed to look at the spectrum and repair of oxidative DNA
base damage in the kidneys and livers of nickel-treated
rats up to 14 days post-injection. The DNA base products
determined and quantified in this study are presented in
Figure 1.
Materials and methods
Materials
Nickel(II) acetate tetrahydrate, sodium acetate trihydrate, sodium and potassium phosphates, hydrochloric acid and other inorganic chemicals were
purchased from J.T.Baker Chemical Company (Phillipsburg, NJ). Formic
acid (88%) was obtained from Malinckrodt (Paris, KY). Acetonitrile and
bis(trimethylsilyl)trifluoroacetamide (BSTFA*) containing 1% trimethylchlorosilane were obtained from the Pierce Chemical Company (Rockford, IL).
Tris, EDTA, Chelex-100 resin (200–400 mesh), Triton X-100, sucrose,
phenylmethane-sulfonyl fluoride, dithiothreitol, isobarbituric acid (5-hydroxyuracil; 5-OH-Ura) and 5-(hydroxymethyl)uracil (5-OHMe-Ura) were purchased from Sigma Chemical Co. (St Louis, MO). Thymine-α,α,α,6-2H4 was
obtained from Merck and Co., Inc./Isotopes (Montreal, Canada). 5-Hydroxy5-methylhydantoin (5-OH-5-Me-Hyd), dialuric acid, isodialuric acid [5,6dihydroxyuracil (5,6-diOH-Ura)], 5-hydroxycytosine (5-OH-Cyt), 4,6-diamino-5-formamidopyrimidine (FapyAde), 7,8-dihydro-8-oxoadenine (8-oxoAde), 2,6-diamino-4-hydroxy-5-formamidopyrimidine (FapyGua), 7,8-dihydro-8-oxoguanine (8-oxo-Gua), 5-hydroxy-5-methylhydantoin-1,3-15N2-2-13C,
dialuric acid-1,3-15N2-2,4-13C2, 5-hydroxyuracil-1,3-15N2-2-13C, 5-(hydroxymethyl)uracil-2,4-13C2-α,α-2H2, 5-hydroxycytosine-1,3-15N2-2-13C, 5,6- dihydroxyuracil-1,3-15N2-2-13C, 4,6-diamino-5-formamidopyrimidine-1,3-15N-213C-(5-aminoformyl-15N,2H), 7,8-dihydro-8-oxoadenine-1,3,7-15N -2,8-13C ,
3
2
2,6-diamino-4-hydroxy-5- formamidopyrimidine-1,3-15N2-(5-aminoformyl-15N)2-13C, and 7,8-dihydro-8-oxoguanine-1,3-15N2-(2-amino-15N)-2-13C were synthesized by Dr V.Nelson from SAIC Frederick (Frederick, MD).
Animals and treatments
Forty-eight male F344/NCr rats, 5 weeks old, obtained from the Animal
Production Area (NCI-FCRDC, Frederick, MD) were housed in polycarbonate
cages on a sawdust bedding (Sanichips, P.G.Murphy Forest Products, Co.,
Mountsville, NJ). They had free access to food (NIH-31 Open Formula 6%
Modified, Zeigler Brothers, Gardners, PA) and drinking water. The rats were
divided randomly into two groups of 24 rats. One group of rats received a
single i.p. injection of 180 µmol of sodium acetate/kg body wt (control group).
Rats from the treatment group received a single i.p. injection of 90 µmol of
Ni(II) acetate/kg body wt. The salts were dissolved in water and injected in
aliquots of 2 ml/kg body wt. The rats were killed by exsanguination (cardiac
puncture) under nitrogen narcosis 12 h and 1, 3, 7 and 14 days post-injection
(eight rats per group at day 1, four rats at each other time). Their kidneys and
livers were quickly removed and stored in liquid nitrogen until analysis.
271
K.S.Kasprzak et al.
Fig. 1. Oxidatively damaged DNA bases determined by the GC/MS-SIM according to Dizdaroglu (12).
Chromatin preparation
Nuclei were isolated according to Lilja et al. (7) with some modifications
introduced by Tsapakos et al. (8). Chromatin was prepared from isolated
nuclei according to Mee and Adelstein (9). Both procedures have been
described in more detail previously (2).
