Journal of Heredity Advance Access originally published online on March 30, 2005
Journal of Heredity 2005 96(4):388-395; doi:10.1093/jhered/esi051
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Genetic Coadaptation of the Amylase Gene System in Drosophila melanogaster: Evidence for the Selective Advantage of the Lowest AMY Activity and of Its Epistatic Genetic Background
From the Department of Biology, Graduate School of Sciences, Kyushu University, 6-10-1, Hakozaki, Higashi-ku, Fukuoka 812-8581, Japan (Araki, Yoshizumi, Inomata, and Yamazaki); Department of Zoology, Oregon State University, 3029 Cordley Hall, Corvallis, OR 97333 (Araki); and Research Institute of Evolutionary Biology, 2-4-28, Kamiyoga, Setagaya-ku, Tokyo 158-0098, Japan (Yamazaki)
Address correspondence to Hitoshi Araki, Department of Zoology, Oregon State University, 3029 Cordley Hall, Corvallis, OR 97333, or e-mail: arakih{at}science.oregonstate.edu.
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Abstract |
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In natural populations of Drosophila melanogaster, an amylase isozyme with the lowest
-amylase activity (AMY1,1) is predominant. To evaluate the selective significance of AMY1,1 and its regulatory factor(s), we examined selection experiments in laboratory populations on two distinct food environments. After 300 generations, AMY1,1 became predominant (89%) in a glucose (a product of AMY)-rich environment, while an isozyme with higher -amylase activity, AMY1,6, became predominant (83%) in a starch (substrate)-rich environment. We found that the identical alleles of the amylase (Amy) gene, which encodes each of AMY1,1 and AMY1,6, were shared between the two populations in the different food environments, employing the nucleotide sequencing of the duplicated Amy genes. Nevertheless, AMY1,6 homozygotes selected in the starch-rich environment had a twofold higher AMY enzyme activity than those selected in the glucose-rich environment, suggesting a coadaptation of the coding region and its regulatory factor(s) on the genetic background. Such a difference in AMY enzyme activity was not detected between AMY1,1 homozygotes, suggesting that the effect of the genetic background is epistatic. Our results indicate that natural selection is working on the Amy gene system as a whole for flies to adapt to the various food environments of local populations.
How organisms adapt to variable environments has long been discussed (Gupta and Lewontin 1982; Kang and Gauch 1996). A direct approach to address this issue is a laboratory experiment in which fitness of individuals is directly estimated under controlled conditions. In Escherichia coli, this approach has been applied to estimate the fitness of mutants in different environments (e.g., temperature and food) and succeeded in detecting significant differences of fitness among mutants (Cooper and Lenski 2000; Remold and Lenski 2001). However, these studies focus on the selective significance of a specific gene or mutation, while differences in a genetic background, which contains unlinked cis and trans regulatory regions, may play an important role in the adaptation of a gene system (Remold and Lenski 2004).
-amylase (AMY), which is encoded by the amylase gene (Amy), hydrolyzes starch in food. Because AMY is required to interact with changes in the food environments in natural populations, Amy is a candidate gene under natural selection. Indeed, several laboratory experiments of Drosophila (De Jong and Scharloo 1976; Hickey 1977; Powell and Andjelkovic 1983) indicated that Drosophila with high AMY enzyme activity has a selective advantage in starch-rich environments. However, their interpretations of the results have been called into question, mainly because they used only a few strains in their experiments and a strong linkage disequilibrium is expected (Doane 1980). The selective significance of this gene is supported by the studies employing recent molecular population genetics and molecular evolutionary approaches (Araki et al. 2001; Inomata et al. 1995b; Zhang et al. 2003). More importantly, the previous selection experiments do not explain why AMY1,1, a haplotype with the lowest AMY enzyme activity in Drosophila melanogaster (Doane 1969), is predominant in natural populations (Hickey 1979; Singh et al. 1982). The symbol AMYX,Y represents a haplotype in which AMYX and AMYY isozymes are encoded by the duplicated Amy genes in this species, Amy-p and Amy-d, respectively. To date, little was known about the selective advantage of AMY with the lowest activity in natural populations.
