Journal of medical and applied macology revista de macologia

J. Med. 6 Appl. Malacol., 1994: 6: 00-00
XENOBIOTIC METABOLIZING ENZYMES OF THE FRESHWATER SNAILS
PLANORBELLA DURYI AND LYMNAEA (RADIX) NATALENSIS
Y. S. Naikl, T. Magwere, M. Matiyenga and J. A. Hasler2
ABSTRACT
Very little is known about the ability of snails to metabolise and remove xenobiotics
such as molluscicides. The present study was conducted to determine whether the
freshwater snails Planorbella duryi and Lymnaea (Radix) nntaIensis possess two of the
major enzyme systems of detoxication, namely the P450 monooxygenase system and the
glutathione Stransferases. We were able to measure microsomal cytochrome b5 and
NADPH cytochrome c reductase activity, indicating a functional mixed function oxidase
system. However, probing with three known substrates for the mammalian enzyme, we
were unable to detect any cytochrome P450 mediated activity. Glutathione Stransferase
activity was detectable in cytosolic fractions. In general, the activities detectable in snails
were much lower than in mammalian liver preparations. The existence of these enzymes
in snails suggests that studies should be undertaken to observe the interaction between
these enzymes and xenobiotics such as candidate molluscicides.
Key words: Planorbella duyi, Lyrnnaea (Radix) natalensis, cytochrome P450, glutathione S
transferase.
INTRODUCTION
x
Xenobiotic metabolizing enzymes (XME) play an important role in the metabolism and
removal of potentially harmful compounds (Jakoby & Ziegler, 1990). The cytochrome P-450
dependant monooxygenases convert hydrophobic compounds to water soluble compounds
(Guengerich, 1990), while the glutathione S-transferases remove potentially harmful electrophilic compounds (Mannervik & Danielson, 1988). These two families of enzymes are distributed widely in nature (Nelson et al., 1993: Beutler & Eaton, 1992) and it is thought that
they have been retained during evolution to cope with an ever increasing diversity of compounds encountered by aquatic and terrestrial animals (Nelson e f al., 1993; Beutler & Eaton,
1992).
An increasing number of aquatic organisms have been identified which possess these
enzyme systems. The enzyme systems of aquatic molluscs appear to be inducible by foreign
compounds, a property which they share with their mammalian counterparts (Livingstone,
et al., 1989). Aquatic mollusc species have also been used to assess environmental water
pollution in studies similar to those conducted using fish (Livingston
1987). Although information on marine molluscs is increashg, relat
about the xenobiotic metabolizing enzymes of freshwater snails, although recently there has
been a report on the XME of the freshwater snail Lymnaea stagnalis (Linnaeus) Hubendick,
1951) (Wilbrink et al., 1991a; Wilbrink et al., 1991b). Knowledge of the existence of these
enzyme systems is important for two reasons. Firstly, freshwater snails are known to transmit a number of parasitic diseases in tropical countries. Since chemical means are used to
eradicate these snails and since these chemicals are likely to be metabolised by the XME, it is
possible that resistance to these chemicals may be mediated by the XME. Secondly, these
aquatic snail species may be used to monitor water pollution. The demonstration of the
existence of the enzymes in these snail species would therefore open the way for their development as possible biomarkers to monitor water pollution.
We report here on some of the enzymes of two freshwater snails, namely Planorbell@ryi
(Wetherby) and Lymnaea (Radix) nafalensis (Gauss). Two rodent species have been included
for comparison.
'Department of Biochemistry, University of Zimbabwe, P. 0.Box MP167 Mount Pleasant, Harare,
Zimbabwe.
2Author to whom correspondence should be sent.
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Naik, Magwere, Matiyenga and Hasler
2
MATERIAIS AND METHODS
Chemicals
All biochemicals were obtained from the Sigma Chemical Co., St. Louis, Missouri, U.S.A.
Ethoxyresorufin was synthesized according to the method of Prough et al. (1978).
Rats and Mice
Eight-week old male Sprague Dawley rats or BALB/c mice were allowed standard laboratory chow
and water ad libitum and were exposed to natural day-night cycles. Animals were sacrificed by cervical
dislocation. The livers were perfused in situ with ice-cold saline until they had blanched, then were
removed, blotted dry and weighed. They were then homogenised in 5 volumes of 0.05 M Tris-HC1
buffer, pH 7.4 containing 0.15% KC1 (Buffer A) in a Potter-Elvehjem homogenizer using a teflon pestle.
The homogenate was treated as described below.
Snails
Planorbella duryi or Lymnaea natalensis were originally obtained from the field and cultured in
outdoor cement aquaria for a period of over a year before study. The volume in each aquaria was
maintained at approximately 10 litres each. The daily temperatures of the water in the aquaria during
the experimental period (May-July)ranged from 10°C in the early morning to about 20°C in the afternoon. The diet of the snails consisted of only fresh locally grown garden lettuce.