Hydrolysis and derivatization
To chromatin samples containing 100 µg of DNA (as predetermined by UV
measurements), 2 nmol of thymine-α,α,α,6-2H4 and 0.5 nmol each of 5hydroxy-5-methylhydantoin-1,3-15N2-2-13C, dialuric acid-1,3-15N2-2,4-13C2,
5-hydroxyuracil-1,3-15N2-2-13C, 5-(hydroxymethyl)uracil-2,4-13C2-α,α-2H2,
5-hydroxycytosine-1,3-15N2-2-13C, 5,6-dihydroxyuracil-1,3-15N2-2-13C, 4,6diamino-5-formamidopyrimidine-1,3-15N-2-13C-(5-aminoformyl-15N,2H), 7,8dihydro-8-oxo-adenine-1,3,7-15N3-2,8-13C2, 2,6-diamino-4-hydroxy-5-formamidopyrimidine-1,3-15N2-(5-aminoformyl-15N)-2-13C and 7,8-dihydro-8oxoguanine-1,3-15N2-(2-amino-15N)-2-13C were added as internal standards.
Samples were lyophilized and hydrolyzed with 0.5 ml of 60% formic acid in
evacuated and sealed glass tubes for 30 min at 140°C (10). Under these
hydrolysis conditions, dialuric acid-1,3-15N2-2,4-13C2 yields quantitatively 5hydroxyhydantoin-1,3-15N2-2,4-13C2 (11). The hydrolysates were lyophilized
and then trimethylsilylated under nitrogen in polytetrafluoroethylene-capped
hypovials (Pierce Chemical Co., Rockford, IL) with 100 µl of a mixture of
nitrogen-bubbled BSTFA and acetonitrile (4/1, v/v) by heating for 30 min
at 120°C.
Gas chromatography–mass spectrometry with selected ion monitoring (GC/
MS-SIM)
Analyses of the derivatized hydrolysates of chromatin samples by GC/MS
with selected ion monitoring (SIM) were performed as previously described
(10–12). An aliquot of derivatized sample was injected into the injection port
of a gas chromatograph using an autosampler. A split ratio of 1:15 was used
at the injection port. This resulted in approximately 0.25 µg of hydrolyzed
and derivatized DNA going through the GC column during each analysis and
a detection limit of approximately 3 pmol of base product/mg DNA. The
quantification of DNA base products was performed by isotope dilution mass
spectrometry using their stable isotope-labeled analogues as internal standards
(11). Thymine-α,α,α,6-2H4 was used to determine the amount of thymine and
DNA in hydrolyzed samples (13). The results of DNA determination by this
method and by UV measurements (above) correlated well with each other.
272
Statistical evaluation of the results
Testing for the DNA damage and repair involved regression analysis, with
time as the independent variable and the level of a particular damaged DNA
base as the dependent variable. Of the 10 damaged bases assayed, only six
bases whose levels were apparently affected by nickel in at least one organ
were fully statistically evaluated. For each of the resulting 12 organ/base
combinations, simple linear regressions were run separately for the control
and nickel-treated rats between days 1 and 14. The 12 h point was not
included because no DNA base damage was observed at this time (Table I).
The regression analyses were supplemented by the non-parametric Jonckheere
test for trend (14) in the event that the required assumption of homogeneous
within-cell variances was not always met. A statistically significant negative
regression slope for Ni(II)-treated animals was taken as evidence of DNA
base damage followed by repair over the next 13 days.
In cases in which the slopes for neither the control nor the treated rats
differed significantly from zero, the data were compared by covariance analysis
to confirm that the slopes did not differ from each other and to reveal possible
significant differences in levels of the overall (all time points) adjusted means
between the control and treated rats (15).
A significant difference in adjusted means by covariance analysis was
interpreted as evidence of DNA damage with no repair in 13 days.