The gene regulation of Amy in Drosophila is relatively well studied. For example, the responses of AMY enzyme activity to different dietary carbohydrates in food have been reported (Benkel and Hickey 1986; Inomata et al. 1995a; Inomata and Yamazaki 2000; Yamazaki and Matsuo 1984). The cis regulatory factors were identified upstream of each of the Amy coding regions (Choi and Yamazaki 1994; Okuyama et al. 1996), while the presence of trans regulatory factor(s) was suggested but not evidenced (Matsuo and Yamazaki 1997).
In this study, we addressed two questions: whether AMY1,1 has the selective advantage in a product-rich environment, and if so, whether any regulatory factor(s) that is responsible for AMY enzyme activity also adapts to different food environments. We tested these hypotheses directly, performing the laboratory selection experiments in D. melanogaster. Selection was determined for two types of food environments which contain distinct dietary carbohydrates—starch (a substrate of AMY) and glucose (a product of AMY). Matsuo and Yamazaki (1984) indicated that the starch-rich cage population adapted by increasing their plasticity for different sources of dietary carbohydrate rather than by changing the AMY1,1 frequency, using the same starch-rich cage population in a relatively small number of generations. With a new population in the glucose-rich environment, we estimated the AMY1,1 frequencies in the cage populations during the selection experiment for more than 300 generations to test the selective significance of AMY1,1 in longer generation time. After selection, the nucleotide sequences of the Amy gene regions in the samples from the laboratory populations were obtained to see if the two cage populations share the same Amy alleles and its linked regulatory regions. Nucleotide sequences of a microsatellite locus, DROYANETSB, were also obtained to estimate the effective population size in the cage populations. Furthermore, we measured the AMY enzyme activities of the flies, which shared the same Amy alleles but had different genetic backgrounds from the cage populations, to evaluate the effect of selection on the genetic backgrounds that affect AMY enzyme activity. Based on the results obtained, we discuss how possibly flies with the lowest AMY activity can adapt to natural environments and how such adaptations can shape the genetic variation of the organism in natural populations.
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Materials and Methods |
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Cage Population Experiments
The materials used in the cage experiments were the same as those in Matsuo and Yamazaki (1984). In short, experimental materials for the cage experiments were from an original cage population (with normal medium, see below) that was derived from 400 isofemale flies (collected by T. Yamazaki at Akayu, Yamagata, Japan in 1974). The AMY1,1 frequency in this population was estimated as 0.93 (Yamazaki et al. 1984). During the first approximately 150 generations in the establishment of the original cage population, the AMY1,1 frequency in the cage population dramatically decreased, but became stabilized around 0.5 after about 80 additional generations (see Figure 1). We started the selection experiment after an additional 10 generations. Prior to the selection experiment, we examined 200 salivary gland chromosomes from the cage population to confirm that inversions, which potentially change the Amy allele frequency simply by the linkage effect, had been completely eliminated.
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After the 240 generations since the establishment of the original cage population, two different types of experimental cage populations, starch cages and normal cages, were established by sampling approximately 20,000 eggs from the original cage population. After the 312 generations in the original cage population, glucose cages were established from the normal cage by the same method. Each type of cage had two replicates at the starting point, and all six cages were expected to have nearly the same genetic constitution. Unfortunately some cages were accidentally damaged, and they were reestablished from the replicate of the same type of populations (once for the normal cage and twice for both the starch and glucose cages; see Figure 1). All populations were maintained in the same bucket-type cages (diameter 25 cm) at 25°C. Twenty-four generations per year are expected under these conditions. Into each 55 ml vial, 10.0 ml of food was poured, and 30 vials were kept in every cage. The oldest portion of the vials (6 of 30), which remained in a cage for about 20 days, was replaced with a new set every 3 or 4 days so that continuous generations of flies are maintained in the cages.
