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Biochemical Genetics, Vol. 42, Nos. 3/4, April 2004 (°
High Genetic Variability of Esterase Loci in Natural
Populations of Parus major, P. caeruleus, and P. ater
Simon Driesel,1 Lutz Bachmann,2 Michael Stauss,1 Gernot Segelbacher,3
Doris Flach,1 Jürgen Tomiuk,1,4 and Jost Kömpf1
Received 31 December 2002—Final 7 March 2003
In Parus major, P. caeruleus, and P. ater the genetic variation of 16 isozyme loci
was determined. The focus was on esterases that show high phenotypic variation
in natural populations of these species. The degree of heterozygosity of the “nonesterase” loci was 0.029 ± 0.008 (P. major); 0.023 ± 0.012 (P. caeruleus), and
0.034 ± 0.034 (P. ater). Including the esterase loci with up to six alleles per
locus the overall degree of heterozygosity increased to 0.130 ± 0.056 (P. major);
0.143 ± 0.067 (P. caeruleus), and 0.194 ± 0.090 (P. ater). We explain the high level
of variability of esterases by gene amplification and subsequent selection for high
allelic heterogeneity. Substrate specificity of loci is assumed to allow for multiple
resistance against various toxic components. Large allelic variation of esterases,
therefore, increases the fitness of Parus species and allows for utilizing new food
resources.
KEY WORDS: enzyme polymorphism; esterase; passerine birds; isoelectric focusing; ecotoxicology.
INTRODUCTION
Descriptions of protein polymorphisms in humans (Harris, 1966) and in natural
populations of Drosophila pseudoobscura (Hubby and Lewontin, 1966) initiated
two decades of intensive population genetic research on protein variability such as
1
Division of General Human Genetics, Institute of Anthropology and Human Genetics, University of
Tübingen, Wilhelmstrasse 27, D-72074 Tübingen, Germany.
2 Department of Zoology, Natural History Museums and Botanical Garden, University of Oslo, Oslo,
Norway.
3 Max Planck Research Centre for Ornithology, Vogelwarte Radolfzell, Schloss Möggingen, Radolfzell,
Germany.
4 To whom correspondence should be addressed; e-mail: juergen.tomiuk@uni-tuebingen.de.
109
C 2004 Plenum Publishing Corporation
0006-2928/04/0400-0109/0 °
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the analyses of temporal and spatial dynamics of populations in various species.
Furthermore, comparative studies of enzyme variability gave insight into speciation
processes (e.g., Ayala, 1976).
The neutral theory of molecular evolution (Kimura, 1983) can explain the high
level of protein polymorphism observed in natural populations without selection
(see Nei and Graur, 1984). However, for some loci selective mechanisms affecting
the fitness of individuals and/or populations have to be assumed (e.g., Nevo, 1978).
Transferrins, esterases, and peptidases can serve as examples. The large phenotypic
variation of these enzymes/proteins in many animals is expected to reflect selection
for different activity, structure, and/or substrate specificity of the different allelic
products; in short, adaptation to natural environmental conditions (e.g., Hiraizumi
et al., 1992; Parkash and Yadav, 1993; Smith and Small, 1982).
The products of different esterase genes can have locus-specific functions. In
rats, for example, it is assumed that a particular acetylhydrolase plays a significant
role in the regulation of the platelet-activating factor during the late stages of pregnancy (Matsubara et al., 1997). A significant association between fitness traits and
esterase variants is found in Drosophila mulleri (Lourenço et al., 2001) where a relatively high adaptability of genotypes can be related to the presence of a slow allele
at the esterase-4 locus. A classical example is insecticide resistance of peach-potato
aphids (Myzus persicae) that depends on esterase activity (Devonshire, 1977). Indeed esterases are important detoxifying enzymes when utilising specific natural
resources. These few examples may illustrate the importance of esterase functions
on individual development and fitness in a particular habitat. Furthermore, esterases
that indicate exposure of an organism to particular environmental contaminants
may also be used as biomarkers in ecotoxicology (Walker, 1995).
In this paper we present a comprehensive analysis of allozyme variation in
Parus major, P. caeruleus, and P. ater. We analyze in particular esterases that might
as in other species (e.g., Parkash and Yadav, 1993) show high phenotypic variation
in natural populations of Parus species as well.
