Open Access

Gene alterations at Drosophila inversion breakpoints provide prima facie evidence for natural selection as an explanation for rapid chromosomal evolution

BMC Genomics201213:53

DOI: 10.1186/1471-2164-13-53

Received: 24 August 2011

Accepted: 1 February 2012

Published: 1 February 2012



Chromosomal inversions have been pervasive during the evolution of the genus Drosophila, but there is significant variation between lineages in the rate of rearrangement fixation. D. mojavensis, an ecological specialist adapted to a cactophilic niche under extreme desert conditions, is a chromosomally derived species with ten fixed inversions, five of them not present in any other species.


In order to explore the causes of the rapid chromosomal evolution in D. mojavensis, we identified and characterized all breakpoints of seven inversions fixed in chromosome 2, the most dynamic one. One of the inversions presents unequivocal evidence for its generation by ectopic recombination between transposon copies and another two harbor inverted duplications of non-repetitive DNA at the two breakpoints and were likely generated by staggered single-strand breaks and repair by non-homologous end joining. Four out of 14 breakpoints lay in the intergenic region between preexisting duplicated genes, suggesting an adaptive advantage of separating previously tightly linked duplicates. Four out of 14 breakpoints are associated with transposed genes, suggesting these breakpoints are fragile regions. Finally two inversions contain novel genes at their breakpoints and another three show alterations of genes at breakpoints with potential adaptive significance.


D. mojavensis chromosomal inversions were generated by multiple mechanisms, an observation that does not provide support for increased mutation rate as explanation for rapid chromosomal evolution. On the other hand, we have found a number of gene alterations at the breakpoints with putative adaptive consequences that directly point to natural selection as the cause of D. mojavensis rapid chromosomal evolution.


Inversion breakpoints mutation rate chromosomal evolution transposable elements gene duplication gene transposition position effects


Chromosomal inversions are a common feature of genome evolution in many groups of animals and may play a significant role in adaptation, speciation and sex chromosome evolution [14]. The rate of rearrangement fixation varies significantly within and between animal groups [2, 5]. The genus Drosophila shows one of the highest rates in all eukaryotes [68] at least partially because special cytological mechanisms in Diptera allow heterozygotes for paracentric inversions to circumvent the production of aneuploid gametes [1]. A striking extent of variation in rearrangement rate has been reported among different Drosophila lineages [6, 912]. For instance, the fixation rate of inversions is higher in the Sophophora subgenus than in the Drosophila subgenus [10]. Also particular lineages such as D. miranda or D. yakuba exhibit an unusually rapid rate of chromosomal evolution [9, 11]. Four factors may contribute to the variation among lineages in the rate of chromosomal rearrangement: generation time, population size, mutation rate and fitness effects of rearrangements. However, the actual reason for such variation is unclear and different studies invoke different explanations [912].

Chromosomal inversions can be generated by two major mechanisms. The first of them is ectopic recombination (or non-allelic homologous recombination, NAHR) between transposable elements (TEs) [1315], segmental duplications [16, 17] or short repeat sequences [18]. When ectopic recombination occurs between two copies of a TE inserted in opposite orientation at two different chromosomal sites, the resulting inverted chromosomal segment will be flanked by two chimeric TE copies bounded by exchanged target site duplications (TSD) [14, 15]. The second mechanism is chromosomal breakage and erroneous repair of the free ends by non-homologous end-joining (NHEJ) [19]. Breakages can be simple double-strand breaks (DSB) or staggered single-strand breaks (SSB). In the second case, the consequence is the generation of inverted duplications at both sides of the inverted segment [11, 20]. Thus, inversions generated in this way can be recognized by duplicated DNA segments (originally single-copy) in inverted orientation flanking the inverted chromosomal segment. The relative contribution of the two mechanisms to the generation of natural Drosophila inversions is not yet clear. In Dipterans, clear-cut evidence for the implication of TEs in their generation has been found for a few polymorphic inversions [15, 2123] but has never been found for fixed inversions [6, 11, 24, 25]. On the other hand, breakage and repair by NHEJ may be the prevalent mechanism in D. melanogaster and its close relatives [11].

