Skip to main content
PMC home page
Genetics logoLink to Genetics
. 2008 Jun;179(2):1021–1027. doi: 10.1534/genetics.107.078345

Recurrent Deletion and Gene Presence/Absence Polymorphism: Telomere Dynamics Dominate Evolution at the Tip of 3L in Drosophila melanogaster and D. simulans

Andrew D Kern *,1, David J Begun †
PMCID: PMC2429855  PMID: 18505885

Abstract

Although Drosophila melanogaster has been the subject of intensive analysis of polymorphism and divergence, little is known about the distribution of variation at the most distal regions of chromosomes arms. Here we report a survey of genetic variation on the tip of 3L in D. melanogaster and D. simulans. Levels of single nucleotide polymorphism in the most distal euchromatic sequence are approximately one order of magnitude less than that typically observed in genomic regions of normal crossing over, consistent with what might be expected under models of linked selection in regions of low crossing over. However, despite this reduced level of nucleotide variation, we found abundant deletion polymorphism. These deletions create at least three gene presence/absence polymorphisms within D. melanogaster: the putative G-protein coupled receptor mthl-8 (which is the most distal known or predicted gene on 3L) and the unannotated mRNAs AY060886 and BT006009. Strikingly, D. simulans is also segregating deletions that cause mthl8 presence/absence polymorphism. Breakpoint sequencing and tests of correlations with segregating SNPs in D. melanogaster suggest that each deletion is unique. Cloned breakpoint sequences revealed the presence of Het-A elements just distal to unique, canonical euchromatic sequences. This pattern suggests a model in which repeated telomeric deficiencies cause deletions of euchromatic sequence followed by subsequent “healing” by retrotranposition of Het-A elements. These data reveal the dominance of telomeric dynamics on the evolution of closely linked sequences in Drosophila.

THE recognition that changes in gene number might be important material for evolution came very early in the history of genetics (Haldane 1932; Bridges 1935; Muller 1935). Ohno (1967) went so far as to suggest that evolution by gene duplication was the most important factor in genomic evolution. Indeed, whole genome analyses have demonstrated conclusively that gene number often differs greatly between species (e.g., Adams et al. 2000; Holt et al. 2002). Although gene copy-number variation is typically thought of as having a lower bound of one, surprisingly, some single-copy genes (or gene families) that are present in most species in a clade, nevertheless, appear to be absent from others, raising questions about the functional consequences and population genetic mechanisms responsible for such patterns (Hahn et al. 2007). Such interspecific differences must have their origin as intraspecific variation (Begun and Lindfors 2005; Greenberg et al. 2006; Wang et al. 2006; Begun et al. 2007a,b; Dopman and Hartl 2007).

Duplications and deletions are thought to be more common in highly repetitive parts of the genome (e.g., Achaz et al. 2001), raising the possibility that patterns of gene presence/absence variation in such regions may be related to their population genetic and genetic properties. However, the structural, functional, and evolutionary characterization of the repetitive sequences of eukaryotic heterochromatin, which populate the centromeric as well telomeric regions of chromosomes, have remained elusive. In spite of these difficulties, variation both within and between species in telomeric or subtelomeric regions appears to be the rule rather than the exception in plants (Broun et al. 1992), worms (Caenorhabditiselegans Sequencing Consortium 1998), yeast (Winzeler et al. 2003), and humans (reviewed in Vergnaud 1999). Most telomeric variation seems to take the form of duplications and deletions (e.g., Winzeler et al. 2003) rather than the point mutations that are more common in other chromosomal locations. Thus the forces that create telomere variation may be different than forces acting on other parts of the genome.

Patterns of population genetic variation at the ends of Drosophila chromosomes are largely uncharacterized. The distal region of the Drosophila melanogaster X chromosome shows reduced levels of heterozygosity but relatively low levels of linkage disequilibrium (LD) (Begun and Aquadro 1995; Langley et al. 2000). The analysis of the tip of the X chromosome has recently been extended to the subtelomeric sequences, which show rapid divergence, including high levels of insertions and deletions, low heterozygosity, and evidence of extensive recombination (Anderson et al. 2008). Here we report on an unusual case of gene presence/absence variation in D. melanogaster and D. simulans. These gene polymorphisms are located in the most distal region of chromosome arm 3L and are associated with large deletions of subtelomeric sequence.