For each experiment, a single pool of 48-60 young adult snails was used. On the day of the sacrifice,
snails were washed three times with tap water to remove soil and leaf particles. The snails were left in
clear distilled water for about 10 minutes in order to allow the snails to pass out as much green faecal
matter as possible. This green particulate matter normally contaminates the microsomal and cytosolic
fractions and this procedure was undertaken to help minimize it. After this period, the snails were
again washed twice with distilled water.
Each snail was removed from its shell using forceps. The intestines were immediately excised and
removed. The remaining body of the snail was quickly rinsed in icecold Buffer A. The soft tissues of
the snail thus obtained were pooled and then homogenised in Buffer A using a Potter-Elvehjern
fitted with a teflon pestle. The homogenate was treated as desaibed below.
Specimens (shells) of Planorbella d w y i and Lymnaea (Radix) natalensis have been deposited in the
Museum of Zoology, University of Michigan, cat. nos. 353640 and 353637, respectively.
.-Preparation of microsomal and cytosolic fractions
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Liver homogenates were centrifuged at 9,000 x g for 15 minutes and the resultant supernatant was
centrifuged for 60 min at 105,000 x g to obtain microsomes (pellet) or cytosol (supernatant). Whole
snail homogenates were centrifuged at 10.000 x g (Lymnaea,ztaIensis) or 20,000 x g ( H e l ~ d u r y ifor
)
30 minutes and the resultant supernatant in each case was then centrifuged at 105,000 x g for 60 min to
obtain the microsomes (pellet) or cytosol (supernatant).
All microsomal pellets were homogenised in TEG solution (0.1 M Tris-HCl, 1 rnM EDTA and
20%v/v glycerol). Microsomal suspensions and cytosolic fractions were all stored at -70°C until
required.
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Protein was measured according to the method of Lowry, et al. (1951), using bovine serum albumin
as standard. The following assays were performed on microsomal fractions. Cytochrome b5
(Estabrook & Werringloer, 1978), NADPH cytochrome c reductase (Omura & Takesue, 1970), ethoxycoumarin 0-deethylase (ProughLjet al., 1978), ethoxyresorufin Odeethylase (Prough et aL, 1978), pentoxyresorufin 0-deethylase as for EROD except that pentoxyresorufin was used as substrate (Prough et
al., 1978). Glutathione Stransferase activity was measured in cytosolic fractions according to the
method of Habig et al. (1974) at pH 6.5.
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RESULTS
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Table 1 shows the cytochrome b5 (B5) levels, cytochrome c reductase (CRED) activities as
well as the ethoxycoumarin 0-dealkylase (ECOD), ethoxyresorufin 0-dealkylase (EROD)
and pentoxyresorufin 0-dealkylase (PROD) activities in microsomes of snails and rodents
as well as the glutathione S-transferse activity in supernatants of these species.
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Xenobiotic Metabolizing Enzymes of Snails
*
DISCUSSION
Our results indicate that the freshwater snails p ssess the XME found in most other organcontent in snail microsomes since
isms. We were unable to measure the cytochrome
the difference spectrum consistently gave a peak at 420nm and no peak at 450nm. The
reason for this may be explained by the fact that the microsomes in our study appeared to be
heavily contaminated with green pigment, presumably chlorophyll from the lettuce on
which they fed. Wilbrink,;ef al., (1991a) also using a freshwater pond snail, have commented
that feeding snails on a l e h c e diet contaminated their microsomes and hence they provided
an alternative diet. The fact that the snails do possess microsomal B5 and CRED activity
indicates that a functional mixed function oxidase system is likely to operate. However,
probing with known substrates for the mammalian enzyme we were unable to detect any
activity (EROD/PROD/ECOD). This may be due to several possible factors. The method of
microsomal isolation and the assay conditions used by us are those that have been devised
for mammalian systems and may be less than ideal for preparing and probing snail
microsomes. On the other hand, it may be that the enzymes are expressed in amounts too
low for detection. Lastly, the lack of activity may also have been due to the unsuitability of
the chosen substrates for detecting snail enzymes. Although ethoxycoumarin, ethoxyresorufin and pentoxyresorufin are well established substrates for various isoforms of P450 from
mammalian organs they may be less than ideal for the detection of snail enzymes. Wilbrink
ef al. (1991a) also were unable to detect EROD activity but were able to detect PROD activity
as well as aminopyrine N-demethylase activity using Lymnaea stagnalis. Thus, it would seem
unlikely that all three substrates used by us were unsuitable for the detection of snail
enzymes. However, the differences between our study and theirs may also be due to snail
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Naik, Magwere, Matiyenga and Hasler
4
species differences as well as different culture conditions including diet. Clearly, further
studies using more highly purified microsomes and a wider selection of substrates are
needed to further characterise the monooxygenases of the snail species we are studying.
The glutathione S-transferase activity was detectable more easily and appears to be
expressed at higher levels. We have been able to measure significant activity in cytosolic
fractions and partially characterise the crude cytosols using a variety of different substrates
and inhibitors (Naik et al., 1994).
