FUNDAMENTAL AND APPLIED TOXICOLOGY 3 7 , 1 6 8 - 180 (1997)
ARTICLE NO. F A 9 7 2 3 1 3
Dichloromethane Metabolism to Formaldehyde and Reaction of
Formaldehyde with Nucleic Acids in Hepatocytes of Rodents and
Humans with and without Glutathione S-Transferase T1 and M1 Genes
Mercedes Casanova,* Douglas A. Bell.f and Henry d'A. Heck*
*Chemical Industry Institute of Toxicology, P.O. Box 12137, Research Triangle Park, North Carolina 27709; and ^National Institute
of Environmental Health Sciences, P.O. Box 12233, Research Triangle Park, North Carolina 27709
Received December 12, 1996; accepted April 9, 1997
Dichloromethane Metabolism to Formaldehyde and Reaction of
Formaldehyde with Nucleic Acids in Hepatocytes of Rodents and
Humans with and without Glutathione S-Transferase 77 and Ml
Genes. Casanova, M., Bell, D. A., and Heck, H. d'A. (1997). Fundam. Appl. Toxicol. 37, 168-180.
Metabolism of dichloromethane (DCM) to formaldehyde
(HCHO) via a glutathione S-transferase (GST) pathway is
thought to be required for its carcinogenic effects in B6C3F,
mice. In humans, this reaction is catalyzed primarily by the
protein product of the gene GSTTl, a member of the Theta class
of GST, and perhaps to a small extent by the protein product
of the gene GSTM1. Humans are polymorphic with respect to
both genes. Since HCHO may bind to both DNA and RNA
forming DNA-protein crosslinks (DPX) and RNA-formaldehyde adducts (RFA), respectively, these products were determined in isolated hepatocytes from B6C3F, mice, F344 rats,
Syrian golden hamsters, and humans to compare species with
respect to the production of HCHO from DCM and its reaction
with nucleic acids. Only mouse hepatocytes formed detectable
amounts of DPX, the quantities of which corresponded well
with quantities of DPX formed in the livers of mice exposed to
DCM in vivo [Casanova, M., Conolly, R. B., and Heck, H. d'A.
(1996). Fundam. Appl. Toxicol. 31,103-116]. Hepatocytes from
all rodent species and from humans with functional GSTTl and
GSTM1 genes formed RFA. No RFA were detected in human
cells lacking these genes. Yields of RFA in hepatocytes of mice
were 4-fold higher than in those of rats, 7-fold higher than in
those of humans, and 14-fold higher than in those of hamsters.
The RFA:DPX ratio in mouse hepatocytes incubated with DCM
was approximately 9.0 ± 1.4, but it was 1.1 ± 0.3 when HCHO
was added directly to the medium, indicating that HCHO generated internally from DCM is not equivalent to that added externally to cells and that it may occupy separate pools. DPX were
not detected in human hepatocytes even at concentrations
equivalent to an in vivo exposure of 10,000 ppm; however, the
possibility that very small amounts of DPX were produced from
DCM cannot be excluded, since HCHO was formed in human
cells. Maximal amounts of DPX|iVer that might be formed in
humans were predicted from the amounts in mice and the relative amounts of RFA in hepatocytes of both species. With predicted DPXUTtr as the dosimeter, the unit risk, the upper 95%
0272-0590/97 $25.00
Copyright O 1997 by the Society of Toxicology.
All rights of reproduction in any form reserved.
confidence limit on the cancer risk, and the margin of exposure
were calculated at several concentrations using the linearized
multistage and benchmark dose methods. Since the actual delivered dose is smaller than that predicted, the results suggest that
DCM poses at most a very low risk of liver cancer to humans.
O 1997 Sotlety of Toilcology.
Species-dependent carcinogenic responses to dichloromethane (DCM) as well as individual human differences in
the metabolism of this compound pose a challenge in efforts
to estimate the carcinogenic risk of DCM to humans. DCM
induced hepatocellular and pulmonary adenomas and carcinomas in B6C3F, mice exposed to concentrations of 2000 or
4000 ppm and increased the incidence of benign mammary
tumors in F344 rats (Mennear et al, 1988), but it did not
cause cancer in Syrian golden hamsters exposed to concentrations as high as 3500 ppm (Burek et al., 1984). Epidemiological studies of workers exposed to DCM have not shown
a correlation between exposure and the induction of lung
or liver cancer (Hearne et al., 1987, 1990), but inherent
differences in the rate at which humans can metabolize DCM
via a glutathione S-transferase (GST)-dependent pathway
have raised the possibility of genetic differences in susceptibility to DCM-induced carcinogenesis (Thier et al, 1993;
Hallier et al, 1993; Pemble et al, 1994).
The GST pathway was recognized as the route most likely
to be responsible for the carcinogenicity of DCM (Green,
1983; Andersen et al, 1987). The activity of this pathway
varies across species in a manner that is consistent with their
susceptibilities to tumor induction by DCM (Green, 1989;
Reitz et al, 1989). This pathway is thought to result in the
generation of a reactive, presumably mutagenic intermediate.
The amount of DCM metabolized via this pathway, calculated using a physiologically based pharmacokinetic (PBPK)
model, was proposed as a surrogate for the dose of this
intermediate in different tissues (Andersen et al, 1987). The
intermediate was not identified, however, and no DNA adducts resulting from exposure of liver or lung tissues of
168
ADDUCT DOSIMETRY OF DICHLOROMETHANE IN HEPATOCYTES
mice to [I4C]DCM were demonstrated (Green et al, 1988;
Ottenwalder and Peter, 1989). Subsequently, formaldehyde
(HCHO) generated from DCM via the GST pathway was
shown to form DNA-protein crosslinks (DPX) in mouse
liver but not in the livers of Syrian golden hamsters (Casanova et al, 1992, 1996). Since mice but not hamsters are
susceptible to the induction of hepatocellular tumors (Burek
et al., 1984; Mennear et al., 1988), these results suggested
that DPX may be involved in the carcinogenic mechanism.
To understand more clearly the metabolism and covalent
binding of DCM in different species, hepatocytes from humans, mice, hamsters, and rats were evaluated with respect
to their ability to metabolize DCM to HCHO and to form
DNA-protein crosslinks. The hepatocyte model for metabolism and covalent binding was validated by comparing the
yields of DPX obtained in mouse hepatocytes with those
previously observed in mouse liver in vivo (Casanova et
al, 1996). As a comparative dosimeter for risk assessment,
however, DPX were found to be of limited utility because
they were detectable only in mouse hepatocytes. A more
sensitive biomarker was needed for studies involving human
cells. In this article, an alternative biomarker for DCM metabolism, RNA-formaldehyde adducts (RFA), is described
that is detectable in hepatocytes from humans, mice, hamsters, and rats.
The study of human cells and tissues is important for risk
assessment, but people are known to be genetically variable
with respect to their ability to metabolize DCM (Bogaards
et al, 1993; Hallier et al, 1993, 1994; Pemble et al, 1994;
Norppa et al, 1995). This variability is related to the presence or absence of a Theta-class glutathione 5-transferase
known as GSTTl-1 (Meyer et al, 1991; Jemth et al, 1996),
the protein product of the gene GSTT1. [A glutathione 5transferase of the Mu class, the protein product of the gene
GSTM1, was reported to metabolize DCM in rats (Blocki et
al, 1994); however, the specific activity of this enzyme
in homogenates of human liver with DCM appears to be
negligible (Bogaards et al, 1993).] The proportion of individuals in different ethnic groups who carry functional alleles for GSTT1 varies from approximately 36% in Chinese
to about 80% in Caucasians and 90% in Mexican-Americans
(Nelson et al, 1995). Whether individuals who carry this
gene ("conjugators") are (Thier et al, 1993; Pemble et al,
1994) or are not (Hallier etal, 1993) at greater risk of cancer
from exposure to DCM than those who do not carry the
gene ("nonconjugators") is controversial. Therefore, in
studies that involved human cells, the presence or absence
of GSTT1 and GSTM1 genes was determined.