Several explanations have been put forward for the spread of inversions in populations [3]. Although in principle inversions could be neutral or underdominant and spread by genetic drift, this is probably unusual in Drosophila species given their elevated effective population size, of the order of 106[26, 27]. The traditional explanation for the adaptive significance of inversions is based on their recombination-reducing effect [28] that keeps together alleles at loci with epistatic effects on fitness, the "coadaptation" hypothesis [29]. An alternative model proposes that inversions capture a set of locally adapted genes and protect them from recombination with immigrant chromosomes [4, 30]. Finally, inversions may spread in populations due to the direct mutational effects associated with their breakpoints, the "position effect" hypothesis [31]. This latter hypothesis has received so far little attention [32] but the relatively high gene density and compact structure of Drosophila genome (> 90% of euchromatin has functional annotations) [33, 34] make position effects most likely. Available genomic sequences [35] provide the opportunity to investigate the structure of inversion breakpoints and ascertain their functional consequences.

Drosophila mojavensis has been an excellent model for the study of the genetics of ecological adaptation and speciation for more than fifty years [3638] and it is now a useful model for genomic studies as the complete genome sequence is available [35, 39]. D. mojavensis is a cactophilic species in the repleta group endemic to the deserts of the Southwestern USA and Northwestern Mexico, chiefly the Sonoran Desert (Arizona, Baja California and Sonora) the Mojave Desert and Santa Catalina Island in southern California. Natural populations are genetically differentiated and use different primary host plants, Stenocereus gummosus (pitaya agria) in Baja California, Stenocereus thurberi (organ pipe) in Arizona and Sonora, Ferocactus cylindraceous (California barrel) in Southern California and Opuntia spp. on Santa Catalina Island [4042]. The ecological conditions of the Sonoran Desert are extreme (dry, arid and hot according to Köppen classification [43]) as attested by the fact that only four Drosophila species are endemic [41]. Accordingly, D. mojavensis is unusually thermotolerant and desiccation resistant [4447]. In addition, D. mojavensis is the exclusive inhabitant of its chief host plants over most of its distribution range, in part because they contain large amounts of unusual lipids and triterpene glycosides that make them unsuitable for other Drosophila species [48, 49].

The salivary gland chromosomes of D. mojavensis and its close relatives D. arizonae and D. navojoa were cytologically analyzed and the D. mojavensis standard chromosomal arrangement seemingly contain ten fixed inversions compared to Primitive I (the ancestor of the repleta group), one in chromosome X (Xe), seven in chromosome 2 (2c, 2f, 2g, 2h, 2q, 2r and 2s) and two on chromosome 3 (3a and 3d) [50, 51]. Five inversions (3d, Xe, 2q, 2r and 2s) are exclusive to D. mojavensis whereas the rest are shared by other cactophilic Drosophila of the mulleri complex (see Figure 1). Thus, D. mojavensis is a chromosomally derived species that contains the highest number of fixed inversions in the entire mulleri complex [52]. Only one of D. mojavensis inversions (Xe) has been previously characterized at the molecular level [53]. Here we characterize all inversions fixed in D. mojavensis chromosome 2, the most dynamic of the five major chromosomes, and explore the causes of its rapid chromosomal evolution. Using comparative mapping of BAC-end sequences from D. buzzatii onto the D. mojavensis genome (see Figure 1), we identify the breakpoint regions of all inversions. We then annotate them by comparison with the genome of D. virilis, the closest relative with a sequenced genome [35] that represents the ancestral (non-inverted) arrangement. Our results provide information on the multiple causes that generated these inversions, reveal unreported associations of inversion breakpoints with duplicated and transposed genes, and shed light on the functional consequences of D. mojavensis inversions. Overall, our results suggest that rapid chromosomal evolution in D. mojavensis is not due to an increase in the rate of inversion generation but to its adaptation to the extremely harsh environment of the Sonoran Desert that was accompanied by strong natural selection.
Figure 1