Composition of Three Types of Media
The components of the media were as follows: normal medium: sugar 14.0 g, molasses 28.0 g, cornmeal 110 g, killed yeast 20.0 g, agar 6.0 g, propionic acid 4.0 ml in 1.0 L of water; starch medium: starch 155 g, killed yeast 20.0 g, agar 3.0 g, propionic acid 4.0 ml in 1.0 L of water; glucose medium: glucose 155 g, killed yeast 20.0 g, agar 6.0 g, propionic acid 4.0 ml 1.0 L of water. Nipagin 1.0% was supplemented in every media as a mold inhibitor. After approximately 150 generations of all the selection experiments (1992), the composition of the starch and glucose media was changed to a carbon source (starch or glucose) 100 g, killed yeast 50.0 g (amounts of agar, propionic acid, and Nipagin were not changed). In the enzyme activity assay, this new composition of the media was also used to breed the flies.
Samples for Sequencing
Isofemale lines were established in 1996 from each of the two starch-rich and two glucose-rich cage populations. The CyO/Pm balancer stock was used to obtain Amy isogenic lines. Ten isogenic lines from each cage, which were homozygous for the second chromosome and free of either lethal or sterility genes, were selected at random for nucleotide sequencing (20 lines per food environment). All methods and primers for sequencing of Amy-p, Amy-d, and DROYANETSB are the same as in Araki et al. (2001). After polymerase chain reaction (PCR) amplification, the PCR products were directly sequenced using an ABI 377 automated DNA sequencer (Applied Biosystems, Foster City, CA).
Nucleotide Data Analysis
We used the ProSeq program version 2.56 (available at http://helios.bto.ed.ac.uk/evolgen/filatov/proseq.html) for editing the sequences and the program CLUSTALX (Thompson et al. 1997) for multiple alignments. Gene diversities (identical to average allele heterozygosities; see Nei 1987) were estimated using the DnaSP program version 3.14 (Rozas and Rozas 1999). The effective population size of the cage population (Ne) was estimated based on the equation
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Samples for the AMY Enzyme Activity Assay
To investigate the effect of selection on AMY enzyme activity and on the genetic background in the flies, we measured AMY activity in AMY homozygous lines established from the cage populations after the selection experiment. Eight AMY1,1 and 10 AMY1,6 homozygous lines were established from the starch cages (S-cage background), and 12 AMY1,1 and 10 AMY1,6 homozygous lines were established from the glucose cages (G-cage background) by the following method. In 2000, we sampled females from the starch cages and the glucose cages to establish isofemale lines. Mating pairs were designed using a single virgin male and a virgin female from the different isofemale lines derived from the same cages. Out of the offspring by these mating pairs, only offspring of parents were identified to be homozygous for the identical Amy gene by the single nucleotide polymorphism (SNP)-based PCR detection method were selected. Because we obtained the nucleotide sequences of the most frequent AMY1,1 and AMY1,6 alleles in the cage populations, we could distinguish these alleles by the PCR method specifically designed for each allele. Because the Amy-p locus encoded only the AMY1 isozyme and most of the Amy-p alleles were perfectly linked to the Amy-d alleles in our samples (see Results), we focused the SNPs on the Amy-d locus to distinguish the AMY1,1 and AMY1,6 alleles in the samples.