MATERIALS AND METHODS
Chemicals. [14C]Dichloromethane (10.4 mCi/mmol; radiochemical purity, 99%) was obtained from DuPont-New England Nuclear (Boston, MA)
in break-seal tubes. For in vitro incubations of hepatocytes with dichloro-
169
methane, [14C]DCM was added to cell suspensions from stock aqueous
solutions, which were prepared in the following manner. A break-seal tube
containing 12 mCi (74 /il) of [14C]dichloromethane and a small (8-ml)
scintillation vial were attached to a vacuum line with Tygon tubing greased
with silicone. The line was evacuated, liquid N2 was placed around the
scintillation vial, and [I4C]DCM was transferred from the break-seal rube
to the vial. When transfer was complete (approximately 10 min), the vial
was detached from the line, placed in ice, and capped with a Mininert valve
(Pierce, Rockford, IL) containing a short 26-gauge needle to provide a vent
for relief of internal pressure as the contents warmed from - 1 9 6 to 0°C.
After the [14C]DCM had thawed, the vial was briefly opened, and 7 ml of
ice-cold distilled water was added to the vial. The vial was immediately
recapped, the needle was removed, and the Mininert valve was closed.
[l4C]Dichloromethane was dissolved in the water by sonicating and shaking
the vial. Analysis of the vial contents was performed by transferring a 25fj.\ aliquot of the solution to a 25-ml volumetric flask containing methanol
and determining the I4C content of the flask by liquid scintillation spectrometry. This analysis showed that the transfer efficiency of [I4C]DCM from the
break-seal tube to the scintillation vial was 99% and that the concentration of
[l4C]dichloromethane in the stock solution was 0.16 M.
[l4C]Formaldehyde (10.0 and 53 mCi/mmol, respectively; radiochemical
purity at least 99%) was purchased from DuPont-New England Nuclear
as aqueous solutions. The vials were opened and the solutions were placed
in screw-capped Reacti-Vials (Pierce, Rockford, IL) that contained a Teflonlined septum. Stock solutions were prepared by diluting the contents of the
vials with distilled water. [l4C]Formaldehyde concentrations of the stock
solutions (29.5 and 8.47 mM, respectively) were determined by liquid scintillation spectrometry.
Chemicals used for the isolation and purification of DNA and RNA from
hepatocytes and for the analysis of [l4C]formaldehyde covalently bound to
DNA and RNA were reported previously (Casanova et al., 1996). All other
chemicals were of the purest grade available commercially.
Laboratory animals. Male B6C3F,/CrlBR mice, Syrian golden hamsters [LAK:LVG(SYR)], and Fischer-344 rats [CDF(F344)/CrlBR] were
purchased from Charles River (Raleigh, NC, or St Constant, Quebec, Canada). Animals were acclimated to the animal facility (room temperature
24°C, relative humidity 50%) for at least 12 days before use and were
maintained on a 12-hr light:dark cycle. All animals received NIH-07 laboratory diet (Zeigler Bros., Gardners, PA) and tap water ad libitum. The animals
were free of rodent pathogens detected in standard viral screens (Microbiological Associates, Bethesda, MD).
Hepatocyte isolation from laboratory animals. Hepatocytes were isolated by a modification of the method of Seglen (1976), as described by
Kedderis and Held (1996). Briefly, animals were anesthetized with Nembutal (0.2 ml per 100 g body weight ip) at approximately 8:00 AM, the peritoneal cavity was opened, and the vena cava was cannulated with a 20-gauge
Angiocath (Becton Dickinson, Sandy, UT). The portal vein was cut, and
perfusion was begun in situ at 5 ml/min (mice), 15 ml/min (rats), or 8 ml/
mm (hamsters) using 37°C Hanks' balanced salt solution (without calcium
or magnesium) containing ethylene glycol ftiX/3-aminoethyl ether)iV.iV.Af.yV-tetraacetic acid (EGTA) (0.5 mM) and Af-2-hydroxyethylpiperazine-/V'-2-ethanesulfonate (Hepes) buffer (10 mM), pH 7.3. After 7 min, the
perfusate was switched to 37°C Williams' medium E (WME) containing
collagenase (0.3 mg/ml) and Hepes buffer (10 mM), pH 7.3. Perfusion was
continued for 10 to 12 min, after which the liver was carefully removed
and placed in a Petri dish containing 20 ml of collagenase solution. The
capsule was gently pulled away with forceps, and the cells were dispersed.
The cell suspension was filtered through two layers of sterile gauze and
diluted with cold WME, and the cells were collected by centrifugation at
50g for 1 min. The preparation was enriched in viable hepatocytes by
centrifugation (8 min) through Percoll (10 ml WME, 9 ml Percoll, and 1
ml of Hanks' solution). Hepatocytes whose viability was less than 80%
based on trypan blue exclusion were not used. The hepatocytes were incubated with [I4C]DCM as described below.
170
CASANOVA, BELL, AND HECK
Human hepatocytes.
Suspensions of freshly isolated (uncultured) human hepatocytes were purchased from the Human Cell Culture Center,
Woodbine, Georgia. Samples were used only if they met certain criteria,
including (1) they were derived from postpubertal subjects to maximize
expression of GST [as described for the rat (Lamartiniere, 1981)]; (2) the
hepatocytes appeared morphologically (microscopically) normal and their
viability (trypan blue exclusion) was at least 70%; and (3) the donors did
not have degenerative liver disease, and no liver trauma was suffered in
accidents (verified by the supplier). Several samples provided by the supplier were rejected owing to low viability or because they were derived
from a prepubertal subject; three were accepted. Two of these were obtained
from organ donors who were accident victims; the third sample was obtained
following a lobectomy to remove a biliary obstruction. Sample 1 was a
male Caucasian, 17 years of age; samples 2 and 3 were female Caucasians,
46 and 34 years of age, respectively. The hepatocytes were suspended in
WME containing 1 fiM insulin, 15 mM Hepes, 100 U/ml penicillin, 100
/jg/ml streptomycin, 12.5 mM L-glutamine, supplemented with 10% lowendotoxin calf serum, and then were air-shipped to the Chemical Industry
Institute of Toxicology in 50-ml centrifuge tubes packed in ice in wellinsulated containers and were delivered by courier. Hepatocytes were collected by low-speed centrifugation and resuspended in WME. The samples
were incubated with [I4C]DCM as described below within 2 to 3 hr of their
arrival. Viabilities of the three samples (trypan blue exclusion) at the start
of the experiments were 80, 74, and 87%, respectively. Viabilities at the
end of the experiments were 75, 70, and 80%, respectively.
Incubation of hepatocytes with ["CVichloromethane. Hepatocytes (4
x 106/ml, 3.0 ml) suspended in WME were placed in 30-ml Erlenmeyer
flasks capped with Mininert valves under an atmosphere of 5% CO 2 in
air. An aliquot of the stock aqueous solution of [ H C]DCM ranging from
approximately 0.05 to 0.3 ml in volume was injected into each flask through
the valve, and the flasks were incubated with shaking (60 cycles/min) at
37°C for 2 hr. Two or three replicate flasks of hepatocytes were used at
each concentration of [I4C]DCM, because preliminary experiments showed
that the viability of hepatocytes was not maintained if single, larger flasks
were used. The final concentration of [14C]DCM in each flask was determined by withdrawing 50 /il from the headspace after 1.5 hr and analyzing
the DCM concentration by gas chromatography as described previously
(Casanova et at, 1996). Amounts of [I4C]DCM added to the incubation
flasks were sufficiently large that the vapor phase concentration remained
virtually unchanged throughout the 2-hr incubation, except for the initial
partitioning between gas and liquid phases. The concentration of [MC]DCM
in the aqueous phase containing hepatocytes was calculated from the headspace [MC]DCM concentration using the partition coefficient
[DCM],,
= 6.0 at 37°C.
The partition coefficient was determined at the above temperature using
flasks containing 3 ml of WME plus mouse hepatocytes, 3 ml of WME
plus heat-inactivated (70°C for 20 min) mouse hepatocytes, or 3 ml of
WME without cells. The presence of cells appeared to slightly increase the
solubility of DCM in WME, since the partition coefficient in the presence
of cells was 12% larger than in the absence of cells. The partition coefficient
measurements showed that equilibration of DCM between liquid and vapor
phases was complete within 10 min and thereafter remained constant for
at least 2 hr. For the given volumes of liquid (ca. 3.2 ml) and vapor (ca.
26.8 ml) in each flask, approximately 42% of the DCM was dissolved in
the aqueous phase.