Phylogenetic relationships and divergence times for seven species of the Drosophila subgenus. Six species (D. buzzatii, D. longicornis, D. mojavensis, D. arizonae, D. navojoa and D. mulleri) belong to the repleta species group and the chromosome 2 inversions fixed in the D. mojavensis and D. buzzatii lineages are indicated [51, 52]. Primitive I is the most recent common ancestor of D. mojavensis and D. buzzatii [52]. D. virilis is the outgroup species and belongs to the virilis species group. Phylogenetic relationships and divergence times are taken from [38, D.C.S.G. Oliveira, F.C. Almeida, P. O'Grady, W.J. Etges, M.A. Armella and R. DeSalle, personal communication].


Identification of syntenic segments and breakpoint regions

We sequenced the ends of 1,152 D. buzzatii chromosome 2 BAC clones [54] and 1,870 BAC-end sequences (BES) mapped onto D. mojavensis chromosome 2 (see Methods for details). By comparing the chromosomal localization of the markers, we identified 20 syntenic segments (Additional file 1). D. mojavensis scaffold 6540, corresponding to chromosome 2 [55], is 34,148,556 bp long (coordinates begin at centromere). The most proximal marker in our map (segment 20) was located at position 1,721,255 bp whereas the most distal marker (segment 1) was located at position 34,039,404, i.e. only 109 kb from the end of the scaffold. The largest segment was number 15 with 5,926.5 kb and 426 markers whereas the smallest one was number 16 with 50.5 kb and 9 markers. The second-smallest segment was number 7 with 80.7 kb and 2 markers. This latter segment was exceptional as it was detected using comparative information from BAC clone 1B03 that has been fully sequenced [56]. In general, the markers were distributed homogeneously along the chromosome as indicated by the highly significant correlation (r2 = 0.95, P < 0.001) between segment size and number of markers. The 20 syntenic segments amount to 30,830,590 bp, representing ~90.3% coverage of chromosome 2. The missing 3,317,966 bp are distributed between the endmost chromosomal regions (~5.3%) and the 19 breakpoint regions (~4.4%).

Estimating the genomic distance

The order, size and orientation of the 20 conserved syntenic segments are shown in Figure 2. This breakpoint graph [57] contains nine cycles (represented with different colors), namely eight rectangles and a more complex cycle comprising two concatenated rectangles, suggesting that eight inversions and a more complex rearrangement are fixed in chromosome 2 since the divergence between D. buzzatii and D. mojavensis. GRIMM software [58] indicated that a minimum of 10 inversions are needed to transform the D. buzzatii chromosome 2 into that of D. mojavensis (Figure 3). Because there are 20 syntenic segments and 19 breakpoints, this implies one breakpoint reuse. Previous work in our laboratory [12] determined that three inversions, 2m, 2n and 2z7, have been fixed in chromosome 2 of D. buzzatii since its divergence from Primitive I, the most recent common ancestor with D. mojavensis (Figure 1). Furthermore, the breakpoints of these three inversions have been isolated and sequenced [59]. Inversions 2m and 2n are arranged in tandem and share the middle breakpoint. Thus we identified the complex cycle in the breakpoint graph (Figure 2) as corresponding to the 2mn rearrangement and determined that seven inversions have been fixed in D. mojavensis since divergence from Primitive I. These seven inversions entail 14 breakpoints, i.e. they have independent breakpoints. GRIMM software [58] was run again to compare the arrangement of D. mojavensis chromosome 2 with that of Primitive I (inferred by subtraction of the three inversions fixed in D. buzzatii). The result was the single scenario shown in Figure 3.
Figure 2