Results
The parenteral administration of Ni(II) acetate resulted in
oxidative DNA base damage in the kidneys and livers of male
Fischer rats. Of the 10 damaged bases quantified (see Figure
1 for structures and names), the levels of four bases were
apparently increased in the kidney (8-oxo-Gua, 8-oxo-Ade, 5OH-Hyd, and 5,6-diOH-Ura), and levels of five bases were
increased in the liver (8-oxo-Gua, FapyGua, 5-OHMe-Ura, 5OH-Hyd, and 5,6-diOH-Ura). The statistical analysis that
followed confirmed the significance of those increases for
most of the base products at either day 1 post-injection or for
the overall adjusted means (days 1–14), but left some doubts
DNA base damage and repair in nickel-treated rats
Table I. Levels of DNA base products in kidney and liver chromatin 12 h and 1 day after treatmenta (mol/105 mol of parental base 6 SE, n 5 4–8)
12 h
DNA base product
5-OHMe-Ura
5-OH-5-Me-Hyd
5-OH-Ura
5-OH-Cyt
5,6-diOH-Ura
5-OH-Hyd
8-oxo-Ade
FapyAde
8-oxo-Gua
FapyGua
1 day
Kidney
C
Ni
C
Ni
C
Ni
C
Ni
C
Ni
C
Ni
C
Ni
C
Ni
C
Ni
C
Ni
4.35
4.29
29.79
31.32
5.19
5.79
27.38
28.96
5.82
7.48
10.81
12.87
5.22
7.32
7.12
8.28
13.44
16.23
12.10
13.97
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
Liver
1.82
1.40
7.95
9.77
1.32
1.53
4.73
3.76
1.05
1.12
1.80
2.26
0.60
0.70
1.99
0.92
3.58
0.55
2.62
1.91
5.66
6.19
44.91
40.53
14.43
10.84
6.52
5.82
9.45
8.81
59.00
69.20
5.75
6.15
9.80
8.46
8.31
8.51
4.88
4.26
Kidney
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
0.64
0.19
2.12
7.95
1.40
1.74
0.32
1.27
1.40
0.92
6.92
7.34
0.69
0.44
1.07
0.68
1.04
0.89
0.25
0.20
2.22
2.93
29.59
36.08
4.09
5.69
30.19
31.32
5.95
8.98
9.24
16.31
7.64
10.45
8.45
6.98
12.18
17.69
14.23
25.13
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
Liver
0.39
0.38
4.80
12.7
0.60
1.67
6.23
7.70
0.70
1.49b
0.54
2.06c
0.52
1.52b
1.83
1.35
1.22
2.05c
1.34
8.11
5.66 6 0.04
11.01 6 0.76d
56.35 6 9.63
76.28 6 3.53
14.60 6 1.46
29.08 6 1.55e
7.25 6 0.87
7.68 6 1.02
9.22 6 0.77
21.06 6 2.32d
59.90 6 23.0
86.50 6 5.40b
7.14 6 0.22
9.48 6 0.73e
12.71 6 1.29
12.90 6 0.12
10.20 6 0.98
14.95 6 1.52d
4.75 6 0.42
9.84 6 1.35d
aC,
control rats treated with sodium acetate; Ni, rats treated with Ni(II) acetate.
significant versus control by Student’s t-test at day 1, but significant by covariance analysis (P 5 0.04 or better) when pooled with the results for days 3,
7 and 14 versus pooled control values (see Figures 2–5).
cP , 0.05 or better by t-test versus control at day 1, also significant by covariance analysis (P 5 0.018 or better) when pooled with the results for days 3, 7
and 14 versus pooled control values.
dP , 0.05 or better by t-test versus control at day 1; not a subject for covariance analysis because of significant negative slope of the time curve for Nitreated rats compared with ’0’ slope for the control rats (see Figures 4–6).
eP , 0.05 or better by t-test versus the controls at day 1, but excluded from further consideration because of high experimental errors at later days.
bNot
about the increase of 8-oxo-Ade in the kidney (see below).
Table I gives the levels of all 10 DNA base products in
chromatin of the two organs 12 h and 1 day after treatment.
The statistically significant differences between control and
nickel-treated rats are indicated.