MATERIALS AND METHODS

Fly stocks/sequencing:

North American D. melanogaster isofemale lines used for our initial detection of mthl8 deletion polymorphism came from lines collected in California by the authors. African-extracted chromosome lines were derived from isofemale lines collected in Zimbabwe. A set of 50 North American D. melanogaster third chromosome extraction lines from Pennsylvania were kindly provided to us by Andy Clark; these lines were isogenic for chromosomes X and II. These lines were used for all sequencing, Southern blotting, and qRT–PCR experiments reported in the article. A sample of 30 inbred D. simulans lines derived from females collected in Winters, California (S. Nuzdhin) were used in Southern blot and PCR experiments. All polymorphism data sequencing was performed via sequencing of PCR products in both directions on an ABI 3700 sequencer. Raw data were analyzed using phred (Ewing et al. 1998), and phrap (Ewing and Green 1998). Sequences were then examined by eye using consed (Gordon et al. 1998). Polymorphic sites in multiple alignments were verified using the MACE package of scripts (W. Gilliland, unpublished results). All primer sequences and locations are given in supplemental Table 1. All coordinates are relative to GenBank accession no. AE003467. The primers listed in supplemental Table 1 include those used for surveys of sequence polymorphism, mapping of deletion breakpoints, as well as subsequent RT–PCR experiments. New sequences reported in this article can be found in GenBank under accession nos. EU394734–EU394868. Alignments are available from A.D.K. upon request.

Sequence analysis:

All alignments were done using either CLUSTALW 1.82 (Thompson et al. 1994) and/or DIALIGN 2.2 (Morgenstern 1999) and were subsequently edited by hand. In all cases alignment was simple and without major ambiguity. All polymorphism and divergence statistics were estimated using software developed by A.D.K. Source code in Ruby is available upon request. Homology-based searching was performed using BLAT and the genome assemblies of D. simulans (droSim1) and D. yakuba (droYak1) are available from the University of California Santa Cruz (UCSC) genome browser. Most statistical analysis was performed using R, version 2.1 (http://cran.r-project.org/). Composite likelihood estimation of the population recombination rate was performed using Ldhat, version 2.0 (McVean et al. 2002).

Southern analysis:

DNA preparations were performed using a scaled down version of the protocol of E. J. Rehm (BDGP: http://www.fruitfly.org/about/methods/inverse.pcr.html). Individual flies were ground with a micropestle in a 1.5-ml microfuge tube with 40 μl of solution A (100 mm Tris–HCl, pH 7.5, 20 mm EDTA, pH 8.0, 100 mm NaCl, 0.5% SDS), incubated at 65° for 30 min and then mixed with 160 μl of solution B (1.43 m KC2H3O2, 4.29 m LiCl). Preparations were incubated for 10 min on ice and then spun for 15 min at maximum speed in a microcentrifuge. DNA was precipitated from the supernatant with 120 μl of isopropanol, washed with 70% ethanol, dried briefly (∼5–10 min) at room temperature, and finally resuspended in 10 μl ddH2O.

Genomic DNA was digested to completion with HinfI, separated on 1.0% agarose/TBE gels, and transferred onto Nytran SuPerCharge (Schleicher & Schuell, Keene, NH) membranes via the alkaline method. The mthl8 probe (∼900 bp) was PCR amplified from D. melanogaster or D. simulans depending on the origin of the target DNA, using the CG32475F7 and CG32475R7 primers (62° annealing temperature) and radiolabeled with [32P]dCTP by random priming. Membranes were hybridized overnight at 60° and then washed twice at 55°.

qRT–PCR experiment:

The purpose of our qRT–PCR experiment was to determine whether subtelomeric deletions affect transcription of nearby regions of the chromosome (cf. Beissmann et al. 2005). A subset of Pennsylvania third chromosome extraction lines (n = 16), that represented both deleted and nondeleted classes, was reared at 25°. White prepupae were collected, flash frozen in liquid nitrogen, and stored at −80° until RNA extraction. Total RNA was extracted from 10 prepupae using a Trizol prep (Gibco, Calsbad, CA). Two independent RNA preps were made for each line. RNAs were purified and DNased using the RNAeasy kit and the RNase-free DNase set available from QIAGEN (Valencia, CA). cDNA was synthesized using first-strand Taqman RT reagents (Applied Biosystems, Foster City, CA). SYBR green PCR mix (ABI) was used for detection of amplification products on an ABI PRISM 7700. Two genes were assayed, Lsp-1-γ (the closest gene to mthl8) and Gapdh-1 (chromosome 2), which served as the control. Primers were designed using ABI's Primer Express software and are available upon request. Gapdh-1 primers were PCR tested for cross-amplification with its paralog; none was detected. For each combination of sample and primers, three replicate reactions were carried out (i.e., technical replicates). Additionally, two minus-RT controls, and two no-template controls were run for each sample. Melting curve analysis was performed at the end of each PCR run to verify product specificity. Statistical analysis was performed on Ct measurements of Lsp1γ that had been normalized against the housekeeping gene Gapdh-1 (i.e., the Δ-Δ-Ct method; Livak and Schmittgen 2001).