Dot plot comparing the order of chromosome 2 markers between D. mojavensis and D. buzzatii. The x-axis represents the position of all markers in the physical map of D. buzzatii [54], while in the y-axis markers were ordered according to their coordinates in D. mojavensis scaffold 6540. Each syntenic segment is identified by a number with inverted segments indicated by a negative sign. Also shown is the breakpoint graph [57] generated by connecting consecutive syntenic segments along D. buzzatii chromosome 2 with continuous lines and those along the D. mojavensis chromosome 2 with dashed lines. The result is eight rectangles and a more complex cycle comprising two concatenated rectangles (yellow) that correspond to the 10 inversions fixed between the two species.
Figure 3

Order and orientation of the 20 chromosome 2 conserved segments during the divergence between D. buzzatii and D. mojavensis. The genomic distance between the two species calculated using GRIMM software was 10 inversions. When inversions 2m, 2n and 2z 7 fixed in D. buzzatii [12, 59] were subtracted, GRIMM software yielded a single scenario with seven inversions (2c, 2f, 2g, 2h, 2q, 2r and 2s) fixed from Primitive I to D. mojavensis.

In order to compare the inversions proposed by GRIMM with those detected previously using cytological methods (see Introduction), we located on the D. buzzatii physical map [54] those clones that mapped on each D. mojavensis breakpoint region and identified the chromosomal bands involved in each case. We corroborated the inversion breakpoints identified by cytogenetics and those detected by bioinformatics with an accuracy of three to five bands. Rearrangements detected cytologically and those proposed by GRIMM (Figure 3) did not only match in number but also the regions involved in each of them were in agreement, allowing for the differences between the precision of both techniques. However, three cytological breakpoint coincidences were not corroborated at the sequence level. The general agreement between cytogenetics and bioinformatics is remarkable because often these two approaches to chromosomal evolution seem to provide discordant results [60, 61]. For instance, in Drosophila, comparative mapping has sometimes revealed fixed inversions overlooked by previous cytological studies [11, 62, 63].

Delimitation and annotation of breakpoint regions

Among the seven chromosome 2 inversions fixed in the D. mojavensis lineage, three (2f, 2g and 2c) are shared between diverse species of the mulleri complex and must be between 7 and 11 myr old (Figure 1); another one (2h) is shared between D. mojavensis and D. arizonae only and should be between 2 and 6 myr old (Figure 1); the remaining three inversions (2q, 2r and 2s) are exclusive of D. mojavensis and thus must be relatively young (less than 2 myr, Figure 1). We initially identified the 14 breakpoint regions of these seven inversions as those sequences between syntenic segments (Additional file 2). These regions varied between 9,776 bp and 480,695 bp. In order to narrow down the size of these regions, the corresponding sequences were blasted against the D. virilis genome (see Methods), which represents the parental (non-inverted) chromosome (Figure 1). We expect that breakpoint regions for each inversion will appear in D. mojavensis genome as AC (distal) and BD (proximal) but in D. virilis genome as AB (distal) and CD (proximal). Similarity comparisons of AC, BD, AB and CD sequences allowed us to reduce the size of the breakpoint regions to between 259 bp and 91,812 bp, on average 8.3% of the original breakpoint regions (Additional file 2). Five breakpoint regions were further reduced to about 71.1% of their previous size (on average) by excluding the coding sequences of orthologous genes. Once the new limits for the 14 breakpoint regions in D. mojavensis were established, we analyzed the similarity between the two breakpoint regions of each inversion using BLAST 2 sequences [64]. A summary list of the genes adjacent to the 14 inversion breakpoints is shown in Table 1 and a detailed annotation of the breakpoint regions of each inversion is shown in Figures 4, 5 and 6 for the most recent inversions (2s, 2r and 2q) and Additional files 3, 4, 5 and 6 for the rest. TE content of all the breakpoint sequences (see Methods) is summarized in Additional file 7. Our analysis of the breakpoints provides significant information on the causes and consequences of the seven chromosome 2 inversions fixed in D. mojavensis (Table 1) that we present in the following sections.
Table 1