MATERIAL AND METHODS
Individuals of Parus major (214 adults, 102 families, 698 offspring), P. caeruleus
(197 adults, 99 families, 712 offspring), and P. ater (24 adults, 10 families, 65
offspring) sampled from populations of a large forest area near Tübingen (48◦ 33’N,
9◦ 00’E), South-West Germany during two breeding seasons in 1999 and 2000 were
analyzed.
The birds were caught in their nest boxes, and blood samples (about 50 µL)
were obtained through tapping the Vena ulnaris. Blood was stored in 250 µL
EDTA-buffer in plastic vials and frozen until further processing. Protein electrophoreses were performed on 10 enzymes (16 isozyme loci): lactate dehydrogenase (LDH, 1.1.1.27; 2 isozymes), malate dehydrogenase (MDH, 1.1.1.37; 2
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Fig. 1. Esterase banding patterns of individuals of P. major obtained by PAGIF and subsequent staining
with α-naphthyl acetate (see Appendix; EST-1, EST-2, and EST-3 cannot be discriminated).
isozymes), phosphogluconate dehydrogenase (PGD, 1.1.1.44), NADH-diaphorase
(NADH-DIA, 1.6.2.2), glutamate oxaloacetate transaminase (GOT, 2.6.1.1; 2
isozymes), adenylate kinase (AK, 2.7.4.3), uridine monophosphate kinase (UMPK,
2.7.4.4), phosphoglucomutase (PGM, 2.7.5.1; 2 isozymes), carboxylesterase (EST,
3.1.1.1; 3 isozymes), phosphoglucose isomerase (PGI, 5.3.1.9). The detailed
recipes for electrophoreses and staining procedures are given in the Appendix.
Esterases were electrophoresed in polyacrylamide gels and subsequently
stained with the substrates 4-methylumbelliferyl acetate and α-naphthyl acetate in
order to genotype these enzymes. Using the substrate α-naphthyl acetate, a large
and complex variation of esterase patterns was observed (Fig. 1). The genetics of
these phenotypes could only be explained by the presence of three different loci
(EST-1, EST-2, and EST-3) in each Parus species. The fluorescent locus EST-1
(staining with 4-methylumbelliferyl acetate) was clearly identified by its substrate
specificity whereas family analyses were mandatory to associate the remaining
bands with allelic variation of loci EST-2 and EST-3. The patterns indicate a
monomeric structure of all three isozymes. The substrate specificity of EST-1 suggests homology of this locus among the three species. However, isozyme notation
of EST-2 and EST-3 is arbitrary and does not indicate homology.
The computer programs GENEPOP (Raymond and Rousset, 1995), CERVUS
(Marshall et al., 1998 Slate et al., 2000), and SAS (SAS Institute, 1987) were used
for statistical data analyses.
RESULTS
Allelic variation was studied at 16 isozyme loci (Table I). All isozyme phenotypes
except those of the esterases could be genotyped easily in accordance with the
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Table I. Allele Frequencies of 16 Enzyme Loci in Populations of P. major, P. caeruleus, and P. ater
Locus
na
Hexp
Hobs
PIC
AEP
Allele frequency
f0
m
P. major
LDH-1
LDH-2
MDH-1
MDH-2
PGD
1
1
1
1
2
—
—
—
—
0.038
—
—
—
—
0.038
—
—
—
—
0.037
—
—
—
—
0.019
—
—
—
—
−0.003
26
26
14
14
52
NADH-DIA
GOT-1
GOT-2
AK
1
1
1
2
—
—
—
0.089
—
—
—
0.091
—
—
—
0.083
—
—
—
0.042
—
—
—
—
10
26
26
22
UMPK
PGM-1
1
2
—
0.060
—
0.061
—
0.057
—
0.029
—
—
10
33
PGM-2
EST-1
1
6
—
0.521
—
0.550
—
0.491
—
0.313
—
−0.029
33
209
EST-2
6
0.620
0.633
0.563
0.370
−0.007
207
EST-3
6
0.573
0.576
0.519