RESULTS

Presence/absence polymorphism of mthl8 in D. melanogaster:

The subtelomeric gene mthl8 was initially identified as a putative candidate for presence/absence polymorphism in D. melanogaster populations serendipitously, through hybridization of population samples of D. melanogaster genomic DNA to cDNA arrays (C. Meiklejohn, personal communication). Our PCR analysis of the mthl8 coding region from a California sample of D. melanogaster extracted third chromosomes strengthened the idea that the gene is missing from some chromosomes, as 5 of 21 third chromosome lines repeatedly failed to amplify. Given that PCR failure could result from variants in primer annealing sites as well as from gene presence/absence variation, we used Southern analysis to conclusively determine whether chromosome lines that failed to PCR amplify nevertheless contain a copy of the gene. These genomic Southern blot experiments confirmed that all PCR failures result from deletion of mthl8 coding sequence (data not shown). We then used PCR and Southern analysis on a second, larger sample of extracted third chromosome lines from the Pennsylvania sample, which showed a slightly higher frequency of deleted chromosomes (18/50) relative to the California sample. An analysis of 20 African third chromosome extraction lines revealed that mthl8 presence/absence variation is not restricted to cosmopolitan populations, as 5 of 20 lines showed no evidence of mthl-8 coding sequence.

Presence/absence variation of mthl8 in other Drosophila:

Given the mthl8 presence/absence polymorphism in D. melanogaster, we were interested in establishing the phylogenetic distribution of the locus. We screened 30 inbred lines of D. simulans, again using PCR and Southern blots, and found 8 of the 30 lines to be homozygous for deletion of mthl8. One of the lines screened for mthl8 deletion, sim6, was part of the recent effort to carry out light shotgun sequencing on a number of D. simulans genomes (http://www.dpgp.org, Begun et al. 2007b). As expected from PCR and Southern results, in silico homology-based searches showed no evidence of the mthl8 coding region in the sequenced genome. However, unique flanking sequence with homology to D. melanogaster could be identified near the tip of 3L. These data demonstrate that D. simulans is also polymorphic for the presence of mthl8.

A BLAT search of the mthl8 protein against the April 2004 assembly of the D. yakuba genome (http://genome.ucsc.edu) provided no good evidence of an ortholog on 3L, but did point to a genomic region (chr 3L: 338,820-342,024) containing a clear ortholog of mthl10, a diverged paralog of mthl8. In D. pseudoobscura however there is an annotated ortholog, GA16932 (November 2004 assembly). Thus the gene was likely present in the last common ancestor of D. melanogaster and D. pseudoobscura, which suggests it has an important function in at least some lineages.

Analysis of mthl8 deletion breakpoints in D. melanogaster:

We used a collection of amplification primers spaced throughout the tip of 3L to coarsely map deletion breakpoints (Figure 1 and supplemental Table 1). Amplification of some products and not others for individual deleted chromosomes provided map information with respect to the presence of “canonical” sequence as inferred from the D. melanogaster reference sequence. Note that the dense spacing of PCR primer pairs typically resulted in a contiguous set of failed primer pairs, which strongly supports the deletion hypothesis. A striking diversity of breakpoints was observed, some of which suggest the presence of deletions of >40 kb of euchromatic sequence (Figure 1).

Figure 1.—

Figure 1.—

Open in a new tab

Deletion polymorphism at the tip of 3L. Many chromosomes extracted from nature do not carry the putative G-protein coupled receptor mthl-8. The diversity of 3L subtelomeric sequences that we have found in North American chromosomes are shown. Browser tracks from top to bottom are as follows: (1) The positions of PCR markers used to map deletion breakpoints (black), (2) PCR markers also used to collect nucleotide polymorphism data (red; refer to Table 1 for locus names), (3) sequence determined to be present in numerous mthl8 deletion lines (black; line names in left-hand column), (4) protein-coding genes (blue), (5) D. melanogaster mRNAs (black), and (6) 12-way multiz alignment conservation scores (gray).

In an effort to sequence across deletion breakpoints, we designed forward PCR primers against known telomeric sequences (HeT-A and TART) and reverse primers from the most distal, unique euchromatic region of 3L that, on the basis of our PCR mapping, would amplify for a given strain. PCR reaction conditions for these experiments were optimized for extremely long products. Successfully amplified products were sequenced directly. In all, we were able to sequence across 7 of the 18 deletions in our sample. Figure 2 presents a schematic of the structure of deletion breakpoints from three chromosomes. These three chromosomes are representative of the two broad classes of deletions we observed, complete deletions of mthl8 and partial deletions of mthl8. Partial deletions were originally identified via Southern blotting because our probe sequence did not cover the 3′ portion of the coding sequence. Some of the larger deletions also remove one or two unannotated mRNAs (BT006009 and AY060886), both of which have been recovered as full-length cDNA inserts. Thus the largest deletions may be removing three single-copy genes from the genome.