Incubation of mouse hepatocytes with [l4C]formaldehyde
Hepatocytes
from B6C3F, mice (4 X 107ml, 3.0 ml) suspended in 3 ml of WME were
placed in 30-ml Erlenmeyer flasks capped with Mininert valves under an
atmosphere of 5% CO 2 in air. An aliquot from one of the stock aqueous
solutions of [l4C]formaldehyde ranging from 39 to 300 fi\ in volume was
injected into each flask through the valve, and water was added to bring
the final liquid volumes of all flasks to 3.3 ml. The flasks were incubated
with shaking at 37°C for 2 hr. Two replicate flasks of hepatocytes were used
at each concentration of [l4C]formaldehyde. At the end of the incubation, the
flasks were opened, replicate flasks were combined, and the cells were
collected as described above.
The concentrations of [MC]formaldehyde in the aqueous phase in these
experiments were assumed to be equal to the amounts of [l4C]formaldehyde
added to each flask divided by the aqueous volume; i.e., volatilization of
HCHO from the aqueous to the vapor phase was assumed to be negligible.
This assumption was justified on the basis of the Henry's law constant for
HCHO in water at 37°C. The value of this constant can be calculated from
the data of Betterton and Hoffmann (1988) as 1.23 X 103 M atm"1; this
corresponds to the partition coefficient [ H C H O J ^ ^ ^ t H C H O ] , ^ , = 3.13
X 104. Thus for the given volumes of liquid (3.3 ml) and vapor (26.7 ml)
in each flask, more than 99.9% of the HCHO is expected to be dissolved
in the aqueous phase.
Analysis ofcovalently bound [I4C]formaldehyde in DNA and RNA. At
the end of the incubation with [MC]DCM or [l4C]formaJdehyde, the flasks
were opened in a hood, replicate flasks were combined, and the volume
was brought to 30 ml with cold WME. The cells were collected by centrifugation, and the supernatant was discarded. Lysing solution (9 ml) containing
8 M urea and 1% sodium dodecyl sulfate was added to the pellet, and the
cells were ruptured by vortexing. The nucleic acids were sheared by several
passages through 18- and 20-gauge needles, and proteins were digested
with proteinase K to solubilize the DNA that was crosslinked to proteins
(Casanova-Schmitz and Heck, 1983; Casanova et ai, 1989). The solutions
were extracted with an equal volume of chloroform/Zioamyl alcohol/phenol
(CIP), then the aqueous phase containing nucleic acids was extracted with
diethyl ether. Nucleic acids (DNA and RNA) were purified by hydroxyapatite chromatography (Casanova and Heck, 1987) DNA was incubated with
ribonuclease A (0.33 mg/ml) and washed by ultrafiltration to remove residual nbonucleotides.
The nucleic acids were heated (70°C) overnight, then were enzymatically
hydrolyzed (2-3 hr) to deoxyribonucleosides or ribonucleosides in 0.1 M
Tris buffer, which resulted in the quantitative release of [14C]formaldehyde
(Casanova et ai, 1989) DNA was hydrolyzed as described previously;
RNA was hydrolyzed using the same enzymes as were used for DNA plus
ribonuclease A (0.67 mg/ml). Analysis of the nucleic acid hydrolyzates by
high-performance liquid chromatography (HPLC) (Casanova et ai, 1989)
showed that both DNA and RNA were completely hydrolyzed to their
corresponding nucleosides and that no l4C-labeled adducts were detectable
in either hydrolyzate. The released [l4C]formaldehyde was collected in a
single tube, acidified to pH 4.8 with acetate buffer, and precipitated with
dimedone in the presence of carrier unlabeled HCHO (Casanova el ai,
1989). Precipitation efficiencies of [l4C]formaldehyde under the experimental conditions were determined separately in each experiment and were at
least 90%. Dimedone precipitates were collected on filters, the precipitates
were dissolved in Hionic-Fluor (Packard, Meriden, CT), and the quantities
of ['4C]formaldehyde were determined by liquid scintillation spectrometry.
Analysis of DNA and RNA was performed by integration of deoxyribonucleoside or ribonucleoside peak areas at 260 nm, and the amount of
nucleoside in each peak was computed from standard curves that spanned
the range of interest.
Contribution of RNA-Protein crosslinks to RNA-formaldehyde adducts. To determine whether the [l4C]formaldehyde covalently bound
to RNA was due to the formation of RNA-protein crosslinks, hepatocytes were incubated for 2 hr with [I4C]DCM (concentration in the
aqueous phase = 3.2 mM), and then the cells were lysed with sodium
dodecyl sulfate-8 M urea as described above. The lysed cells were
extracted with CIP before digesting the proteins with proteinase K,
which partitioned the nucleic acids into aqueous (AQ) and interfacial
(IF) fractions; proteins were mainly extracted into the organic phase
(Casanova-Schmitz and Heck, 1983; Casanova-Schmitz et at, 1984;
Casanova et al., 1989). Nucleic acids crosslinked to proteins were
assumed to be present in the IF fraction, as shown previously (Casanova-Schmitz and Heck, 1983; Casanova-Schmitz et ai, 1984; Casanova et at, 1989). The nucleic acids in the IF fraction were solubilized
171
ADDUCT DOSIMETRY OF DICHLOROMETHANE IN HEPATOCYTES
by digesting the proteins with proteinase K. Both AQ RNA and IF RNA
(a)
were purified by hydroxyapatite chromatography and were washed by
o
ultrafiltration (Casanova-Schmitz and Heck, 1983; Casanova-Schmitz
DC 150- o
et al., 1984). Following enzymatic hydrolysis of the RNA, the quantity
O)
of released [14C]formaldehyde was determined as described above (Cas100anova et al., 1989).
E
S
Analysis of glutathione S-transferase genes in human hepatocytes. A
polymerase chain reaction (PCR) method was used to detect the presence
of GSTT1 in human cells. Another glutathione S-transferase gene, GSTM1,
was also identified, although as mentioned previously, the protein product
of this gene may not be involved in the metabolism of DCM (Bogaards et
al, 1993; Blocki et ah, 1994). The multiplex PCR procedure used has
been published previously (Chen el al., 1996). Briefly, this method is a
modification of methods used for analysis of GSTM1 (Bell et al., 1993)
and GSTTI (Pemble et al., 1994) that includes both sets of GST primers
in the same PCR with an annealing temperature of 62°C. /J-Globin primers
were added as an internal reference control to monitor the success of each
reaction.
RESULTS
DNA -protein Crosslinks in Mouse, Hamster, Rat, and
Human Hepatocytes Incubated with
['4' C]Dichloromethane
The results of experiments designed to detect covalently
bound [l4C]formaldehyde in the DNA of mouse, hamster,
rat, and human hepatocytes are depicted in Fig. 1. The range
of aqueous DCM concentrations was chosen on the basis of
PBPK model simulations (Andersen et al., 1987) that predicted the time-weighted-average concentrations of DCM
expected to occur in mouse liver during a 6-hr inhalation
25
20
o
o
A
X
g>15
•
MOUSE
RAT
HAMSTER
HUMAN 1
HUMAN 2
HUMAN 3
o
1
2
3
4
[DCM] In medium (inH)
5
6
FIG. 1. Formation of DNA-protein crosslinks (DPX) in hepatocytes
of different species incubated (2 hr) with [I4C]DCM. DPX formation was
detectable in hepatocytes of male B6C3F, mice but not in hepatocytes of
F344 rats, Syrian golden hamsters, or humans with or without functional
GSTTI genes (see text). A DCM concentration of 1.9 mM is equivalent to
the time-weighted-average concentration expected to occur in mouse liver
dunng a 6-hr exposure to 4000 ppm, according to a PBPK model (Andersen
eta!., 1987).
u.
50-
E
UOUSE
BAT
HAUSTEfl
O
/ Q
y/
•
•
/
/
/
H-1 GSTT1(+)
H-2 Q3TT1(+)
H-3 6STT1(-)
O
o ..--*'
e"
0-
A
1 2
3 4 5 6
[DCM] In medium (mM)
0
1 2
3 4 5 6
[DCM] In medium (mM)
FIG. 2. Formation of RNA-formaldehyde adducts (RFA) in hepatocytes of different species incubated (2 hr) with [I4C]DCM. (a) RFA formation in hepatocytes of male B6C3F, mice, F344 rats, and Syrian golden
hamsters; yields of RFA in mice are approximately 4-fold higher than in
rats and 14-fold higher than in hamsters, (b) RFA formation in hepatocytes
of three humans. Hepatocytes of humans 1 and 2 produced RFA and both
contained functional GSTTI (see text); human 3 did not produce RFA and
did not contain functional GSTTI (see text). The yields of RFA in mice (a)
are approximately 7-fold higher than in humans with GSTTI.
exposure. For example, the predicted time-weighted-average
concentration of DCM in mouse liver during a 6-hr exposure
to 4000 ppm is 1.9 mM, which is near the midpoint of the
concentration-response curve (Fig. 1). Concentrations of
[I4C]DCM in the aqueous phase were calculated from the
measured concentrations in the vapor phase by assuming
a liquid/vapor phase partition coefficient equal to 6.0 (see
Materials and Methods).