0.331
−0.009
205
PGI
4
0.184
0.182
0.173
0.090
1.000
1.000
1.000
1.000
0.981
0.019
1.000
1.000
1.000
0.955
0.045
1.000
0.970
0.030
1.000
0.655
0.175
0.081
0.072
0.012
0.005
0.536
0.288
0.087
0.046
0.036
0.007
0.597
0.249
0.083
0.051
0.015
0.005
0.899
0.082
0.012
0.007
0.038
214
P. caeruleus
LDH-1
LDH-2
MDH-1
MDH-2
PGD
1
1
1
1
2
—
—
—
—
0.082
—
—
—
—
0.083
—
—
—
—
0.077
—
—
—
—
0.038
—
—
—
—
—
26
26
14
14
24
NADH-DIA
GOT-1
GOT-2
1
1
1
—
—
—
—
—
—
—
—
—
—
—
—
—
—
—
10
14
14
1.000
1.000
1.000
1.000
0.958
0.042
1.000
1.000
1.000
(Continues)
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Table I. (Continued)
Locus
na
Hexp
Hobs
PIC
AEP
Allele frequency
f0
m
AK
2
0.121
0.125
0.110
0.055
—
16
PGM-1
PGM-2
EST-1
1
1
2
—
—
0.457
—
—
0.559
—
—
0.352
—
—
0.176
—
—
−0.102
22
22
195
EST-2
4
0.680
0.738
0.626
0.425
−0.057
195
EST-3
5
0.740
0.728
0.692
0.494
0.006
195
PGI
5
0.070
0.072
0.069
0.036
0.937
0.063
1.000
1.000
0.649
0.351
0.467
0.233
0.200
0.100
0.349
0.249
0.197
0.197
0.008
0.964
0.013
0.013
0.008
0.002
−0.009
195
P. ater
LDH-1
LDH-2
PGD
1
1
3
—
—
0.307
—
—
0.333
—
—
0.269
—
—
0.148
—
—
—
20
20
9
NADH-DIA
GOT-1
GOT-2
PGM-1
PGM-2
EST-1
1
1
1
1
1
4
—
—
—
—
—
0.754
—
—
—
—
—
0.500
—
—
—
—
—
0.678
—
—
—
—
—
0.477
—
—
—
—
—
—
9
14
14
10
10
14
EST-2
4
0.779
0.583
0.699
0.500
—
12
EST-3
4
0.492
0.500
0.414
0.240
—
14
PGI
1
—
—
—
—
—
16
1.000
1.000
0.833
0.111
0.056
1.000
1.000
1.000
1.000
1.000
0.357
0.286
0.179
0.179
0.292
0.250
0.250
0.208
0.678
0.250
0.036
0.036
1.000
Note. n a – number of alleles; Hexp and Hobs – expected and observed degrees of genetic heterozygosity;
PIC – polymorphism information content; AEP – average exclusion probability; f 0 – frequency of
null alleles; m – sample size.
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electrophoretic patterns described by Harris and Hopkinson (1976). Using the
99% criterion (Hartl and Clark, 1989) the degree of polymorphism of the “nonesterase” loci ranged between 0.11 and 0.31 (0.308 P. major; 0.250 P. caeruleus;
0.111 P. ater). The observed and expected degrees of heterozygosity (Hobs and
Hex p , respectively) were relatively low in all species (Hexp ± SE: 0.029 ± 0.015
P. major; 0.023 ± 0.012 P. caeruleus; 0.034 ± 0.034 P. ater). As a consequence the
average parental exclusion probability of “non-esterase” loci was low amounting
to about 15%.
The esterases showed high phenotypic and genetic variability in all three
Parus species (Fig. 1, Table I). Up to six alleles per locus were observed and the expected degrees of heterozygosity ranged between 0.457 and 0.779 (0.521 ≤ Hexp ≤
0.620 and 0.550 ≤ Hobs ≤ 0.633 for P. major; 0.457 ≤ Hexp ≤ 0.740 and 0.559 ≤
Hobs ≤ 0.738 for P. caeruleus; 0.492 ≤ Hexp ≤ 0.779 and 0.500 ≤ Hobs ≤ 0.583
for P. ater). The probability of excluding one parent was about 0.75 considering
all three loci. Considering all 16 enzymes, the expected degrees of heterozygosity increased considerably to exceed 13% and the average exclusion probability
(AEP) reached ∼80% (0.760 P. major, 0.790 P. caeruleus, 0.831 P. ater). However, in a previous study we showed close linkage between esterase loci (Stauss
et al., 2003) and, therefore, the use of average values for esterase loci seems to
be appropriate (Hexp = 0.067 and AEP = 0.450 for P. major; Hexp = 0.069 and
AEP = 0.444 for P. caeruleus; Hexp = 0.098 and AEP = 0.494 for P. ater).