First, we constructed two kinds of allele-specific primer sets: MDH500-1 and –7D (AMY1 specific), and MDH500-6 and –7D (AMY6 specific) to amplify only the Amy-d regions encoding AMY1 and AMY6 isozyme, respectively. The sequences of these primers were as follows: MDH500-1: 5'-TACAAGAGAACGTAGTG-3'; MDH500-6: 5'-TACAAGAGAACGTAGCA-3', and –7D: 5'-GCCATATTATGCGTAAC-3'. Using these primer sets, we checked the Amy alleles in a pair of flies after a cross and established AMY1,1 and AMY1,6 homozygous lines from offspring, of which parents were homozygous for the identical allele. Because these parents were isofemale lines sampled from the cage populations, and because our method employed no chromosome from out of the cage populations, unlike many other genetic methods for establishing isogenic lines, all the genomes of the AMY homozygous lines derived from the cage populations. Therefore, measuring the AMY enzyme activities in these lines enabled us to evaluate the effect of selection on their genetic backgrounds, which potentially include unlinked cis and/or trans regulatory factors of Amy. Because we found two Amy-d alleles encoding the AMY1 isozyme in the cage populations, we selected the predominant AMY1 allele by the same method as above, using two additional allele-specific primer sets: MDH100-1 and MDT30 (major-allele specific), and MDH100-1 and MDT30B (minor-allele specific), where MDH100-1 is 5'-CTCACAAATCACTCCCGGCT-3', MDT30 is 5'-TAATTGCTCGGCTTCTC-3', and MDT30B is 5'-TAATTGCTCGGCTTTTC-3'. We confirmed their isozyme patterns by polyacrylamide gel electrophoresis (see Araki et al. 2001 for details). In this way, we could establish the AMY1,1 and AMY1,6 homozygous lines, of which genetic backgrounds were directly derived from the glucose and starch cage populations.
Measurement of AMY Enzyme Activity and Protein Assay
Five 4-day-old female flies from each line were homogenized in 500 µl of distilled water by sonication. The homogenate was centrifuged at 10,000 rpm for 5 min and the supernatant was assayed for enzyme activity and total protein content. Commercially available kits—amylase B-test (Wako Pure Chemical Industries, Osaka, Japan) and BCA protein assay reagent (Pierce, Rockford, IL) were used to measure AMY activity and protein content, respectively [see Inomata et al. (1995a) for details]. AMY activity was expressed in glucose units, where 1 glucose unit represents 10–4 µmol glucose released/min at 37°C. The AMY specific activity was defined as the activity per microgram of total protein content, and a specific activity for a sample was obtained by averaging the AMY activities over the five independent replicates. The three-way analysis of variance (ANOVA), which tests the dependency of the variations in the AMY specific activity on the genetic background, haplotype, and medium on which flies were reared, was performed using StatView version 4.5.
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Results |
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Frequency Changes of AMY1,1
The frequency changes of the AMY1,1 haplotype in the three types of cages (normal, starch, and glucose) are shown in Figure 1. The initial frequencies of the starch and glucose cage populations (generation 0) were 0.460 and 0.465, respectively. After 384 and 312 generations, the AMY1,1 frequencies in the starch and glucose cage populations became 0.170 and 0.893, respectively. The trajectories in Figure 1 show that the haplotype frequencies in the starch and glucose cages were clearly differentiated in opposite directions, while the frequencies in the normal cages showed no particular trend (0.508 in 1999). In the starch cages, only the AMY1,1 and AMY1,6 haplotypes were found after the selection, and the latter was predominant. In spite of the lack of replication (see Materials and Methods), our selection experiment in the starch-rich environment showed the same response as in some of the previous studies (De Jong and Scharloo 1976; Hickey 1977). Although random genetic drift can also explain the frequency changes of AMY in the cage populations, the stability of AMY1,1 frequency in the normal cage population and the agreement of the directional change in AMY1,1 frequency in the substrate-rich environment in the previous studies supported that the selective disadvantage of AMY1,1 had driven the reduction of the AMY1,1 frequency in the starch-rich environment. The selection in the glucose-rich environment, on the other hand, resulted in a high frequency of AMY1,1, suggesting the selective advantage of the naturally predominant AMY isozyme in a product-rich environment.