Figure 2.—

Figure 2.—

Open in a new tab

Schematic of deletion breakpoint structure. Each bar represents one chromosome extracted from a natural population. The red hatched box represents HeT-A sequences, blue boxes represent mthl-8 coding sequence, and gray is euchromatic noncoding sequence. Breakpoint identities are labeled and correspond to Figure 1 labeling. See Figure 1 for relative positions.

Interestingly, in each deleted chromosome, euchromatic sequences directly abut HeT-A retrotransposons. HeT-A elements are known to be telomere specific in their genomic distribution (Rubin 1978; Karpen and Spradling 1992; Levis et al. 1993) and have been shown to “heal” broken chromosome ends through transposition (Beissmann et al. 1990, 1992). Thus, these segregating deletions may represent independent terminal deletions that were healed by HeT-A transposition. Furthermore, these data suggest that D. melanogaster deletions are likely to be independent from D. simulans terminal deletions rather than ancient variants that were segregating in the common ancestor.

Population genetic variation at the tip of 3L:

The extensive deletion polymorphism in the subtelomeric region of 3L was surprising given the fact that cytological section 61A is thought to be a region of reduced crossing over in D. melanogaster (Ashburner 1989). To further investigate the distribution of variation in the region, we obtained polymorphism data from Pennsylvania chromosomes for four loci in the mthl8 region (see Figure 1). Table 1 shows the results from this survey. Average heterozygosity at the tip of 3L (mean θπ = 0.0012, mean θW = 0.0015) is about an order of magnitude lower than that observed in regions of normal recombination in D. melanogaster populations (Andolfatto 2001). This pattern is consistent with the reduced levels of variation that have been observed earlier in areas of low crossing over in D. melanogaster populations (e.g., Aguade et al. 1989; Begun and Aquadro 1991, 1995; Berry et al. 1991; Langley et al. 1993, 2000). A general picture of divergence at the tip of 3L can be outlined using a comparison of the current D. melanogaster and D. simulans assembly (dm3 and droSim1) alignments that have been done as part of a 15-way multiz alignment available at UCSC (http://genome.ucsc.edu). Using the mafNets that underlie this track, we find that in the 50-kb region surrounding mthl8 (chr 3L: 2903-52,902), ∼89% of bases are aligned after accounting for repeat masking. Using the droSim1 assembly from UCSC, D. simulans orthologs of mthl8 and Lsp1γ regions used in our polymorphism resequencing survey could be found, but no obvious D. simulans orthologs of the intergenic f25 and f72 loci were detected, which suggests an unusually high rate of indel divergence in this genomic region, similar to what was observed in the subtelomeric sequence of the X chromosome (Anderson et al. 2008). The conservation track in Figure 1 demonstrates this visually. Jukes-Cantor corrected divergence per nucleotide site is ∼0.21. This number is at the high end of sequence divergence between D. melanogaster and D. simulans, and should be treated cautiously, as sequence assembly at the tips of Drosophila chromosomes is difficult. Nevertheless, the high divergence suggests that reduced nucleotide polymorphism in D. melanogaster is not explained by reduced neutral mutation rates. Values of Tajima's D statistic (Tajima 1989) show no evidence for a strong skew toward rare alleles, as might be expected under a hitchhiking model (e.g., Braverman et al. 1995).

TABLE 1.

Summary statistics of DNA polymorphism from mthl8 region of D. melanogaster

Population Locus N bp S H θW θπ Tajima's D Lower 95% D Upper 95% D
Pooled Lsp1γ 46 472 3 4 0.0014 0.0023 1.2360 −1.4537 1.7102
f72 25 842 4 5 0.0013 0.0006 −1.5288 −1.5288 1.6172
f25 41 722 3 3 0.0013 0.0004 −1.4354 −1.5711 1.6769
mthl8 23 685 5 6 0.0020 0.0016 −0.6397 −1.5299 1.6199
Average 0.0015 0.0012
Deleted Lsp1γ 18 472 3 4 0.0018 0.0025 0.9257 −1.7130 1.7152
f72 7 842 1 2 0.0005 0.0004 NA NA NA
f25 15 722 2 2 0.0009 0.0008 NA NA NA
Average 0.0011 0.0012
Undeleted Lsp1γ 28 472 2 3 0.0011 0.0020 NA NA NA
f72 18 842 3 4 0.0011 0.0007 −1.1313 −1.7130 1.6113
f25 26 722 3 3 0.0014 0.0007 −1.3110 −1.5127 1.6375
mthl8 23 685 5 6 0.0020 0.0016 −0.6397 −1.5299 1.6199
Average 0.0012 0.0011
Open in a new tab