As shown in Fig. 1, DPX were detectable only in mouse
hepatocytes. In these cells the yield of DPX appeared to
increase in direct proportion to the concentration of DCM
in the medium to concentrations at least as high as 2 mM. The
line shown in the figure is a linear least-squares regression
(through the origin) of the mouse DPX data on the DCM
concentration. Although a simple linear model fits the DPX
response of mouse hepatocytes satisfactorily, this does not
rule out the possibility of a more complex concentration
dependence. Note that DPX were not detected in human
hepatocytes even at aqueous DCM concentrations as high
as 5 mM, which is equivalent to the time-weighted-average
concentration predicted to occur in mouse liver during a 6hr exposure to a DCM concentration > 10,000 ppm.
RNA-Formaldehyde Adducts in Mouse, Hamster, Rat,
and Human Hepatocytes Incubated with
[I4C\Dichloromethane
The results of the analysis of RFA in mouse, hamster,
and rat hepatocytes are shown in Fig. 2a. Hepatocytes from
all species formed measurable amounts of RFA, with mouse
hepatocytes producing about 4-fold higher yields of RFA
than rat hepatocytes and about 14-fold higher yields than
hamster hepatocytes. Formation of RFA appeared to be linearly related to the concentration of DCM in the medium.
172
CASANOVA, BELL, AND HECK
The lines drawn in the figure are linear least-squares regressions (through the origin) of the RFA data on the DCM
concentration.
Results obtained using three samples of human hepatocytes are depicted in Fig. 2b. Note that the RFA data for
human hepatocytes (Fig. 2b) are shown on an expanded scale
relative to that used for rodent hepatocytes (Fig. 2a). Two
of the human samples, denoted as H-l and H-2, produced
measurable quantities of RFA. The third sample, denoted
as H-3, produced no detectable RFA, indicating that the
hepatocytes in this sample did not metabolize [I4C]DCM to
[l4C]formaldehyde. As discussed below, samples H-l and
H-2 were obtained from individuals whose DNA contained
GSTT1 and GSTM1 genes; sample H-3 was obtained from
an individual whose DNA lacked both genes. The yields of
RFA in the two human hepatocyte samples that produced
this adduct (H-l and H-2) were about one-seventh of those
in mouse hepatocytes.
Although the hepatocytes in sample H-3 did not metabolize [I4C]DCM to [14C]formaldehyde, they did metabolize
[I4C]DCM to [l4C]formate, presumably via the cytochrome
P450 2E1 pathway for DCM (Pankow et al, 1991a,b), implying that these hepatocytes were metabolically competent.
HCHO is not an intermediate in the P450 pathway (Gargas
et al., 1986). The production of [l4C]formate from [14C]DCM
was shown by the fact that the hepatocytes incorporated
14
C (derived from [l4C]formate by one-carbon biosynthetic
pathways) into adenosine and guanosine (approximately 800
pmol equivalents of I4C per mg of RNA).
On average, the ratio of RFA to DPX in mouse hepatocytes was about 9.0 ± 1.4 (JT ± SE, n = 7). If DPX had
been formed in GS777-positive human cells, and if the
RFA:DPX ratio in human cells were the same as in mouse
cells, then DNA-protein crosslinks should have been
detected in human hepatocytes; i.e., the expected yield of
DPX in human cells would have been above the detection
limit of the analytical method. (The same conclusion also
applies to cells from the rat and hamster; i.e., DPX should
also have been found in these species.) Therefore, the
RFA:DPX ratio in human, rat, and hamster hepatocytes
was greater than 9.0 ± 1.4.
Contribution of RNA-Protein
Formaldehyde Adducts
Crosslinks to RNA-
To estimate the amount of RNA-protein crosslinks that
may be present in RFA, mouse hepatocytes were incubated
with [14C]DCM, and then the RNA was fractionated into
aqueous and interfacial portions by extraction with CIP (see
Materials and Methods). The AQ RNA and IF RNA were
separately purified by hydroxyapatite chromatography. After
enzymatic hydrolysis of the RNA, the amount of [ I4 C]formaldehyde covalently bound to each RNA fraction was
determined (Table 1).
TABLE 1
Covalent Binding of [uC]Fonnaldehyde Derived from [UC]Dichloromethane to Aqueous and Interfacial RNA from Mouse
Hepatocytes0
RNA fraction
Quantity of
RNA (jig)
Concentration of
[14C]formaldehyde
bound to RNA
(pmol/mg)
Interfacial RNA
Aqueous RNA
75 ± 34
2460 ± 2 4 0
360+150
104 ±
8
Distribution of total
[uC]forma]dehyde
bound to RNA
(%)
6.5 ± 3.0
93.5 ± 3.0
° Hepatocytes were incubated with [I4C]DCM (3.2 ITIM) for 2 hr, and
then were lysed and extracted with CIP. Interfacial RNA was solubilized
by digestion of interfacial proteins with proteinase K. RNA fractions were
purified by hydroxyapatite chromatography and then enzymatically hydrolyzed to nucleosides in 0.1 M Tris buffer, which released covalently
bound [HC]formaldehyde. RNA and [l4C]formaldehyde were analyzed by
HPLC and liquid scintillation spectrometry. values shown are X± SE, n =
3 sets of hepatocytes, each set isolated from a different mouse.
As shown in the table, approximately 93% of the total
[l4C]formaldehyde in the AQ and IF RNA was associated
with the aqueous fraction. This formaldehyde was unlikely
to have been bound as RNA-protein crosslinks, because
proteins crosslinked to RNA would probably have been extracted from the aqueous phase into the interface with CIP,
as was previously observed with DNA (Casanova-Schmitz
and Heck, 1983; Casanova et al., 1989), or into the organic
phase. Although most of the [14C]formaldehyde bound to
RNA was present in the AQ RNA fraction, the concentration
of covalently bound [14C]formaldehyde (pmol [14C]formaldehyde/mg RNA) was approximately fourfold higher
in the IF RNA than in the AQ RNA (Table 1). The higher
concentration of [l4C]formaldehyde in the IF RNA suggests
that a portion of this [l4C]formaldehyde may have been
bound as RNA-protein crosslinks.
DNA—Protein Crosslinks and RNA-Formaldehyde
Adducts in Mouse Hepatocytes Incubated with
[NC]Formaldehyde
The amounts of [14C]formaldehyde that are covalently
bound to RNA or to DNA may depend on both the cell type
and the source of [l4C]formaldehyde. To investigate whether
the source of [l4C]formaldehyde may affect the amounts
bound to nucleic acids, mouse hepatocytes were incubated
for 2 hr with [14C]formaldehyde that was added directly to
the cell suspension. The HCHO concentrations were chosen
on the basis of preliminary studies to provide yields of DPX
that were similar to those obtained with [I4C]DCM, and
to ensure that hepatocyte viability (trypan blue exclusion)
remained relatively high (at least 70%) thoughout the 2-hr
incubations. The yields of DPX and RFA that were derived
from externally added [l4C]formaldehyde are shown in Fig.
ADDUCT DOSIMETRY OF DICHLOROMETHANE IN HEPATOCYTES
3a. These results can be compared with the yields of DPX
and RFA that were obtained by incubating mouse hepatocytes with [14C]DCM. The latter results are reproduced in
Fig. 3b.
Figure 3 demonstrates that the relative amounts of HCHO
bound to mouse DNA and RNA depend on whether the
formaldehyde was added directly to the medium or was generated internally as a metabolite of DCM. When HCHO was
added to the medium (Fig. 3a), the ratio of RFA to DPX
was 1.1 ± 0.3 (Z± SE, n — 9), which was almost an orderof-magnitude smaller than the ratio obtained when the hepatocytes were incubated with DCM (9.0 ± 1.4; see above).
Thus, at a given yield of DPX, the yield of RFA was much
smaller when HCHO was added externally to cells than when
it was generated internally by metabolism of DCM. Moreover, the concentration dependence of DPX and RFA formation in mouse hepatocytes was apparently nonlinear when
HCHO was added directly to the medium (Fig. 3a), but it
appeared to be linear when HCHO was generated intracellularly from DCM (Fig. 3b).