DISCUSSION
Allozyme variation is frequently used to characterize the genetic structure and
dynamics of natural populations. Genetic variation of birds based on allozyme data
is in the same order of magnitude as in other vertebrates (P = 0.30 and H = 0.05)
(Nevo et al., 1984). Ward et al. (1992) found similar estimates of genetic variability
in bird species (H = 0.07). However, smaller values are described in 4 species of
Strigiformes (P = 0.16 and H = 0.03) and 10 species of Charadriiformes (0.01 ≤
H ≤ 0.04 (Randi et al., 1991). Our results more closely fit with the higher estimates
given by Nevo et al. (1984) and Ward et al. (1992): i.e., in the three Parus species the
average degree of polymorphism is about 30% and the average degree of genetic
heterogeneity is close to 7% (using average values for the three esterase loci).
However, estimates of genetic heterozygosity and polymorphism can be biased,
since high variability of only a few loci can considerably alter overall estimates
as in that of the esterase loci in the present study. In fact we found that the degree
of heterozygosity and the number of alleles with relatively high frequencies at
esterase loci are within the range of microsatellites.
The effects of genetic variability of enzyme loci on the fitness of individuals
and populations have been extensively discussed. Genetic heterogeneity can have
a general impact on the fitness of individuals as has been shown by theoretical
studies (Berger, 1976; Charlesworth, 1991; Ohta, 1971; Turelli and Ginzburg,
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1983). It is therefore not surprising that in some case studies a strong correlation
of allozyme variability and fitness parameters was observed (e.g., Allendorf and
Leary, 1986; David et al., 1995; Koehn et al., 1988; Mitten et al., 1986; Pogson
and Zouros, 1994; Thelen and Allendorf, 2001). In the mosquito Culex pipiens
qualitative and quantitative variation of esterases has been interpreted as a consequence of selection (Guillemaud et al., 1999; Raymond et al., 1989, 1993). Gene
amplification and regulatory mechanisms affect insecticide resistance and fitness
of individuals. The same is true for the aphid Myzus persicae (Field et al., 1988).
In birds, studies on esterase functions and fitness of individuals did not yield
consistent results. In nestling European starlings (Sturnus vulgaris) Parker and
Goldstein (2000) could not find a strict correlation between mortality and different esterases buffering organophosphate and carbamate toxicity. Accordingly,
Sanchez et al. (1997) found no correlation of serum B esterase activities and biometric parameters in the four bird species Sylvia melanocephala, Serinus canaria,
Parus caeruleus, and Erithacus rubecula. In contrast, Fossi et al. (1996) described
interspecies differences in B esterases for seven bird species and correlated low B
esterase activity to large birds and food specialists. Furthermore, cholinesterases
and carboxylesterases showed seasonal, diurnal, and developmental variations in
activity in some birds (Thompson, 1993). The high genetic variation of esterase
loci in the three Parus species studied here can affect individual and population
fitness. High level of structural variability can be explained by gene amplification
and subsequent selection for high allelic heterogeneity. We assume that substrate
specificity can lead to multiple resistance against different toxic components. The
following argument may support our interpretation of selection for high genetic
variation at esterase loci. Under selective neutrality, gene silencing is expected and
null alleles at duplicated esterase loci should be common (Nei and Roychoudhury,
1973). Our estimates of very low null allele frequencies, however, do not support
such an evolutionary scenario (Table I). Indeed further studies show that allelic
variation of esterase loci may have profound effects on the fitness of P. major
females depending on the habitat quality (Stauss et al., in press). Therefore, we
propose that gene duplications and the large allelic variation of each esterase locus has evolved and has been maintained in Parus species through adaptation to
environmental toxicity, e.g., food resources.
APPENDIX
Electrophoretic methods for the detection of enzyme variability in three Parus
species.
Electrophoreses
1. Horizontal starch gel electrophoresis: 14% Connaught starch using a discontinous system (0.4 M citric acid–NaOH with pH 6.2 in the traces and
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30 mM Histidine/HCl–NaOH with pH 6.0 in the gel), 7 V/cm, 16 h,
8◦ C.