Chief features of inversion breakpoint regions in D. mojavensis


Breakpoints and adjacent protein-coding genes in D. mojavensis

TE copies at co-occurrent breakpoints

Inversion-associated inverted duplications

Preexisting duplications in parental genome

Transposition-associated genes and D. virilis lineage specific genes (underlined)

Gene gains (bold) and putative position effects









GstD1a GstD1b


















Dmoj\GI22722 CG32344

CG2846 Dvir\GJ23779









7.1 kb from AB











1 kb from AB

4.3 kb from CD















Histone clusters
















Figure 4

Annotation of inversion 2s distal and proximal breakpoint regions in D. virilis (non-inverted chromosome) and D. mojavensis (inverted chromosome). Genes are depicted as solid boxes (exons) linked by polygonal lines (introns) with the 5' and 3' ends showing the direction of transcription. Genes adjacent to the distal (AB) and proximal (CD) breakpoints of D. virilis are colored in green and blue, respectively. Orthologs in the distal (AC) and proximal (BD) D. mojavensis breakpoint regions are colored accordingly. Red curly brackets with an arrow indicate the breakpoint junctions. TE insertions are shown as solid rectangles: blue (Bu T5), purple (Homo 3) or yellow (Galileo). Some TE insertions are flanked by TSDs insertions depicted in boxes above (or below) them. Dotted sections are not drawn to scale.
Figure 5

Annotation of inversion 2r distal and proximal breakpoint regions in D. virilis (non-inverted chromosome) and D. mojavensis (inverted chromosome). TE insertions shown as solid rectangles: yellow (Galileo), green (Invader) or brown (Homo6). The black box in the D. mojavensis BD region represents a 90.2 kb-block containing interspersed histone clusters and TEs. Other symbols as in Figure 4.
Figure 6

Annotation of inversion 2q distal and proximal breakpoint regions in D. virilis (non-inverted chromosome) and D. mojavensis (inverted chromosome). Inverted duplications in the D. mojavensis breakpoints are enclosed within dotted boxes, light blue (1 kb) or orange (4.3 kb). These duplications were presumably generated by staggered single-strand breaks in the parental chromosome represented by dotted red lines flanked by red arrows. Dmoj\GI22075, a novel gene not present in D. virilis breakpoint regions, was generated by the partial duplication of CG1208. Two unidentified TE insertions are found between the duplicated sequences in D. mojavensis BD region. Other symbols as in Figure 4.

Generation of chromosomal inversions

In order to test for the implication of TEs in the generation of the seven inversions, we analyzed the TE content of the breakpoint regions and detected copies of a TE at both co-occurrent breakpoints in three inversions: 2s, 2r and 2c (Table 1). One of them, inversion 2s, provides compelling evidence for the implication of the transposon BuT5[65] in its generation. At the distal breakpoint, a 981-bp copy of BuT5 was found bounded by the 9-bp sequences AAGGCAAGT and CTGTATAAT (Figure 4). At the proximal breakpoint, we uncovered a 27-bp BuT5 fragment comprising 12 bp identical to one end and 15 bp identical to the other end, and thus resembling the footprints that transposons often leave behind at the donor site following excision [66, 67] bounded by the 9-bp sequences ACTTGCCTT and ATTATACAG. These sequences can be interpreted as TSD produced at the time of the transposon insertion and its exchanged arrangement (ACTTGCCTT and CTGTATAAT are the inverted complementary versions of AAGGCAAGT and ATTATACAG, respectively) provides unequivocal evidence for the generation of inversion 2s by ectopic recombination between BuT5 copies. Recently work in our laboratory has shown that an inversion fixed in D. uniseta, another related species that belongs to the buzzatii complex [56], has also been generated by ectopic recombination between BuT5 copies.