Kidney
The slopes of the time-dependent curves for 8-oxo-Gua, 8oxo-Ade, 5-OH-Hyd and 5,6-diOH-Ura in Ni(II) acetatetreated rats and the respective control rats were not significantly
different from zero by the regression and non-parametric tests
for trends (the P-values are given in Figures 2 and 3) and
from each other (treated/control for each base) by the covariance analysis (P 5 0.70, 0.39, 0.82, 0.86 for 8-oxo-Gua, 8oxo-Ade, 5-OH-Hyd and 5,6-diOH-Ura, respectively). Therefore, the adjusted means for each base level in the treated and
control rats could be compared by the covariance analysis.
The latter revealed highly significant increases by nickel
treatment in levels of three bases, with P 5 0.018 for 8-oxoGua (31% increase), 0.001 for 5-OH-Hyd (73%) and 0.002
for 5,6-diOH-Ura (60%). These results indicate a significant
oxidative damage to the guanine and cytosine residues of renal
DNA and no repair in 2 weeks. The adjusted means for 8oxo-Ade also differed between the two groups of rats with
P 5 0.043, but inspection of the raw data indicated that this
result depended largely on high 8-oxo-Ade levels in only a
few nickel-treated rats at days 1 and 3. Therefore, a positive
conclusion regarding oxidative damage to the adenine moiety
of renal DNA must be drawn with caution.
No significant differences were observed between nickeltreated and control rats for the remaining six DNA base
products quantified in renal DNA, including FapyGua, Fapy-
Ade, 5-OH-Cyt, 5-OH-Ura, 5-OHMe-Ura and 5-OH-5-MeHyd.
Liver
Following 128% (P , 0.008 by t-test), 107% (P , 0.03) and
94% (P , 0.0004) increases at day 1, the levels of 5,6diOH-Ura, FapyGua and 5-OHMe-Ura, respectively, decreased
markedly at later days. The slopes of the time-dependent
curves for these base products in nickel(II) acetate-treated rats
were significantly different from zero by both the regression
analysis and Jonckheere test. This, compared with ‘zero’ slopes
of the corresponding control curves (see Figures 4–6 for the
P-values) makes a convincing case for DNA base damage
(cytosine, thymine, and guanine in particular) followed by
repair. Although the slope of the 8-oxo-Gua curve in nickeltreated rats differed significantly from zero only by the
Jonckheere test, making a weak case for guanine damage with
repair, the inspection of the raw data supported such a
conclusion since the 8-oxo-Gua level was markedly elevated
by 46% at day 1 after treatment (P , 0.05 by t-test), and was
next quickly brought down to the control level (Figure 4). The
slopes of the 5-OH-Hyd curves for the treated and control rats
did not differ from zero (P-values given in Figure 5) and from
each other (P 5 0.45 by covariance analysis). That allowed
for using the covariance analysis to test statistical significance
of an apparent difference between the adjusted means of 5OH-Hyd levels in treated and control rats. Such analysis
yielded a positive result (39% increase with P 5 0.017). Thus,
the 5-OH-Hyd level was significantly increased by nickel,
indicating no repair of this damage in 2 weeks post-treatment.
No meaningful differences were found between nickeltreated and control rats for the remaining five DNA base
273
K.S.Kasprzak et al.
Fig. 2. Levels of 8-oxo-Gua and 8-oxo-Ade in renal DNA. Means 6 SE
(n 5 4–8). C, control rats (solid line); T, nickel-treated rats (dashed line).
PR and PJ, P-values of regression and Jonckheere analyses, respectively.
The P-values from the covariance analysis of adjusted means, indicating
significant difference between levels are 0.018 for 8-oxo-Gua and 0.043 for
8-oxo-Ade.
products quantified in the liver. Although levels of two of
them, 8-oxo-Ade and 5-OH-Ura, were significantly elevated
by Ni(II) acetate at day 1 (Table I), high experimental errors
at later days did not allow for drawing any firm conclusions
concerning the persistence or repair of these products. The
levels of FapyAde, 5-OH-Cyt, and 5-OH-5-Me-Hyd were
definitely not increased by Ni(II) treatment at any time point.
Discussion
To our best knowledge, the results of the present experiment
reveal for the first time the persistence of nickel(II)-induced
oxidative DNA base damage in different animal organs. Some
indications of a prolonged character of such damage after a
single nickel(II) injection have been observed by us before,
but only in the first 2 days after treatment (1,2). The present
results extend the original observations to a 14-day period of
time and allow for making comparisons in regard to the
spectrum and repair of oxidative DNA base damage in a target
(kidney) and non-target (liver) organ for nickel(II)-initiated
carcinogenesis.