Population refers to chromososme classes defined by presence/absence of mthl8. Locus is the locus surveyed. N is the number of alleles sampled. bp is the average length of the sequence within the sample. S is the number of segregating sites in the sample. H is the number of haplotypes observed. θπ and θW are estimates of nucleotide polymorphism. Tajima's D is a measure of the site frequency spectrum; the lower and upper 95% confidence intervals were determined by 105 coalescent simulations without recombination and conditioned on the observed number of segregating sites.

Remarkably, these data suggest that the high level of deletion variation is physically distal to a region of severely reduced nucleotide heterozygosity. Given this, it is of interest to ask whether chromosomes belonging to the “deleted” class are in any way distinct from nondeleted chromosomes in terms of the distribution of nucleotide variation. Table 1 shows polymorphism summary statistics from the surveyed loci for deleted and nondeleted chromosomes. Levels of polymorphism are not significantly different between the classes of chromosomes (Wilcoxon rank-sum tests, θπ P-value = 0.6531; θw P-value = 0.6579). Thus, deleted chromosomes do not appear to constitute a distinct subset of the population sample.

Patterns of linkage disequilibrium between the deletions and segregating sites in the region are presented in Figure 3, which displays a diagonal matrix of significance levels of nonrandom association between polymorphic sites. Little linkage disequilibrium is seen in the region, both between deletions and SNPs, as well as among SNPs. All Fisher's exact tests of association between deletions and SNPs had a nonsignificant P-value. Consistent with the pairwise analysis, composite likelihood estimates of the population recombination rate (4Nr; Hudson 1983; McVean et al. 2002) for the data shown in Figure 3 were ≥100, indicating a large amount of recombination in the history of the sample.

Figure 3.—

Figure 3.—

Open in a new tab

Polymorphism and linkage disequilibrium in the mthl8 region. Regions surveyed for polymorphism are shown on the second track in black and labeled according to the naming conventions presented in Table 1. Gene exon structure is shown in blue, where both mthl8 and Lsp1-γ can be seen. Lines extending down to the triangular matrix represent the positions of SNPs used in the analysis of LD. The matrix of squares represents the statistical significance determined by Fisher's exact test (uncorrected for multiple tests); white, P > 0.1; yellow, 0.05 < P < 0.1; red, P < 0.05.

Transcriptional analysis of subtelomeric deletions:

To test whether deletions affect transcript levels of the neighboring gene, Lsp1γ, we performed qRT–PCR on white prepupae from 16 third chromosome extraction lines of D. melanogaster (8 deleted; 8 undeleted). While a significant line effect was found in our experiment (ANOVA, P < 0.00001), no effect of deletion was observed (ANOVA, P = 0.128). This indicates that although deletions move Lsp1γ closer to heterochromatic sequences near the telomere, there is no large effect on Lsp1γ transcript abundance.

DISCUSSION

Large amounts of duplication and deletion polymorphism within and between species are characteristic of the telomeric and subtelomeric regions of plants, worms, yeast, and humans (Broun et al. 1992; Caenorhabditiselegans Sequencing Consortium 1998; Vergnaud 1999; Winzeler et al. 2003). The data reported here and in Anderson et al. (2008) show that Drosophila, too, can be added to the list of organisms harboring extensive variation near the tips of chromosomes. The unique repetitive structure of telomeres, which plays a role in replicating the ends of linear chromosomes, is likely to be responsible for this extensive brand of genetic variation. Remarkably, levels of nucleotide heterozygosity in the euchromatic sequence adjacent to the subtelomeric deletions were severely reduced compared to levels observed in regions of normal crossing over. A similar pattern was recently reported for the X chromosome (Anderson et al. 2008). The fact that nucleotide divergence between species is typical (or slightly elevated) relative to that observed throughout most of the genome supports the idea that reduced nucleotide heterozygosity is a result of linked selection. Given the low rates of crossing over at the tips of Drosophila autosomes (Ashburner 1989), one might expect relatively high levels of linkage disequilbrium in the surveyed region. However, we observed little correlation between segregating sites, or between segregating sites and indel polymorphisms, which is indicative of extensive recombination in our study region. This pattern is reminiscent of observations from the tip of the X chromosome (Begun and Aquadro 1995; Langley et al. 2000; Anderson et al. 2008), where low levels of linkage disequilibrium in an area with demonstrably low levels of crossing over have been found. A possible explanation for this pattern is that genomic regions experiencing low crossing over experience relatively normal levels of gene conversion. The maintenance of duplication and deletion variation in subtelomeric regions could then be interpreted as a result of both a high mutation rate to new variants and a relatively high rate of exchange in these regions.