Nucleotide Sequences of the Amy Regions in Selected Populations
We obtained nucleotide sequences of the duplicated Amy coding regions (1482 bp each) and the 5' flanking regions (656 bp for Amy-p and 609 bp for Amy-d) from 20 isogenic lines from each type of the two cage populations in 1996. The known cis regulatory elements of Amy were located in the flanking regions (Choi and Yamazaki 1994). We also obtained sequences from an unlinked microsatellite locus, DROYANETSB, for comparison. As generally observed in laboratory populations, the genetic variations in the cage populations were reduced (22–57%) in all the food environments and in all the loci investigated compared with those in the Akayu population (Araki et al. 2001), from which the original cage population was established (Table 1). Of interest, however, is that the two distinct cage populations shared the same Amy alleles. All the alleles found in the Amy-p, Amy-d, and DROYANETSB in the starch cage populations were also found in the glucose cage populations. These results suggest that a reduction in effective population size had probably occurred in the original cage population before the selection experiment. Based on the relative ratio of gene diversities of DROYANETSB in the cage populations to those in the natural population (Table 1) and on generation time in the cage populations in 1996 (Figure 1), the effective population size in the cage populations was estimated as 174–303 (see Materials and Methods). Although a greater number of loci are required to better estimate the effective population size, this estimate seems reasonable comparing with the actual population size in the cages (
1000/population), which sometimes varied dramatically.
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Besides sharing of the common alleles of the Amy between populations, the SNPs in the two Amy regions revealed that the Amy loci and their 5' flanking regions were almost completely linked in the cage populations, while there was only one recombinant between the Amy-p and Amy-d loci in a starch cage population (data not shown).
AMY Enzyme Activity of Samples from the Selected Populations
The shared Amy alleles between the cage populations and the almost perfect linkage of the Amy region allowed us to investigate the putative effect of unlinked regulatory factors that may also be selected during the experiments. To investigate the putative contribution and differentiation of the unlinked regulatory factors, we measured the AMY enzyme activities in the AMY homozygous lines for the two predominant haplotypes in the cage populations, which derived from the two cage populations (AMY1,1 + glucose cage [G]-background, AMY1,1 + starch cage [S]-background, and AMY1,6 + G-background and AMY1,6 + S-background). Note that the lines were established from the cage populations directly, so that their genetic backgrounds were identical to those in the cage populations (see Materials and Methods). We measured the AMY enzyme activities of the lines on glucose and starch media, respectively. The effects of the genetic background, AMY haplotype, and medium on the enzyme activities were simultaneously analyzed by a three-way ANOVA (Table 2), and their mean enzyme activities are shown in Figure 2.
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As previously reported, the AMY1,6 lines had higher AMY activity than the AMY1,1 lines (Matsuo and Yamazaki 1986) (corresponding to H in Table 2), and the AMY activity was repressed on the glucose medium compared with that on the starch medium in total (Benkel and Hickey 1986; Inomata et al. 1995a) (M in Table 2). In addition, the genetic variation of the inducibility, which was defined as the ratio of the amount of enzyme production in one environment relative to that in a control environment (Yamazaki and Matsuo 1984), between AMY1,1 and AMY1,6 was detected (H x M). Furthermore, a significant effect of the genetic backgrounds (B) and a significant interaction between the genetic backgrounds and the haplotypes (B x H) were also detected. These results suggest a significant role of unlinked regulatory factors on the genetic background and an epistatic interaction between the AMY haplotype and the genetic background. The characteristic of the epistatic interaction is clearly represented in Figure 2. The AMY1,6 homozygotes with the S-background were approximately two times higher in AMY enzyme activity than those with the G-background, but such a difference in AMY activity was not detected between the AMY1,1 homozygotes with the different cage backgrounds. Based on Table 2 and Figure 2, we conclude that a genetic background that reduces AMY enzyme activity (G-background) was favored in the glucose-rich environment, while the other background that increases AMY activity (S-background) was favored in the starch-rich environment.
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Discussion |
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Selective Significance of the Predominant AMY Isozyme
In this study we determined the selective significance of AMY1,1 in D. melanogaster in a glucose-rich environment. The estimated effective population size of the Akayu population was 2.3 x 104 based on the gene diversity of DROYANETSB (Araki et al. 2001), assuming the selective neutrality, 3.0 x 10–4 per generation, of the mutation rate at this locus (Schlötter et al. 1998) and an equilibrium state with the stepwise mutation model (Ohta and Kimura 1973). The two orders of the reduction in effective population size in the cage populations (174–303, 0.76–1.3%) and the loss of independent replication results in the stronger effect of random genetic drift on the selection experiment. Nevertheless, we observed a clear differentiation in the isozyme frequency (Figure 1) and the solid differences in AMY enzyme activities of flies from the two food environments (the top and bottom lines in Figure 2 for predominant sets of starch and glucose cage populations, respectively).