The results of the PCR analysis of glutathione 5-transferase genes in the three human hepatocytes samples are
shown in lanes 1, 2, and 3 of Fig. 4. Both H-l and H-2 were
found to possess GSTT1 and GSTMJ genes, and since the
cells produced formaldehyde from DCM (Fig. 2b), GSTT1
was clearly functional. In contrast, H-3 did not appear to
contain either gene. All human samples produced a fragment
from the /3-globin gene (internal reference standard), which
showed that the PCR procedure was successful and that
measurable amounts of DNA were produced. The presence
of GSTT1 in samples H-l and H-2 and the absence of this
gene in sample H-3 are consistent with the fact that RFA
derived from DCM were detected only in the first two sam-
IDINQ
eld)
20
I
200(b)
o
± a 150
CD o
y
EHYD
g nuci
MI'S
100-
91
50-
"I
0
0.5
1
1.6
[HCHO] In medium (mM)
2
o
Q
/°
RFA/
/
o
/
/i-— ••-•-—
0
B 5
6 M
-QSTT1
-
0-GLOBIN
QSTM1
FIG. 4. DNA from human liver samples H-l (lane 1), H-2 (lane 2),.
and H-3 (lane 3) was tested for the presence of the GSTT1 and GSTM1
genes. Results from multiplex PCR analysis of GSTT1 and GSTM1 are
shown. GSTT1 null genotypes lack a 480-bp PCR product (lanes 3 and 6).
GSTM1 null individuals lack a 215-bp PCR product (lanes 3 and 5). All
samples produced a fragment from the /9-globin gene (internal reference
standard). Lanes 5 and 6 are positive/negative controls for GSTT1 and
GSTM1. B designates PCR reagent negative control. M is * X 174 //inDIII
marker.
pies. The results of Figs. 2b and 4 suggest that samples H1 and H-2 were derived from conjugators, but that sample
H-3 was obtained from a nonconjugator.
PCR Analysis of GSTT1 and GSTM1 in Human
Hepatocytes
O
M 1 2 3
•
2
4
6
[DCM] In medium (mM)
FIG. 3. Covalent binding of HCHO to nucleic acids in mouse hepatocytes. (a) Formation of DPX and RFA in hepatocytes incubated (2 hr) with
[l4C]formaldehyde; the RFA:DPX ratio is 1.1 ± 0.3. (b) Formation of DPX
and RFA in hepatocytes incubated (2 hr) with [UC]DCM; the RFA:DPX
ratio is 9.0 ± 1.4.
DISCUSSION
The Hepatocyte Model: Validation and Limitations
To validate the hepatocyte model, the DPX yields measured in vivo in mice exposed to 4000 ppm (Casanova et
ai, 1996) were compared with those obtained in vitro in
mouse hepatocytes incubated with [I4C]DCM at an aqueous
concentration equivalent to the time-weighted-average concentration of DCM predicted to occur in mouse liver at
this exposure concentration. As noted previously, the PBPK
model (Andersen et al., 1987) predicts that the timeweighted-average concentration of DCM in the liver of a
28-g mouse during a 6-hr exposure to 4000 ppm should be
about 1.9 mM.
The yield of DPX in mouse liver at the end of a 6-hr
exposure to 4000 ppm of DCM was approximately 29.5 ±
8.9 pmol/mg DNA (JT± 95% CL) (Casanova et al., 1996).
In comparison, the yield of DPX in mouse hepatocytes at
the end of a 2-hr incubation with 1.9 mM DCM was 6.6 ±
2.2 pmol/mg DNA (Fig. 1). Assuming that the yield of DPX
increases linearly with time, the expected yield of DPX following a 6-hr exposure of hepatocytes would be 19.9 ± 6.7
pmol/mg DNA. The predicted yield is about 70% of the
observed in vivo value. These results support the hepatocyte
model as a semiquantitative tool for the investigation of
DPX formation. The in vivo-in vitro difference in DPX
yields may be due to uncertainties in the intracellular concentrations of DCM or to differences in the metabolic rate constants of DCM and HCHO.
174
CASANOVA, BELL, AND HECK
Although the yields of DPX in mouse hepatocytes were
similar to those in mouse liver, the formation of DPX was
an apparently linear function of the DCM concentration in
vitro to concentrations as high as 2 DIM (equivalent to an in
vivo exposure of ~4000 ppm) (Fig. 1), but it was a nonlinear
function of the DCM concentration in vivo in this concentration range (Casanova et al., 1996). However, freshly isolated
hepatocytes are considerably less stable than normal liver
cells, they lack normal cell-to-cell communications and other
interactions with neighboring cells, and they are suspended
in a nutrient medium that differs from that in the liver.
Therefore, the two systems should not necessarily display
identical metabolic or kinetic characteristics. Reasonably
close similarities in metabolic behavior of isolated hepatocytes and whole animals have been reported for a few compounds (Carfagna et al., 1993; Kedderis et al., 1993; Kedderis and Held, 1996).
Species Differences in the Formation of DNA—Protein
Crosslinks in Hepatocytes
Hepatocytes from B6C3F, mice, a species that is susceptible to DCM-induced hepatocarcinogenesis, formed DPX
when incubated with DCM, but no DPX were detected in
the hepatocytes of rats, hamsters, or humans. The in vitro
results for the mouse and hamster are consistent with in vivo
data on DPX formation in which mice but not hamsters
formed DPX (Casanova et al., 1996). The absence of detectable DPX in human hepatocytes (including hepatocytes from
individuals carrying the GSTT1 gene), even at DCM concentrations that in mice would be equivalent to an exposure
concentration > 10,000 ppm, strongly suggests that if DPX
are formed at all in human cells, the amounts must be very
much smaller than in mice. This conclusion also applies to
rats and hamsters. Thus, among all species tested, the mouse
appears to be uniquely capable of forming DPX from DCM.
The present findings on DPX formation in hepatocytes
from mice, rats, hamsters, and humans are consistent with
observations of Graves et al. (1995) regarding the formation
of DNA single-strand (ss) breaks in hepatocytes from these
species. These investigators detected DNA ss breaks in
mouse hepatocytes incubated with DCM at aqueous concentrations 5= 0.4 mM, which are similar to the concentrations
at which DPX were detected in mouse hepatocytes (Fig. 1).
Graves et al. (1995) also observed the formation of DNA
ss breaks in rat hepatocytes, but only at concentrations 5= 30
mM. According to Graves et al. (1995), DCM concentrations
above 30 mM are equivalent to in vivo concentrations >
86,000 ppm, which exceed the acute lethal concentration of
DCM in rodents. Graves et al. (1995) did not observe DNA
ss breaks in either hamster or human (n = 8) hepatocytes
at concentrations as high as 90 or 120 mM, respectively.
Thus, only mouse hepatocytes were capable of forming either DPX or DNA ss breaks at concentrations below 5 mM.
RNA-Formaldehyde Adducts: An Alternative Dosimeter
Related to the Activity of GST with Respect to DCM
RNA exists in cells in a variety of forms, including heterogeneous nuclear RNA, messenger RNA, transfer RNA, and
ribosomal RNA and its precursors (Alberts et al., 1983). The
different forms of RNA are compartmentalized, with most
of the RNA being ribosomal (ca. 75%) and the remainder
being nuclear (ca. 10%) or cytoplasmic (ca. 15%). The particular RNAs to which HCHO is bound may therefore depend on the subcellular locations in which HCHO is produced. The yield of RFA may depend on the rates of HCHO
generation and detoxication, the local concentration and
types of RNA, and the rate of RNA turnover.
RNA-formaldehyde adducts are a novel dosimeter for
formaldehyde that offers the advantage of greater sensitivity
than DPX, at least for measurements of HCHO production
from DCM. The structures of RFA remain to be determined.
As noted previously, the [l4C]formaldehyde linked to RNA
does not seem to have been bound primarily as RNA-protein
crosslinks. Therefore, the HCHO may have been bound as
intramolecular RNA-RNA crosslinks [structures of one relatively stable cyclic guanosine-formaldehyde adduct and
of two guanosine-formaldehyde-guanosine crosslinks have
been reported (Kennedy et al., 1996)] or as RNA crosslinked
to small molecules. Hydroxymethyl adducts are possible but
are regarded as unlikely, because these adducts are unstable
(McGhee and von Hippel, 1975a,b), and they would probably not have survived the purification procedures. The term
RNA-formaldehyde adducts is intended to denote all forms
of HCHO that were detected by the analytical procedure.