Of the 10 damaged renal DNA bases quantified, the overall
adjusted means of three were significantly increased by nickel
treatment: 5-OH-Hyd by 73%, 5,6-diOH-Ura by 60% and 8oxo-Gua by 31%. The level of one more base, 8-oxo-Ade,
was increased by 26% over the control, at the edge of
significance. The first two bases, elevated to the highest extent,
originate from cytosine; an indication that the cytosine residues
274
Fig. 3. Levels of 5-OH-Hyd and 5,6-di-OH-Ura in renal DNA. The symbols
and descriptions are the same as in Figure 2. The P-values from the
covariance analysis of adjusted means are 0.001 for 5-OH-Hyd and 0.002
for 5,6-diOH-Ura.
were the main targets for nickel-mediated oxidative damage.
They were followed by the guanine and possibly adenine
residues. This pattern is consistent with the observation of
Kawanishi et al. (16) about the cytosines as the most sensitive
and adenines as the least sensitive sites of nickel/H2O2-induced
oxidative attack on DNA, resulting in DNA cleavage. It is,
however, surprising that significant damage was not observed
by us for the thymine residues, also found to be sensitive
cleavage sites (16). The renal DNA levels of neither 5-OHMeUra nor 5-OH-5-Me-Hyd, two thymine derivatives quantified
in the present experiment, were elevated by nickel.
Very importantly, the increased levels of 5-OH-Hyd, 5,6diOH-Ura and 8-oxo-Gua persisted in renal DNA up to 14
days post-injection, i.e. even when the nickel dose was almost
fully excreted (3,4,17). The reasons for this are not clear. The
most likely one seems to include inefficient repair of the
damage in the rat kidney. This, in turn, might be associated
with a kidney-specific low capacity of DNA repair relative to
other tissues or with nickel-induced inhibition of the repair
specifically in this organ. The first option seems unlikely, since
fast repair of the 8-oxo-Gua lesion was observed in rat kidneys
following administration of sodium bromate (18) or Fe(III)
nitrilotriacetate (19). The second possibility is supported by
the known inhibition of DNA repair by nickel(II) (20).
The DNA base damage in the hepatic DNA of nickel(II)
acetate-treated rats resulted in a significant increase of levels
of two cytosine derivatives, 5-OH-Hyd (by 39% in terms of
the adjusted means), 5,6-diOH-Ura (by 128% at day 1 postinjection), one thymine derivative, 5-OHMe-Ura (by 94% at
day 1) and two guanine derivatives, 8-oxo-Gua (by 46% at
DNA base damage and repair in nickel-treated rats
Fig. 6. Levels of 5-OHMe-Ura in hepatic DNA. The symbols and
descriptions are the same as in Figure 2.
Fig. 4. Levels of 8-oxo-Gua and FapyGua in hepatic DNA. The symbols
and descriptions are the same as in Figure 2.
Fig. 5. Levels of 5-OH-Hyd and 5,6-diOH-Ura in hepatic DNA. The
symbols and descriptions are the same as in Figure 2. The P-value from the
covariance analysis, necessary for 5-OH-Hyd, is 0.017.
day 1) and FapyGua (by 107% at day 1). Thus, the damage
in liver was more pronounced than in kidney. However, unlike
in the kidney, the liver damage decreased in time to the
background levels for all derivatives except one, 5-OH-Hyd.
This could result from lower accumulation and more complete
clearance of hepatic Ni(II) compared with those in the kidney
or, perhaps, from more efficient repair of oxidative DNA base
damage in the rat liver or both. Whatever its reason, the
persistence of the damage constitutes the most striking difference between these two organs in response to nickel(II), found
in the present experiment. Whether this difference is causally
associated with nickel carcinogenesis or not, remains an open
question.