The observation of genes that have been maintained over evolutionary time, but are polymorphic within species, is interesting. This phenomenon has previously been observed in Drosophila, primarily at reproduction-related genes (Begun and Lindfors 2005; Greenberg et al. 2006; Wang et al. 2006; Begun et al. 2007a,b). In those cases, changes in selection pressures across lineages may leave certain genes susceptible to loss either by drift or natural selection. Here, given the unique chromosomal context of mthl8, it seems likely that a high mutation rate to deletion variants at the tip of chromosome arm 3L is a main contributor to the maintenance of the gene presence/absence variation. The fact that the same phenomenon is observed in D. melanogaster and the sister species, D. simulans, certainly supports the view that the genetic properties of the tip of 3L are important. It is interesting to note that crude PCR surveys of the most distal euchromatic gene sequence of chromosome arm 3R (map205) provided no evidence of common deletions (data not shown), suggesting that genetic and/or population genetic processes acting at tips of different Drosophila chromosome arms may be heterogeneous.

The selective forces acting on the deletion variants remain unclear and may very well be heterogeneous, depending on the extent of deleted sequence. Deletion variants may be maintained by mutation–selection balance, in which case the high frequency of deletions may be attributable to both a high mutation rate and weak selection. For example, the frequency of natural l(2)gl variants is surprisingly high (Golubovsky and Sokolova 1973, Golubovsky 1978, Green and Shepherd 1979), although not nearly as high as the observed mthl8 deletion variants. Interestingly, many natural l(2)gl alleles are the result of terminal deletions of chromosome arm 2L (Mechler et al. 1985), which suggests the possibility that the tip of chromosome 2L also experiences a high mutation rate to deletions (Walter et al. 1995). Alternatively, deletions may be favored by natural selection, at least in some environments. Sokolova and Golubovsky (1979) found that l(2)gl heterozygotes had a viability advantage over wild type at both high and low temperatures. Indeed preliminary data suggest that the mthl8 deletion shows clinal variation in D. melanogaster (Turner et al. 2008), which would be consistent with spatially varying selection acting among D. melanogaster populations. Finally, telomeric variants could plausibly be associated with meiotic phenotypes affecting their fitness.

Telomeric dynamics may create a situation in which the spatial distribution of genes of large fitness effect (e.g., “essential” genes) are excluded from some subtelomeres over evolutionary time, as a high rate of deletion of such genes should result in a significant fitness reduction. Thus, rearrangements that move an essential gene away from a subtelomere that experiences high deletion rates may be favored by natural selection.

Acknowledgments

We thank John Gillespie, Chuck Langley, Matt Hahn, Corbin Jones, and the University of California Davis Evolutionary Discussion Group for valuable comments and discussion. M. Kerber and V. Chen provided technical assistance. A.D.K. was funded by a Howard Hughes Medical Institute predoctoral fellowship and a fellowship from the National Institutes of Health (NIH)/National Human Genome Research Institute. This work was supported by the National Science Foundation DEB-0327049 and NIH GM-071926.