As shown in Fig. 2, there were significant differences
among species with respect to the yields of RFA obtained
in mouse, rat, hamster, and human hepatocytes incubated
with DCM. Specifically, the yields of RFA in rat, hamster,
and human hepatocytes with GSTT1 (H-l and H-2) were
about l/4th, 1/14th, and l/7th, respectively, of those in
mouse hepatocytes. Similarly, Reitz et al. (1989) reported
that the activities of GST with respect to DCM in cytosolic
preparations from livers of rats, hamsters, and humans with
GST activity (n - 3; one of four samples had no measurable
activity) were about l/4th, l/20th, and l/9th, respectively,
of that from livers of mice. Thus, the yields of RFA in
hepatocytes appear to be correlated with the activity of GST
with respect to DCM.
Another parameter that could affect the rate of DCM metabolism in hepatocytes is the intracellular concentration of
glutathione (GSH). The concentration of GSH in hepatocytes
varies among species, with values ranging from 129.6 ±
11.5 nmol/106 cells in B6C3F, mice (Ruch et al, 1989) to
about 50 nmol/106 cells in Sprague-Dawley rats (Jones et
al, 1978), 22.3 ± 4.9 nmol/106 cells in Syrian golden hamsters, and 21.4 ± 2.6 nmol/106 cells in human cells (n = 4)
ADDUCT DOSIMETRY OF DICHLOROMETHANE IN HEPATOCYTES
(Steinmetz et al, 1988).' Hence, the range of GSH concentrations in hepatocytes from mice to hamsters and humans
is approximately 6-fold, which together with differences in
the specific activity of GST (Green, 1989; Reitz et al, 1989)
could explain the differences observed in RFA formation in
hepatocytes of different species.
The generation of RFA from DCM in human hepatocytes
appears to depend on the presence of GSTT1. The number
of human samples used in this study was small and in itself
would not be compelling evidence for an essential role for
this gene; however, the present findings are supported by
several other human studies that have demonstrated interindividual differences in GSTT1 and have shown that the presence or absence of this gene can explain differences in the
metabolism or genotoxicity of DCM: (1) Hallier et al.
(1993), n = 10: five hemolyzate samples lacked GST activity, and sister chromatid exchange (SCE) induction in lympocytes incubated with DCM was inversely related to this
activity; (2) Hallier et al. (1994), n = 13: six hemolyzate
samples lacked GST activity with respect to DCM; (3) Pemble et al. (1994), n = 8: four hemolyzate samples lacked
GST activity with respect to DCM, and the blood samples
lacked GSTT1; (4) Norppa et al. (1995), n = 20: eight erythrocyte samples lacked GST activity with respect to DCM,
and the blood samples lacked GSTT1. Thus, including the
present study, the total number of individual human samples
characterized with respect to DCM metabolism and GSTT1
or susceptibility to SCE induction by DCM in vitro is currently 54, of which 24 lacked the ability to metabolize DCM
or were more susceptible than metabolically competent samples to SCE induction by DCM in lymphocytes.
The most extensive studies of human variability with respect to DCM metabolism suggest the existence of two subpopulations of conjugators with low and high GST activity,
respectively, the difference in mean activities of the two
subpopulations being about a factor of 3, and the overall
range from lowest to highest being about a factor of 10
[Bogaards et al. (1993), n = 22: three liver samples had no
measurable GST activity; Hallier et al. (1994), n = 13 (see
above); Norppa et al. (1995), n = 20 (see above)]. To date,
over 800 humans have been characterized with respect to
the presence or absence of GSTT1 (Nelson et al., 1995), but
comparisons among humans, mice, rats, and hamsters with
respect to the metabolism or genotoxicity of DCM are relatively few [Reitz et al. (1989): 4 human samples; Green
(1989): 12 human samples; Graves et al. (1995): 8 human
samples; this work: 3 human samples], i.e., 27 human sam' Measurements of GSH in hepatocytes of mice (104 nmol/106 cells) and
hamsters (30 nmol/106 cells) undertaken in this laboratory according to
Sedlak and Lindsay (1968) were similar to values reported in the literature.
Note that intracellular glutathione is compartmentalized (Smith et al, 1996)
and exhibits diurnal fluctuations. To minimize the possible effects of such
changes, hepatocytes were isolated from rodents at the same time each day.
175
pies in total, of which only 7 (Reitz et al., 1989; this work)
provide a quantitative interspecies (human-mouse) comparison. The present study is the only one to our knowledge in
which DCM metabolism in human cells from a potential
target tissue was determined together with GSTT1.
Evidence for Different HCHO Pools When HCHO Is
Added Directly to the Medium or Generated
Metabolically from DCM
The ratio of RFA to DPX in mouse hepatocytes treated
with DCM was 9.0 ± 1.4, whereas the RFA:DPX ratio in
cells treated with HCHO was 1.1 ± 0.3 (Fig. 3). The difference in these ratios indicates that HCHO generated internally
from DCM is not equivalent to that added externally to cells,
and it suggests that separate pools and different concentrations of HCHO are produced from these two sources of
HCHO. The reasons for the dependence of the RFA:DPX
ratio on the source of HCHO are not well understood. Possible explanations might include compartmentalization of
DCM metabolism, HCHO metabolism, RNA, or all three.
Moreover, different types of RNA may be bound to HCHO
depending on the intracellular localization of the compound.
Differences in the intracellular concentration of HCHO may
explain the fact that the concentration-response curves for
DPX and RFA formation were nonlinear when HCHO was
added directly to the medium but appeared to be linear when
HCHO was generated internally from DCM.
Further studies should be undertaken to understand the
basis for the differences in covalent binding to nucleic acids
when HCHO is generated intracellularly by metabolism or is
added directly to cells. Given the present findings, however,
caution should be used when interpreting studies designed
to compare the types of DNA damage induced by HCHO
when added externally to cells with those induced by DCM,
which have led some authors to conclude that HCHO is
not responsible for the genotoxicity of DCM (Graves et al.,
1994a,b, 1996; Guengerich et al., 1995; Graves and Green,
1996). Such a conclusion may be unwarranted. The two
forms of formaldehyde do not occupy the same pool, and
their relative reactivities with macromolecules appear to be
quite different.
Implications for Human Risk Assessment
The absence of detectable DPX in human hepatocytes
carrying GSTT1 and GSTMI genes, even at extremely high
concentrations of DCM, indicates that humans are far less
likely than mice to form this potentially mutagenic adduct
from DCM; however, the possibility that very small
amounts of HCHO may react with DNA in human cells
cannot be excluded, since HCHO was generated from
DCM in GS777-positive human hepatocytes. Assuming
that DPX were formed in these cells, the maximal quantities that might be produced in human hepatocytes can be
176
CASANOVA, BELL, AND HECK
HUMAN DOSE ESTIMATE
RISK ASSESSMENT
DCM Exposure
Concentration
(ppm)
Mouse Tumor Response
In Observable Range,
1-exp(-oyxDPX ||ver )
PBPK
model
Calculated AUC liver
(mM x hr)
LMS Method
Calculation of MLE
and UCL for
Lifetime Exposure
(6 hr/day, 5 days/wk)
Extended PBPK model,
mice
Calculated I
Mice
(pmol/mg DNA)
Adjustment for
duration of exposure,
X (8/6 x 48/52 x yr/7O)
Adjustment for
assumed mouse-human
difference In D P X , ^ , x (1/7)
Predicted DPX ||v0 7
GSTT1(+) Humans
(pmol/mg DNA)
BMD Method
Calculation of
ED 10 andLED 10
Linear
Extrapolation
from LED1Q
Adjusted Risk
Upper Bound
on Risk
-> and Margin
of Exposure
FIG. 5. Approaches to the risk assessment of DCM using DNA—protein crosslinks (DPX) as the dosimeter. The left side (human dose estimate)
shows a procedure for estimating DPXUvCT in humans involving (I) prediction of the A l I C ^ for DCM in humans exposed for 6 hr using a PBPK model
(Andersen et ai, 1987); (2) calculation of DPX in mouse liver containing the same time-weighted-average concentration of DCM (Casanova et ai,
1996); (3) estimation of DPX in human liver by multiplying the value of DPXDvCT in mice by 1/7 (see text). The right side (risk assessment) shows two
procedures (LMS method and BMD method) to estimate human risk based on an empirical relationship between DPXLvt, and the mouse tumor response
observed at 2000 and 4000 ppm (Casanova et ai, 1996; Mennear et ai, 1988). In the LMS method (U.S. EPA, 1986), lifetime risk estimates (MLE and
UCL) for exposures 6 hr/day, 5 days/week, are calculated by application of this relationship at the concentrations of interest. The risks are adjusted for
the duration of exposures. In the BMD method (U.S. EPA, 1996), the ED, 0 and LED,0 are calculated from the empirical relationship, and a linear
extrapolation is made from the LED, 0 to zero dose, zero risk. The extrapolation provides the upper bounds on the risk at the exposure concentrations of
interest. The margin of exposure (dimensionless) is the ratio of the LED I0 to the exposure concentration of interest (see text).
estimated by the method outlined in Fig. 5. The resulting
maximal estimate of DPX can be used as an internal
dosimeter to provide an upper bound on the likelihood of
cancer for humans with metabolic characteristics similar
to those of subjects H-l and H-2.