The extent and slow development of the damage in the
kidney and liver observed in the present study do not concur
with the known accumulation and retention patterns of Ni(II)
in these organs. For example, in the rat kidney, Ni(II) content
reaches its peak between 15 min and 2 h and then decreases
to approximately 25% of the peak value in 6 h and 10% in
16 h after injection (3). The passage of parenteral Ni(II)
through liver is faster and at lower concentration levels (3,4).
Thus, if DNA base damage depended immediately on the
gross tissue Ni(II) contents, its maximum could be expected
to occur within the first few hours post-injection, as is the
case of lipid peroxidation in these organs (21). Further, since
virtually the whole injected Ni(II) dose passes through the
kidney (3,4), the damage in this organ should be greater than
that in the liver. Yet, our earlier analyses of 8-oxo-dG in
kidneys of F344 rats and BALB/c mice 3 and 16 h after Ni(II)
injection (1,22; unpublished) and the present analysis of a
wider spectrum of DNA base products at 12 h, all indicate
that the damaging effect develops after the time of maximum
tissue Ni(II) concentration; and the damage in kidney appears
to be less extensive than in liver. The evolution of oxidative
DNA base damage may thus depend not on gross Ni(II) contents
in the tissue, but rather on subcellular uptake, distribution and/
or retention of the metal. A long intracellular retention,
especially in chromatin, may assist in the damage even in the
presence of repair capability. Obviously, more experiments
have to be done to unveil the nature, timing and sequence of
Ni(II)-associated extra- and intracellular events leading to
DNA base damage.
The relevance of the oxidative DNA base damage to nickel
carcinogenesis stems from the promutagenic properties of at
275
K.S.Kasprzak et al.
least some of the DNA base derivatives analyzed in this study.
Thus, 8-oxo-Gua in template DNA is known to cause GC→TA
transversion mutations (23–25). FapyGua is suspected to induce
GC→CG transversions (26). 5-OHMe-Ura in template DNA
codes as thymine and is not markedly mutagenic during DNA
replication (27), but it may be mutagenic owing to incorporative
mutagenicity. For example, when salmonellae were grown in
the presence of 5-OHMe-dU, mutations were observed due to
predominantly GC→AT transitions (28). 8-oxo-Ade induces
AT→GC transitions and AT→CG transversions in mammalian
cells, and its mutagenic potential appears to be similar to that
of 8-oxo-Gua (29). However, mutagenicity that might be
associated with the remaining DNA base products found
elevated in this study, namely 5-OH-Hyd and 5,6-diOH-Ura,
is unknown. Its full evaluation may be a difficult task because
of the complex chemistry of these bases. Thus, 5,6-diOH-Ura
is the enol form of isodialuric acid. The latter, prevalent in
aqueous solution and in DNA (30,31), is converted into 5,6diOH-Ura during derivatization for the GC/MS-SIM analysis
(32). Further complication arises from the fact that 5,6-diOHUra may result, at least in part, from acid-induced deamination
of 5,6-diOH-Cyt (33). Nevertheless, mechanisms have been
proposed for formation of isodialuric acid in DNA exposed to
oxygen radicals (34). In contrast, the occurrence of 5-OH-Hyd
in oxidatively damaged DNA is doubtful. 5-OH-Hyd is formed
during acid hydrolysis of DNA by decarboxylation of alloxan,
which is the prevalent form in oxygen radical-damaged DNA
(11,33). Thus, the two cytosine derivatives, 5,6-diOH-Ura and
5-OH-Hyd, may reflect the presence of three potentially
mutagenic lesions, i.e. 5,6-diOH-Cyt, isodialuric acid and
alloxan, in genomic DNA. Such lesions, if not properly
repaired, may eventually lead to carcinogenesis. Their promutagenic potentials remain, however, to be established.
In conclusion, the present results suggest that longer persistence of promutagenic oxidative DNA base damage by nickel(II)
in the rat kidney compared with that in the liver, may be
associated with susceptibility of the kidney, but not the liver,
to nickel(II)-initiated carcinogenesis.
Acknowledgements
The authors wish to thank Ms T.L.Schroyer for drafting the figures, Drs
D.Christodoulou and L.K.Keefer for helpful critical comments on this manuscript, and Ms Kathy Breeze for editorial assistance.
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Received on July 22, 1996; revised on October 11, 1996; accepted on October
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