Two of our inversions provide evidence for generation by chromosomal breakage and erroneous repair by NHEJ [11, 19, 20] (Table 1). In the breakpoints of inversion 2h we found evidence for a duplication of a 7.1-kb long segment containing three genes, CG1792, Dmoj\GI23402 and pasha (Additional file 3). When the two D. mojavensis breakpoint regions AC and BD were compared, we uncovered 10 blocks of similarity (E-value ≤ 9e-10) 45 to 641-bp long. These blocks are scattered in a 7,151-bp segment containing genes CG1792, Dmoj\GI23402 and pasha in the AC breakpoint, and within a 2,676-bp segment including Dmoj\GI23123 in the BD breakpoint (Additional file 3). This duplication can be explained by staggered SSB at the distal breakpoint in the parental chromosome. In the distal breakpoint of the derived chromosome (AC) the segment seems intact and the three genes fully functional whereas in the proximal breakpoint (BD), this segment has been reduced to 2.7 kb by several deletions (Additional file 3). Because the duplication was caused by the inversion, we estimated the age of the 2h inversion using the divergence of those fragments non-coding for proteins and the Drosophila neutral substitution rate of 0.0111 [68] as 4.4 myr, which is in agreement with the phylogenetic distribution of inversion 2h (Figure 1). In the breakpoints of inversion 2q there are also inverted duplications. In this case, staggered SSB likely occurred at both breakpoints involving a ~1-kb segment at distal breakpoint (AB) and a 4.3-kb segment at proximal breakpoint (CD) (Figure 6). We estimated the age of inversion 2q using the same procedure as before as 1.4 myr, a figure fully compatible with the phylogeny (Figure 1). There is a third case, where duplicated genes (GstD1) in opposite orientation are found at the two inversion 2c breakpoints (Additional file 6). However, this observation is best interpreted as a breakage occurring at a preexisting duplication (see below).

Preexisting gene duplications at breakpoints

We found four cases where inversion breakpoints fall between duplicated genes, i.e. there were preexisting gene duplications at the breakpoint regions (Table 1). In order to determine if this is in concordance with the random expectation, we estimated the number of intergenic regions localized between duplicated genes in D. mojavensis chromosome 2. Genes in this chromosome encode 3,407 out of the 14,595 D. mojavensis predicted proteins (23.34%). Thus there are 3,406 putative intergenic regions in this chromosome. According to the previous established criteria to consider two genes as duplicated copies (see Methods), we detected 215 intergenic regions between duplicated genes along the entire chromosome. We compared the two proportions by three different statistical methods. The χ2 test with and without Yates correction (χ2 = 11.526, P = 0.0007 and χ2 = 8.111, P = 0.0044, respectively) indicated that 4/14 is significantly higher than the proportion expected at random (215/3406). Fisher exact test (P = 0.0098) corroborated this result. It could be argued that breakpoints are distributed at random in non-coding intergenic regions and that duplicated genes accumulate more breakpoints because their mean intergenic distance is longer than that between non-duplicated genes. We tested this possibility by calculating the intergenic distance for both duplicated and non-duplicated genes in D. mojavensis chromosome 2. A t-test showed that means are significantly different (t = 3.84, P = 0.0001), but the mean distance between duplicated genes is actually the shortest one. Thus, we conclude that there is an excess of breakpoints localized between duplicated genes with respect to the random expectation. Previously, another D. mojavensis inversion (Xe) was found to have a breakpoint adjacent to a gene duplication [53] and in primates, rearrangement breakpoints have been sometimes observed in the midst or adjacent to clustered gene families [69, 70].

Two explanations can be put forward for these observations. Firstly, duplicated genes might cause instability and increased rate of DBS [71] or might be breakage "permissive" regions [70]. Alternatively, we suggest that the mobilization of a duplicated gene may entail in some cases beneficial position effects that might help the inversion to be fixed within the species. Two duplicated genes may have their evolution constrained because of shared regulatory sequences, their co-location in the same chromatin regulatory domain, or sequence homogenization by frequent conversion and ectopic recombination events. The re-location of one of the copies to a different chromosomal region might produce beneficial changes in the regulation of expression for one or the two copies and/or release them from evolutionary constraints (see below).