The use of DPX as a dosimeter for risk assessment is
based on the assertion that DPX are a measure of the area
under the curve (AUC) of reactive formaldehyde in target
cells (Hernandez et al., 1994). According to the U.S. Environmental Protection Agency (U.S. EPA), DPX may be used
as a surrogate for the internal dose "whether DPX are mechanistically involved in the carcinogenic process or are simply
an indicator of intracellular exposure" (Hernandez et al.,
1994). Thus a defined role for DPX in the carcinogenic
mechanism is not an essential requirement for using DPX
as a dosimeter.
The left side of the figure describes a method to estimate
the amount of DPX that might be formed in the livers of
GS7T7-positive humans with metabolic capabilities similar
to those of subjects H-l and H-2. The yields of DPX in
human liver can be predicted from the quantities of DPX in
mouse liver and the relative amounts of RFA in hepatocytes
of both species. This calculation assumes that the RFA:DPX
ratio in human hepatocytes is identical to that in mouse
hepatocytes (9.0 ± 1.4) (Fig. 3b); however, since the
RFA:DPX ratio in human hepatocytes was > 9.0 ± 1.4, and
the yield of RFA in hepatocytes from subjects H-l and H2 was about one-seventh of that in mouse cells, it follows
that the yield of DPX in human hepatocytes was less than
one-seventh of that in mouse hepatocytes. Nevertheless, a
value of 1/7 is assumed for the amount of DPX in GSTT1positive human hepatocytes relative to mouse cells. Ideally,
a much larger number of human samples should be measured
to develop a more precise estimate of the maximum yield
of DPX in human hepatocytes; however, the value of 1/7 is
consistent with other data on the relative activity of GST in
liver cytosolic preparations from humans and mice, i.e.,
1/9 (Reitz et al., 1989) and < l / 9 (Green, 1989), and as a
maximal estimate of DPX that might be formed in human
liver, it is health-protective.
The prediction of DPX in human liver involves (1) calculation of the AUC of DCM in human liver (JJ.M X hr) during
a 6-hr exposure to a selected airborne concentration of DCM
using a PBPK model (Andersen et al., 1987); (2) estimation
of the yield of DPX (pmol/mg DNA) that would be formed
in the liver of a male B6C3F, mouse if the liver of the mouse
177
ADDUCT DOSIMETRY OF DICHLOROMETHANE IN HEPATOCYTES
Two approaches to estimate human risk based on DPXhvCT
TABLE 2
Cancer Risk Assessments Based on Maximal Estimates of are shown on the right side of Fig. 5. One approach uses
DNA-Protein Crosslinks in the Liver of a GSTT1 -Positive Human the linearized multistage (LMS) model, which until recently
Exposed to Dichloromethane Using Linearized Multistage (LMS) was the preferred method of the U.S. EPA (1986). The other
and Benchmarkd Dose (BMD) Methods"
approach is based on the benchmark dose (BMD) method
(Crump, 1984), which was proposed by the U.S. EPA (1996)
Airborne DCM concentration (ppm)
as an improvement on the earlier procedure.
Both approaches begin with an empirical relationship beRisk parameter
10
30
100
tween the "extra risk" of liver tumors in male B6C3F, mice
LMS method
exposed to 2000 or 4000 ppm of DCM in a carcinogenicity
bioassay (Mennear et al., 1988) and the concentrations of
Predicted AUQ,™, for DCM
in humans* (JIM X hr)
3.9
12.8
64.2
DPXi.ve,. in male mice exposed to the same concentrations
Predicted maximal DPXu,,,
of DCM for 6 hr. The empirical relationship
4
3
in humans' (pmol/mg DNA)
Lifetime risk' (MLE)
UCL on lifetime risk**
Adjusted MLE'
Adjusted UCL'
2.9 X 10"
5 x 10"6
9x10-*
4 X 10"6
7X10"*
9.6 x
2 X
3 x
1X
2 x
10-*
10"'
10"5
10"'
10"'
4.85 x
9 x
2 x
7x
1x
10~
10"'
lO"4
10"'
lO"4
BMD method
ED,,/ = 5.76 pmol/mg DNA (equivalent to - 2 1 0 0 ppm)
LED,/ = 3.24 pmol/mg DNA (equivalent to - 1 5 7 0 ppm)
Upper bound on lifetime risk*
Adjusted upper bound on risk*
Margin of exposure'
9 X 10"6
7 x 10"6
11100
3 X 10"'
2 x 10"'
3400
1 X 10"4
1 X 10670
" Risk calculations were based on predicted values for DPXbVCT in humans,
although DPX were not detectable in human hepatocytes. The unit risk
based on DPX Ure as the measure of exposure was assumed to be the same
as that in mice [3.25 X 10~2 (pmol/mg DNA)"1] (Casanova et al, 1996).
The corresponding unit risk (in units of per ppm) is 1 X 10"6 ppm"' (see
footnote 2).
* Calculated using a version of the PBPK model of Andersen et al. (1987)
appropriate for a 70-kg human (R. Conolly, personal communication).
' Predicted using extended PBPK model for DCM in mouse liver (Casanova et al, 1996), assuming that DPX,,«, in humans = DPX Um in mice X
1/7 at the same time-weighted-average concentration of DCM in the liver
of both species.
d
Risk estimated for exposure to DCM occurring 6 hr/day, 5 days/week.
' Adjusted risk for exposure occurring 8 hr/day, 5 days/week, 48 weeks/
year, for 45 years.
•'Calculated using Eq. (1) assuming q\ = 1.83 X 10"2 (pmol/mg DNA)"1)
(see text) and extra risk = 0.1.
' Calculated using Eq. (1) assuming q\* = 3.25 X 10"2 (pmol/mg DNA)~'
(see text) and extra risk = 0.1
* Based on linear extrapolation from LED,0 to zero dose, zero risk.
' The margin of exposure (dimensionless) is the ratio of the LED,0 (pmol/
mg DNA) to the exposure concentration of interest expressed in the same
units.
contained the same time-weighted-average concentration of
DCM (Casanova et al., 1996); and (3) adjustment of the
calculated DPX|iver in mice for the assumed amount of DPX
that might be formed in humans (multiplication of mouse
DPX|iVCT by 1/7). These steps are shown on the left side of
Fig. 5. Calculated AUChVCT and predicted maximal DPXliver
values in humans at three airborne concentrations of DCM
(10, 30, and 100 ppm) are shown in Table 2.
extra risk =
= 1 - exp(-,;D).
(1)
was found to apply in the observable range of concentrations
that induced measurable increases in tumor yields and DPX
in mouse liver [see Fig. 4b of Casanova et al. (1996)]. In
Eq. (1), p(D) is the probability of tumors at dose D [D =
DPXhvcr (pmol/mg DNA)], p(0) is the spontaneous liver tumor incidence observed in unexposed male mice, and q\ is
the dose coefficient [(pmol/mg DNA)" 1 ].