In general, the enzyme activities that we were able to measure were much lower in snail
subcellular fractions than in corresponding rodent fractions. This is in agreement with
those values previously reported for other aquatic organisms such as marine molluscs
(Livingstone et al., 1989).
-
ACKNOWLEDGEMENTS
The authors gratefully acknowledge the financial support of the International Program in Chemical
Sciences, Uppsala University, Sweden and the Research Board, University of Zimbabwe.
LITERATURE CITED
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BEUTLER, T.M. & EATON, D.L. 1992. Glutathione Stransferases: amino acid sequence comparison,
classification and phytogenetic relationship. Environmental Carcinogenesis and Ecotoxicology Reyiews,
C10: 181-203.
ESTABROOK, R.W. & WERRINGLOEg) 1978. The measurement of difference spectra: Application to
ds in Enzymology, 52: 212-220.
the cytochromes of microsomes. M
GUENGERICH, F.P. 1990. Enzymatic oxidation of xenobiotic chemicals. CRC Critical ReoiPws in
Biochemistry and Molecular Biology, 25: 97-153.
HABIG, W.H., PABST, M.J. & JAKOBY, W.B. 1974. Glutathione Stransferase. The first step in mercap
turic acid formation. Journal of Biological Chemistry, 249: 7130-7139.
JAKOBY, W.B. & ZIEGLER, D.M. 1990. The enzymes of detoxication. Journal of Biological Chemistry,
265: 20715- 20718.
LIVINGSTONE, D.R., KIRCHIN, M.A. & WISEMAN, A. 1989. Cytochrome P-450 and oxidative
metabolism in molluscs. Xenobiotica, 19: 1041-1062.
LOWRY, O.H., ROSEBROUGH, N.J., FARR, A.L. & RANDALL, R.J. 1951. Protein measurement with
the Folin Phenol reagent. Jownal of Biological Chemistry, 193: 265-275.
MANNERVIK, B. & DANIELSON, U.H. 1988. Glutathione transferases -structure and catalytic activity. CRC Critical Reviews in Biochemistry, 23: 283-337.
NAIK, Y.S., MAGWERE, T., MATIYENGA,
. 1994. Glutathione Stransferases of the
freshwater snails .klpmwduryi
lensis. Partial charact ' '
d interacG
t
h Niclosamide. J O Y ~ofZ~ e d i c a
press;siGG-)
NELSON, D.R., ET AL. 1993. The P450
seqtiences, gene mappmg, acces
sion numbers, early trivial names of enzymes and nomenclature. DNA and Cell Biology, 12: 1-51.
OMURA, T. & TAKESUE, S. 1970. A new method for simultaneous purilication of cytochrome b5 and
NADPH-cytochrome c reductase from rat liver miaosomes. Journal of Biochemistry, 67: 249-257.
PAYNE, J.F., FANCEY, L.L., RAHIMATULA, A. & PORTER, E.L. 1987. Review and perspective on the
use of mixed-function oxvaenase
enzymes in biolopical
,"
., monitoring.- Comparative Biochemistry and
Physiology, 86C: 233-245.
PROUGH, RA., BURKE, M.D. & MAYER, R.T. 1978. Direct fluorometric methods for measuring mixedfunction oxidase activitv. Methods in Enzwwlom. 52: 372-377.
, & VERMUELEN, N.P.E. 1991a. Occurrence
WILBRINK, M., GROOT,E.J., JANSEN, R.:DE ~ R I E SY.
of a cyto&uoke P-450co~takngmixed-function oxidase system in the pond snail Lymnaea stagnalis.
Xenobiotica, 21: 223-233.
WILBRINK, M., VAN DE MERBEL, N.C.& VERMUELEN, N.P.E. 199Ib. Glutathione Stransferase
activity in the digestive gland of the pond snail Lymnaea stagnalis. Comparative Biochemistry and
Physiology, 99C: 185-189.
Journal of Medical and Applied Malacology, P.0.Box 2715, Ann Arbor, Michigan 48106, U.S.A.
Q International Society for Medical and Applied Malacology, 1994
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TABLE 1. Xenobiotic metabolizing enzyme activities in the freshwater snails PLznorbeZZa d w y i
and Lymnuea (Rndix) nafnZensisand two laboratory rodents.
Rodents
Snails
cytochrome b5*
NADPH cytochrome
P-450 reductase"
2.14
io.05
1
*<
129
f6
M
55
N.D.
0.54
ffl.09
N.D.
178
MO
Ethoxycoumarin
Odealkylase-
0
Ethoxyresorufin
0-dealkylase-
0
0
1.37
ffl.17
Pentoxyresorufin
O-dealkylase*"
N.D.
0
N.D.
Glutathione
.Stransferase""
11.15
ffl.35
*
0.17
ffl.03
N.D.
pmol of cytochromeb5/mg protein.
nmol of cytochrome c reduced /min/mg protein.
*** pmol of product/min/mg protein.
**"
pmol of conjugate/min/mg protein.
N.D. = Not done.
Values are mean f:S.D. for duplicate measurements of 2 4 separate experiments.
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