References

  1. Achaz, G., P. Netter and E. Coissac, 2001. Study of intrachromosomal duplications among the eukaryote genomes. Mol. Biol. Evol. 18 2280–2288. [DOI] [PubMed] [Google Scholar]
  2. Adams, M. D., S. E. Celniker, R. A. Holt, C. A. Evans et al., 2002. The genome sequence of Drosophila melanogaster. Science 287 2185–2195. [DOI] [PubMed] [Google Scholar]
  3. Aguade, M., N. Miyashita and C. H. Langley, 1989. Reduced variation in the yellow-achaete-scute region in natural population of Drosophila melanogaster. Genetics 122 607–615. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Anderson, J. A., Y. S. Song and C. H. Langley, 2008. Molecular population genetics of Drosophila subtelomeric DNA. Genetics 178 477–487. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Andolfatto, P., 2001. Adaptive hitchhiking effects on genome variability. Curr. Opin. Genet. Dev. 11 635–641. [DOI] [PubMed] [Google Scholar]
  6. Ashburner, M., 1989. Drosophila: A Laboratory Handbook. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
  7. Begun, D. J., and C. F. Aquadro, 1991. Molecular population genetics of the distal portion of the X chromosome in Drosophila: evidence for genetic hitchhiking of the yellow-achaete region. Genetics 129 1147–1158. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Begun, D. J., and C. F. Aquadro, 1995. Evolution at the tip and base of the X chromosome in an African population of Drosophila melanogaster. Mol. Biol. Evol. 12 382–390. [DOI] [PubMed] [Google Scholar]
  9. Begun, D. J., and H. A. Lindfors, 2005. Rapid evolution of genomic Acp complement in the melanogaster subgroup of Drosophila. Mol. Biol. Evol. 22 2010–2021. [DOI] [PubMed] [Google Scholar]
  10. Begun, D. J., H. A. Lindfors, A. D. Kern and C. D. Jones, 2007. a Evidence for de novo evolution of testis-expressed genes in the Drosophila yakuba/Drosophila erecta clade. Genetics 176 1131–1137. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Begun, D. J., A. K. Holloway, K. Stevens, L. W. Hillier, Y. P. Poh et al., 2007. b Population genomics: whole-genome analysis of polymorphism and divergence in Drosophila simulans. PLoS Biol. 11 e310. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Beissmann, H., J. M. Mason, K. Ferry, M. d'Hulst, K. Valgeirsdottir et al., 1990. Addition of telomere-associated HeT DNA sequences “heals” broken chromosome ends in Drosophila. Cell 61 663–673. [DOI] [PubMed] [Google Scholar]
  13. Beissmann, H., K. Valgeirsdottir, A. Lofsky, C. Chin, B. Ginther et al., 1992. HeT-A, a transposable element specifically involved in “healing” broken chromosome ends in Drosophila melanogaster. Mol. Cell. Biol. 12 3910–3918. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Beissmann, H., S. Parad, V. F. Semeshin, E. N. Andreyeva, Q. Nguyen et al., 2005. Two distinct domains in Drosophila melanogaster telomeres. Genetics 171 1767–1777. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Berry, A. J., J. W. Ajioka and M. Kreitman, 1991. Lack of polymorphism on the Drosophila fourth chromosome resulting from selection. Genetics 129 1111–1117. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Braverman, J. M., R. R. Hudson, N. L. Kaplan, C. H. Langley and W. Stephan, 1995. The hitchhiking effect on the site frequency spectrum of DNA polymorphisms. Genetics 140 783–796. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Broun, P., M. W. Ganal and S. D. Tanksley, 1992. Telomeric array display high levels of heritable polymorphism among closely related plant varieties. Proc. Natl. Acad. Sci. USA 89 1354–1357. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Bridges, C. B., 1935. Salivary chromosome maps. J. Hered. 26 60–64. [Google Scholar]
  19. Caenorhabditis elegans Sequencing Consortium, 1998. Genome sequence of the nematode C. elegans: a platform for investigating biology. Science 11 2012–2018. [DOI] [PubMed] [Google Scholar]
  20. Dopman, E. B., and D. L. Hartl, 2007. A portrait of copy number polymorphism in Drosophila melanogaster. Proc. Natl. Acad. Sci. USA 104 19920–19925. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Ewing, B., and P. Green, 1998. Base-calling of automated sequencer traces using phred. II. Error probabilities. Genome Res. 8 186–194. [PubMed] [Google Scholar]
  22. Ewing, B, L. Hillier, M. C. Wendl and P. Green, 1998. Base-calling of automated sequencer traces using phred. I. Accuracy assessment. Genome Res. 8 175–185. [DOI] [PubMed] [Google Scholar]
  23. Golubovsky, M. D., 1978. The “lethal giant larvae”—the most frequent second chromosome lethal in natural populations of D. melanogaster. Dros. Inf. Serv. 53 179. [Google Scholar]
  24. Golubovsky, M. D., and K. B. Sokolova, 1973. The expression and interaction of different alleles at the l(2)gl locus. Dros. Inf. Serv. 50 124. [Google Scholar]
  25. Gordon, D., C. Abajian and P. Green, 1998. Consed: a graphical tool for sequence finishing. Genome Res. 8 195–202. [DOI] [PubMed] [Google Scholar]