Based on the changes in AMY1,1 frequency, we can estimate the selective coefficient of the AMY1,1 haplotype in the starch- and glucose-rich environments. Assuming directional selection, the selection coefficient of the AMY1,1 homozygote over the AMY1,6 homozygote, s, can be estimated under the assumptions of random mating, infinite population, s << 1, and the two-alleles model (AMY1,1 and the other haplotypes) by the following equation:
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According to our results, the predominance of the AMY1,1 isozyme in natural populations (Hickey 1979; Singh et al. 1982) indicates that the natural populations of D. melanogaster are generally adapted to glucose-rich environments rather than starch-rich environments. Of interest, however, is that AMY1,1 frequency varies among worldwide populations. For example, the AMY1,1 frequency was only 25% in an African population (Hickey 1979), while it was about 93% in a Japanese population (Yamazaki et al. 1984). The differences in AMY frequency in natural populations may reflect differences in the sources of carbohydrate available to them.
Further investigation is required to address a biological reason why Amy with the lowest enzyme activity is advantageous in product-rich environments rather than selectively neutral. One possible explanation is the cost of protein production when the protein is not required, as suggested in alcohol dehydrogenase of D. melanogaster (Parsch et al. 2000). The facts that AMY activity is repressed in the product-rich environment and that the level of messenger RNA (mRNA) is responsible for AMY activity in D. melanogaster (Yamate and Yamazaki 1999) are consistent with this hypothesis.
Epistatic Interaction and Coadaptation Between Amy and Its Regulatory Factors
Matsuo and Yamazaki (1984) found that flies with higher plasticity for different sources of dietary carbohydrates have higher fitness in the starch-rich environment. Our results are basically consistent with this study, as we also found that flies with higher AMY activity on the starch medium and flies with lower AMY activity on the glucose medium have higher fitness. However, the coadaptation of the haplotype and the genetic background indicates the selective significance of the Amy gene system as a whole, including the epistatic regulatory factors. In the cage populations, the adaptive differentiation of the genetic backgrounds may have occurred in the early stage of the selection experiments, for the following reasons: (1) There was no clear difference in AMY activity between AMY1,1 homozygotes from the two cage populations, and hence selection on the genetic background may not work efficiently in a population where the AMY1,1 frequency is high; and (2) the AMY1,1 frequency was relatively low in both food environments in the early stages of the selection experiments. Once AMY1,1 becomes predominant, like in the glucose cage populations after selection and in natural populations, the selective significance of the genetic background may be masked and the background may behave as if it is selectively neutral. This hypothesis reasonably explains why the genetic backgrounds were polymorphic in the natural population, in which the AMY1,1 was predominant.
The presence of the regulatory factors on the genetic background, which is responsible for the variation in AMY activity, was only suggested in Matsuo and Yamazaki (1997). Further studies on such factors, based on their identification followed by functional and population genetic analyses, will reveal how profoundly such factors are involved in the molecular evolution of the Amy gene system, and may demonstrate the biological and evolutionary importance of the epistatic interactions among loci in higher organisms.
Molecular Evolution of the Amy Gene System in Drosophilids
AMY enzyme activity is high in D. melanogaster among the D. melanogaster species subgroup (Shibata and Yamazaki 1994), and an acceleration of fixed nonsynonymous substitutions was reported in this species after speciation (Araki et al. 2001; Shibata and Yamazaki 1995). Our results suggest that the environmental changes can drive the fixation of an Amy allele, and hence give a possible explanation for the molecular evolution of Amy as a result of an adaptation to a new environment related with speciation. Furthermore, Prigent et al. (2003) showed that large variations of the AMY isozymes exist among drosophilids, and suggested the correlation between isozyme frequencies and the feeding habits of the species. The strong selection on the Amy gene system for adaptation to food environments may be a common issue among the drosophilids or higher organisms.