2. Isoelectric focusing in polyacrylamide gels (PAGIF, LKB Multiphor System): gel size 125 × 260 × 0.5 mm, gel solution (T = 5%, C = 3%):
9.2 mL aqua bidest, 2 g saccharose, 2.5 mL 30%-acrylamide, 1.2 mL
2%-bisacrylamide, 0.9 mL carrier ampholytes (see below), 1 mL 0.5%ammonium persulfate, 0.1 mL 3%-TEMED.
3. Isoelectric focusing in agarose gels (AGIF): gel size 125 × 260 × 0.5 mm,
gel solution (1% agarose): 17 mL aqua bidest, 2 g saccharose, 170 mg IEF
agarose (Pharmacia), 1.3 mL carrier ampholytes (see below).
PAGIF of Esterases
a) Parus major
Carrier ampholytes: 400 µL pH 4–6.5 (Pharmalyte), 200 µL pH 5–7
(Fluka), 200 µL pH 4–6 (Fluka), 100 µL pH 6–8 (Pharmalyte); anolyte:
0.1 M H3 PO4 , catholyte: 0.5 M NaOH; electric settings: 1500 V, 10 mA,
5 W, 30 min prefocusing; 7.5 µL sample in application pieces located
1.5 cm from the cathodal strip; focusing until constant voltage.
b) Parus caeruleus
Carrier ampholytes: 200 µL pH 4–6 (Fluka), 300 µL pH 5–7 (Fluka),
200 µL pH 4–6.5 (Pharmalyte), 200 µL pH 5–6.5 (Sigma); anolyte 0.1 M
H3 PO4 , catholyte 0.1 M NaOH; other settings as described for P. major.
c) Parus ater
Carrier ampholytes: 200 µL pH 3.5–10 (LKB), 300 µL pH 4–6
(Fluka), 300 µL pH 5–8 (Fluka), 100 µL pH 4–9 (Fluka); other settings
as described for P. caeruleus.
AGIF of Phosphoglucose Isomerases (All Species)
Carrier ampholytes: 650 µL pH 7–9 (Pharmacia), 650 µL pH 8–9.5 (Fluka);
anolyte 25 mM glutamine-asparagine acid, catholyte 0.5 M NaOH; settings:
1500 V, 25 mA, set power so that the corresponding initial voltage is 300 V;
1 h prefocusing; 7.5 µL sample in application pieces located 1.5 cm from anodal strip; focusing is finished when the cathodally migrating hemoglobin fraction
forms a sharp band.
Electrophoreses of Other Enzymes
The enzmyes AK, UMPK, GOT-1, GOT-2, LDH-1, LDH-2, MDH-1, MDH-2 were
separated by starch gel electrophoresis. PGM and PGD were electrophoresed by
PAGIF using pH-gradient 3.5–10 (LKB). NADH-DIA as well as GOT-1 were
separated by AGIF using pH-gradient 3.5–10 (LKB).
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Staining and Zymograms of Esterases
1. UV-fluorescent staining with 4-methylumbelliferyl acetate (MUA). Staining solution: 5 mg MUA, 0.5 mL acetone, 1.7 mL 0.25 M Na-acetate-acetic
acid pH 4.6. Application with cellulose-acetate foils (50 × 260 mm). After
10 min in a moist chamber at 30◦ C fluorescent bands appear on the foil
under UV (312 nm).
2. Staining of gels with α-naphthyl acetate and Fast Blue RR salt. Staining
solution: 30 mg α-naphthyl acetate in 1 mL acetone and 1 mL H2 O, 100 mg
Fast Blue RR in 4 mL H2 O, 100 mL 125 mM Tris-Histidine/HCl pH
7.4. Bands develop while shaking at room temperature within 30 min.
After washing the gels with water the gels were immersed for 1 h in 5%
glycerol, and dried at 30◦ C (use of β-naphthyl acetate, naphthyl butyrate,
and naphthyl propionate as substrates did not alter the banding patterns).
Staining and zymogram procedures of all other enzymes under study were done
as described by Harris and Hopkinson (1976).
ACKNOWLEDGMENTS
We thank Jochen Blank for his assistance in the field and an anonymous reviewer
for his comments improving a previous version of this manuscript. This work was
supported by a grant from the German Research Foundation No. DFG To 151/21 and the German Academic Exchange Service DAAD No. 13/PPP-N1. G.S.
was supported by a grant of the Max-Planck Research Centre for Ornithology.
L. Bachmann was supported by a grant from the Research Council of Norway
(National Centre for Biosystematics, 146515/420).
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