  26. Green, M. M., and S. H. Shepherd, 1979. Genetic instability in Drosophila melanogaster: the induction of specific chromosome 2 deletions by MR elements. Genetics 92 823–832. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Greenberg, A. J., J. R. Moran, S. Fang and C.-I. Wu, 2006. Adaptive loss of an old duplicated gene during incipient speciation. Mol. Biol. Evol. 23 401–410. [DOI] [PubMed] [Google Scholar]
  28. Hahn, M. W., M. V. Han and S. G. Han, 2007. Gene family evolution across 12 Drosophila genomes. PLoS Genet. 3(11): e197. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Haldane, J. B. S., 1932. The Causes of Evolution. Cornell University Press, Ithaca, NY.
  30. Holt, R. A., G. M. Subramanian, A. Halpern, G. G. Sutton et al., 2002. The genome sequence of the malaria mosquito Anopheles gambiae. Science 298 129–149. [DOI] [PubMed] [Google Scholar]
  31. Hudson, R. R., 1983. Properties of a neutral allele model with intragenic recombination. Theor. Popul. Biol. 23 183–201. [DOI] [PubMed] [Google Scholar]
  32. Karpen, G. H., and A. C. Spradling, 1992. Analysis of subtelomeric heterochromatin in the Drosophila minichromosome Dp1187 by single P-element insertional mutagenesis. Genetics 132 737–753. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Langley, C. H., J. MacDonald, N. Miyashita and M. Aguadé, 1993. Lack of correlation between interspecific divergence and intraspecific polymorphism at the suppressor of forked region in Drosophila melanogaster and Drosophila simulans. Proc Natl Acad Sci USA 90(5): 1800–1803. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Langley, C. H., B. P. Lazzaro, W. Phillips, E. Heikkinen and J. M. Braverman, 2000. Linkage disequilibrium and the site frequency spectra in the su(s) and su(w(a)) regions of the Drosophila melanogaster X chromosome. Genetics 156 1837–1852. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Levis, R. W., R. Ganesan, K. Houtchens, L. A. Tolar and F. M. Sheen, 1993. Transposons in place of telomeric repeats at a Drosophila telomere. Cell 75 1083–1093. [DOI] [PubMed] [Google Scholar]
  36. Livak, K. J., and T. D. Schmittgen, 2001. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) method. Methods 25 402–408. [DOI] [PubMed] [Google Scholar]
  37. McVean, G., P. Awadalla and P. Fearnhead, 2002. A coalescent-based method for detecting and estimating recombination rates from gene sequences. Genetics 160 1231–1241. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Mechler, B. M., W. McGinnis and W. J. Gehring, 1985. Molecular cloning of the lethal(2)giant larvae, a recessive oncogene of Drosophila melanogaster. EMBO J. 4 1551–1557. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Morgenstern, B., 1999. DIALIGN 2: improvement of the segment-to-segment approach to multiple sequence alignment. Bioinformatics 15 211–218. [DOI] [PubMed] [Google Scholar]
  40. Muller, H.J., 1935. The origination of chromatin deficiences as minute deletions subject to insertion elsewhere. Genetica 17 237–252. [Google Scholar]
  41. Ohno, S., 1967. Sex Chromosomes and Sex-linked Genes. Springler-Verlag, Berlin.
  42. Rubin, G. M., 1978. Isolation of a telomeric DNA sequence from Drosophila melanogaster. Cold Spring Harbor Symp. Quant. Biol. 42 1041–1046. [DOI] [PubMed] [Google Scholar]
  43. Sokolova, K. B., and M. D. Golubovsky, 1979. Manifestation, interaction, and population distribution of alleles at the ‘giant larvae’ locus in Drosophila melanogaster. Genetika 15 233–243. [Google Scholar]
  44. Tajima, F., 1989. Statistical method for testing the neutral mutation hypothesis by DNA polymorphism. Genetics 123 585–595. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Thompson, J. D., D. G. Higgins and T. J. Gibson, 1994. Clustal W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res. 22 383–402. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Turner, T. L., M. T. Levine, M. L. Eckert and D. J. Begun, 2008. Genomic analysis of adaptive differentiation in Drosophila melanogaster. Genetics 179 455–473. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Vergnaud, G., 1999. Structure and evolution of human sub-telomeric regions. J. Soc. Biol. 193 35–40. [PubMed] [Google Scholar]
  48. Wang, X., W. E. Grus, and J. Zhang, 2006. Gene losses during human origins. PLoS Biol. 4(3): e52. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Walter, M. F., C. Jang, B. Kasravi, J. Donath, B. M. Mechler et al., 1995. DNA organization and polymorphism of a wild-type Drosophila telomere region. Chromosoma 104 299–341. [DOI] [PubMed] [Google Scholar]
  50. Winzeler, E. A., C. I. Castillo-Davis, G. Oshiro, D. Liang, D. R. Richards et al., 2003. Genetic diversity in yeast assessed in whole-genome oligonucleotide arrays. Genetics 163 79–89. [DOI] [PMC free article] [PubMed] [Google Scholar]

Articles from Genetics are provided here courtesy of Oxford University Press

RESOURCES