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Acknowledgments |
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We thank H. Tachida, A. E. Szmidt, J. M. Cameron, M. S. Blouin, and M. Kreitman for useful discussions, and Y. Matsuo, J. I. Choi, T. Ohba, and T. Mitsunaga for technical support. This work was supported, in part, by research grants from the Ministry of Education and Science and Culture of Japan (to T.Y. and N.I.) and by Research Fellowships of the Japan Society for the Promotion of Science for Young Scientists (to H.A.).
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Footnotes |
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Corresponding Editor: Shozo Yokoyama
Received December 1, 2004
Accepted January 3, 2005
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References |
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Araki H, Inomata N, and Yamazaki T, 2001. Molecular evolution of duplicated amylase gene regions in Drosophila melanogaster: evidence of positive selection in the coding regions and selective constraints in the cis-regulatory regions. Genetics 157:667–677.
Benkel BF and Hickey DA, 1986. Glucose repression of amylase gene expression in Drosophila melanogaster. Genetics 114:137–144.
Choi J and Yamazaki T, 1994. Molecular analysis of cis-regulatory sequences of the alpha-amylase gene in D. melanogaster: a short 5'-flanking region of Amy distal gene is required for full expression of Amy proximal gene. Jpn J Genet 69:619–635.[CrossRef][Medline]
Cooper VS and Lenski RE, 2000. The population genetics of ecological specialization in evolving Escherichia coli populations. Nature 407:736–739.[CrossRef][Medline]
Crow JF and Kimura M, 1970. An introduction to population genetics theory New York: Harper & Row.
De Jong GA and Scharloo W, 1976. Environmental determination of selective significance or neutrality of amylase variants in Drosophila melanogaster. Genetics 84:77–94.
Doane WW, 1969. Amylase variants in Drosophila melanogaster: linkage studies and characterization of enzyme extracts. J Exp Zool 171:321–342.[CrossRef]
Doane WW, 1980. Selection for amylase allozymes in Drosophila melanogaster: some questions. Evolution 34:868–874.[CrossRef]
Gupta AP and Lewontin RC, 1982. A study of reaction norms in natural populations of Drosophila pseudoobscura. Evolution 36:934–948.[CrossRef]
Hickey DA, 1977. Selection for amylase allozymes in Drosophila melanogaster. Evolution 31:800–804.[CrossRef]
Hickey DA, 1979. The geographical pattern of an enzyme polymorphism in D. melanogaster. Genetica 51:1–4.
Inomata N, Kanda K, Cariou ML, Tachida H, and Yamazaki T, 1995a. Evolution of the response patterns to dietary carbohydrates and the developmental differentiation of gene expression of alpha-amylase in Drosophila. J Mol Evol 41:1076–1084.[Web of Science][Medline]
Inomata N, Shibata H, Okuyama E, and Yamazaki T, 1995b. Evolutionary relationships and sequence variation of alpha-amylase variants encoded by duplicated genes in the Amy locus of Drosophila melanogaster. Genetics 141:237–244.[Abstract]
Inomata N and Yamazaki T, 2000. Evolution of nucleotide substitutions and gene regulation in the amylase multigenes in Drosophila kikkawai and its sibling species. Mol Biol Evol 17:601–615.
Kang MS and Gauch HG, 1996. Genotype-by-environment interaction Boca Raton, FL: CRC Press.
Matsuo Y and Yamazaki T, 1984. Genetic analysis of natural populations of Drosophila melanogaster in Japan IV. Natural selection on the inducibility, but not on the structural genes, of amylase loci. Genetics 108:879–896.
Matsuo Y and Yamazaki T, 1986. Genetic analysis of natural populations of Drosophila melanogaster in Japan VI. Differential regulation of duplicated amylase loci and degree of dominance of amylase activity in different environments. Jpn J Genet 61:543–558.