The maximum likelihood estimate (MLE) of q\ and the
upper 95% confidence limit (UCL) on this estimate (denoted
by q\* and defined as the "unit risk") for mice were calculated using the program GLOBLU 86 (December 1990 version; Clement International Corp., K. S. Crump Division,
Ruston, LA) (see footnote d of Table 6 of Casanova et al.,
1996). For ease of reference, the values of both constants
are repeated here: q\ = 1.83 X 10~2 (pmol/mg DNA)"';
q\* = 3.25 X 10~2 (pmoVmg DNA)" 1 . 2
Substitution of the predicted value of DPX|,ver in humans
in Eq. (1) permits the estimation of the "lifetime risk" of
DCM-induced liver cancer for humans (with the same metabolic characteristics as subjects H-l and H-2) who are exposed to DCM for 6 hr/day, 5 days/week, based on the LMS
2
The unit risk (^i*) in units of (pmol/mg DNA) ' can be converted to
units of ppm"' as follows. The pharmacokinetic results obtained for mice
(Casanova et al., 1996) suggest that the yield of DPX is almost linearly
proportional to the exposure concentration at very low DCM concentrations.
Therefore from row 2 of Table 2, assuming a similar proportionality for
humans, the relationship between D P X ^ (pmol/mg DNA) and the exposure
concentration of DCM (ppm) at low airborne concentrations a 3 X 10"'
pmol/mg DNA per ppm. The unit risk for mice in units of (pmol/mg DNA)~'
is assumed to be identical for humans, i.e., 3.25 x 10~2 per pmol/mg DNA.
Hence, the unit risk in units of ppm"' for humans based on DPX,lwr as the
measure of exposure is 3.25 X I0" 1 x 3 x I0" 5 a 1 X 10"6 per ppm.
This unit risk is approximately 1600-fold smaller than that calculated by
the U.S. EPA (1994) using the PBPK model-estimated amount of DCM
metabolized via the GST pathway (Andersen et al., 1987) and adjusting
the dose for humans with a surface area correction factor.
178
CASANOVA, BELL, AND HECK
model (Table 2). This calculation assumes that the same
equation [Eq. (1)] applies to humans as to mice, i.e., that
the unit risk for mice in (pmol/mg DNA)" 1 is the same as
that for humans. This assumption is not necessarily valid;
i.e., a given quantity of DPXnvCT may or may not represent
the same risk for humans as for mice. Thus, humans may
differ from mice not only in the dose delivered to DNA
(assumed to be 1/7, a pharmacokinetic difference), but also
in the consequences that might result from delivery of that
dose (a possible pharmacodynamic difference). Whether humans and mice differ in the latter sense is unknown, but in
the absence of evidence to the contrary, it is usually assumed
that humans and mice are equally sensitive at a given delivered dose.3
The lifetime risk can be adjusted if the duration of exposure is shorter than 70 years. For example, if the exposure
occurs daily in the workplace (8 hr/day, 48 weeks/year) for
45 years [assumed to be one working life (U.S. Occupational
Safety and Health Administration, 1987)], the correction factor would be 8/6 X 48/52 X 45/70 = 0.79 (Fig. 5). The
values of the adjusted risk at three exposure concentrations
of DCM are shown in Table 2.4
In the BMD method, the effective dose corresponding to
a 10% response (ED,0) or the lower 95% confidence limit
on the effective dose corresponding to a 10% response
(LEDIO) can be calculated from Eq. (1) by solving for
DPXUver using either q\ or q\*, respectively, as the dose
coefficient when the extra risk is equal to 0.1. Values of
ED| 0 and LED, 0 for male B6C3F, mice exposed to DCM are
given in Table 2. In the revised risk assessment guidelines of
the U.S. EPA (1996), the LEDIO is taken as a "point of
departure" from which a straight line is extrapolated to zero
dose, zero risk for genotoxic carcinogens (Fig. 5). The linear
extrapolation can be used to calculate the upper bound on
the risk at any given dose at low doses. As shown in Table
2, the upper bounds on the risk at 10, 30, and 100 ppm are
3
The assumption that the unit risk in units of (pmol/mg DNA) ' is the
same for humans and mice is probably a poor one. The spontaneous incidence of liver cancer in male B6C3F, mice is over 40% (Mennear et ai,
1988), which is much higher than the incidence of liver cancer in humans
[<l%(Peters, 1976; Young and Pollack, 1982)]. Thus, mice are genetically
predisposed to develop liver rumors, and potentially mutagenic compounds
that can react with DNA may initiate cell transformation more readily in
mouse liver than in human liver.
4
Alternatively, instead of adjusting the risk based on the duration of
exposure, a time-averaged dose can be calculated. The latter method was
used by the U.S. EPA in its recent risk assessment of formaldehyde, although adverse effects from HCHO are known to be associated with the
dose rate (intensity of exposure) much more closely than with the total
dose (Hernandez et ai, 1994). Time-averaging the dose has the disadvantage
that potentially important nonlinear pharmacokinetic phenomena may be
obscured. Therefore, we have chosen to maintain the dose unchanged and
to adjust the risk based on the duration of exposure.
essentially identical to the UCL on the risk estimated by the
LMS method.5
Another parameter that can be calculated using the BMD
method is the margin of exposure (MOE), which is defined
as the ratio of the LEDio to the exposure concentration at
any given concentration of interest (U.S. EPA, 1996). MOEs
at 10, 30, and 100 ppm are shown in Table 2. According to
the U.S. EPA (1996), knowledge of the margin of exposure
could be of benefit to risk managers attempting to decide
whether a given exposure concentration represents a significant risk. The MOE values shown in the table apply, of
course, only to subjects having metabolic characteristics similar to those of subjects H-l and H-2.
CONCLUSIONS
The present studies have shown that human hepatocytes,
like those of mice, rats, and hamsters, can metabolize DCM
to HCHO, and that one result of HCHO generation is the
formation of RFA. The ability of human cells to metabolize
DCM is dependent on the presence of functional GSTT1
genes. In human cells that carry this gene, the yields of RFA
were about 7-fold lower than the yields obtained with mouse
cells. In rat and hamster hepatocytes the yields of RFA were
approximately 4- and 14-fold lower, respectively, than in
mouse hepatocytes. Similar results were obtained by Reitz
et al. (1989) for the activities (relative to that of mice) of
GST with respect to DCM in liver cytosolic preparations
from humans, rats, and hamsters.
The amount of HCHO bound to RNA relative to the
amount bound to DNA in mouse hepatocytes depends on
whether the HCHO is generated internally by metabolism
of DCM or is added externally to cells. The RFA:DPX ratio
was about 9.0 ± 1.4 when HCHO was produced by metabolism of DCM, but it was 1.1 ± 0 . 3 when HCHO was added
directly to cells. This difference indicates that HCHO produced from DCM is not equivalent to HCHO added directly
to cells. Therefore, to assume that the toxic effects of HCHO
added externally to cells are predictive of those induced by
HCHO when it is generated internally by metabolism may
be incorrect.
Only mouse hepatocytes were able to form detectable
amounts of DPX. DPX were not detectable in GSTT1 -positive human cells, even at aqueous DCM concentrations
equivalent to an exposure concentration > 10,000 ppm. The
RFA:DPX ratio in human cells was greater than that in
mouse cells. If DPX were formed at all in human cells, the
amounts were very much lower than in mouse cells.
3
According to the U.S. EPA (1996), the linear extrapolation procedure
(BMD method) gives upper bounds on the potential risk at low doses that
are not significantly different from the UCLs predicted by the LMS method.
The same conclusion was reached by Krewski et al. (1984). The results in
Table 2 are consistent with these statements.
ADDUCT DOSIMETRY OF DICHLOROMETHANE IN HEPATOCYTES
Whether or not DPX are involved in the carcinogenic
mechanism of DCM, they can be used as a dosimeter for
risk assessment (Hernandez etai, 1994). A quantitative relationship between mouse liver tumors and DPX|iVCT has been
reported [Eq. (1)] (Casanova et ai, 1996). Assuming that
DPX are formed in GS777-positive human liver, that the
amount of DPX in human liver is as high as one-seventh of
that in a mouse liver with the same time-weighted-average
concentration of DCM, and that the relationship between
liver tumors and DPXhver observed in mice also applies to
humans, the risk of liver cancer to humans with the same
metabolic capabilities as those in the present study can be
assessed using predicted values of DPXllver as an internal
dosimeter. Since the predictions of DPXllver are maximal
values, and since humans are far less likely than B6C3F,
mice to develop hepatocellular carcinomas, the resulting risk
assessments should provide an upper limit on the likelihood
of liver cancer induction in people with similar metabolic
characteristics.
ACKNOWLEDGMENTS
We thank Mr. Donald F. Deyo for excellent technical assistance, Ms.
Sara D. Held for instruction in hepatocyte isolation, Dr. Barbara Kuyper
for editorial suggestions, and Drs. Rory B. Conolly and Gregory L. Kedderis
for performing the PBPK model simulations.
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