Matsuo Y and Yamazaki T, 1997. Genetic factors on the second and third chromosomes responsible for the variation of amylase activity and inducibility in Drosophila melanogaster. Genet Res 70:97–103.[CrossRef][Web of Science][Medline]
Nei M, 1987. Molecular evolutionary genetics New York: Columbia University Press.
Ohta T and Kimura M, 1973. A model of mutation appropriate to estimate the number of electrophoretically detectable alleles in a finite population. Genet Res 22:201–204.[Web of Science][Medline]
Okuyama E, Shibata H, Tachida H, and Yamazaki T, 1996. Molecular evolution of the 5'-flanking regions of the duplicated Amy genes in Drosophila melanogaster species subgroup. Mol Biol Evol 13:574–583.[Abstract]
Parsch J, Russell JA, Beerman I, Hartl DL, and Stephan W, 2000. Deletion of a conserved regulatory element in the Drosophila Adh gene leads to increased alcohol dehydrogenase activity but also delays development. Genetics 156:219–227.
Powell JR and Andjelkovic M, 1983. Population genetics of Drosophila amylase. IV. Selection in laboratory populations maintained on different carbohydrates. Genetics 103:675–689.
Prigent S, Renard E, and Cariou ML, 2003. Electrophoretic mobility of amylase in drosophilids indicates adaptation to ecological diversity. Genetica 119:133–145.[CrossRef][Web of Science][Medline]
Remold SK and Lenski RE, 2001. Contribution of individual random mutations to genotype-by-environment interactions in Escherichia coli. Proc Natl Acad Sci USA 98:11388–11393.
Remold SK and Lenski RE, 2004. Pervasive joint influence of epistasis and plasticity on mutational effects in Escherichia coli. Nat Genet 36:423–426.[CrossRef][Web of Science][Medline]
Rozas J and Rozas R, 1999. DnaSP version 3: an integrated program for molecular population genetics and molecular evolution analysis. Bioinformatics 15:174–175.
Schlötter C, Ritter R, and Bram G, 1998. High mutation rate of a long microsatellite allele in Drosophila melanogaster provides evidence for allele-specific mutation rates. Mol Biol Evol 15:1269–1274.[Abstract]
Shibata H and Yamazaki T, 1994. A comparative study of the enzymological features of alpha-amylase in the Drosophila melanogaster species subgroup. Jpn J Genet 69:251–258.[CrossRef][Medline]
Shibata H and Yamazaki T, 1995. Molecular evolution of the duplicated Amy locus in the Drosophila melanogaster species subgroup: concerted evolution only in the coding region and an excess of nonsynonymous substitutions in speciation. Genetics 141:223–236.[Abstract]
Singh RS, Hickey DA, and David J, 1982. Genetic differentiation between geographically distant populations of Drosophila melanogaster. Genetics 101:235–256.
Thompson JD, Gibson TJ, Plewniak F, Jeanmougin F, and Higgins DG, 1997. The CLUSTALX Windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res 25:4876–82.
Yamate N and Yamazaki T, 1999. Is the difference in alpha-amylase activity in the strains of Drosophila melanogaster with different allozymes due to transcriptional or posttranscriptional control?. Biochem Genet 37:345–356.[CrossRef][Web of Science][Medline]
Yamazaki T, Matsuo Y, Inoue Y, and Matsuo Y, 1984. Genetic analysis of natural populations of Drosophila melanogaster in Japan. I. Protein polymorphism, lethal gene, sterility gene, inversion polymorphism and linkage disequilibrium. Jpn J Genet 59:33–49.
Yamazaki T and Matsuo Y, 1984. Genetic analysis of natural populations of Drosophila melanogaster in Japan. III. Genetic viability of amylase inducibility and fitness. Genetics 108:223–235.
Zhang Z, Inomata N, Cariou ML, Da Lage JL, and Yamazaki T, 2003. Phylogeny and the evolution of the amylase multigenes in the Drosophila montium species subgroup. J Mol Evol 56:121–130.[CrossRef][Web of Science][Medline]
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