Association of inversion breakpoints with gene transposition events

Gene content of chromosomal elements is generally conserved in the genus Drosophila although gene order has been scrambled extensively by fixed paracentric inversions [6]. However, there have been a number of genes that have been relocated between or within chromosomal elements by gene transposition or retroposition [7274]. We searched the 28 genes adjacent to D. mojavensis inversion breakpoints in the 12 Drosophila genomes [35] for evidences of gene transposition and found that there are three genes involved in interchromosomal transposition events (Lsp1beta, Dmoj\GI22722 and CG32344) and another one (Dmoj\GI24456) is likely involved in a intrachromosomal transposition (Table 1). Furthermore, two genes are present in the D. virilis breakpoints regions but not in D. mojavensis and thus are likely also transposed genes. Finally, a large DNA block including several clusters of Histone genes have been inserted in the proximal breakpoint of inversion 2r (Figure 5).

In order to test for an association of inversion breakpoints and gene transpositions, we first determined that 69 out of 514 interchromosomally relocated genes [73] are located in D. mojavensis chromosome 2. Then we compared our proportion of interchromosomal gene transpositions (3/28) to the general chromosome 2 proportion (69/3,407). The χ2 test with and without Yates correction (χ2 = 6.42, P = 0.0113; χ2 = 10.22, P = 0.0014), and the Fisher exact test (P = 0.0199) indicated that 3/28 is significantly higher than the relation expected at random. This test is conservative as we did not take into account the putative intrachromosomal transposition of Dmoj\GI24456, the two D. virilis lineage specific genes or the 90-kb insertion at 2r proximal breakpoint (see below). The association of inversion breakpoints and transposed genes is likely the result of the "fragility" or "permissivity" of these regions [8]. A clear example is the 2g distal breakpoint region in D. virilis where three genes (Dvir\GJ23449, Dvir\GJ23779 and CG32344) have transposed to this region from different sources.

The 2r proximal breakpoint (BD) harbors a big block of DNA (~90-kb) not found in any of the D. virilis breakpoints. This block contains at least five tandemly arranged copies of the Histone gene cluster [75, 76]. The exact number of copies cannot be determined due to the presence of a ~10 kb sequence gap bounded by histone genes from different clusters. The block also comprises a large number of fragments annotated as repetitive sequences (110 ReAS elements that amount up to ~45% of the sequence). These elements tend to occur at regular intervals with a periodicity similar to that of the Histone gene clusters. In D. melanogaster the Histone complex (HIS-C) is located in chromosomal arm 2L (Muller element B) and comprises ~100 tandemly arranged copies of a cluster containing five Histone genes (His1, His2B, His2A, His4 and His3) [75, 76]. Histone genes are often involved in transposition events. In the repleta group species, the ancestral and chief HIS-C (named HIS-C1) is likely located at chromosome 3, but there are other derived and probably smaller complexes (named HIS-C2) at chromosomes 3 and 4, implying at least two transposition events [62]. The insertion of a ~90-kb block containing several Histone gene clusters in the 2r proximal breakpoint (BD) seems to represent yet another transposition event, which is probably specific to D. mojavensis. This block is not found in D. virilis in any of the two breakpoints and was not found in D. mulleri or D. buzzatii by in situ hybridization [62]. We suggest that the occurrence of this ~90-kb block is the result of the reintegration of an extrachromosomal circular DNA fragment (eccDNA) replicated by rolling circle replication [77] perhaps at the time of the inversion generation (when DSBs were available). This hypothesis explains the fact that this large insertion contains tandemly repeated coding (Histone) genes and TEs.

Gene gains and changes in gene structure and/or expression

Two novel genes have been generated at the D. mojavensis breakpoints (Table 1). The gene Dmoj\GI23123, localized at 2h proximal breakpoint (BD), comprises two exons encoding a 94-aa protein (Additional file 3). A similarity search indicated that it is related to the gene pasha (partner of drosha, CG1800), that is found in the distal breakpoint (AC). pasha has five exons and encodes a 655-aa protein with a double-strand RNA binding domain that is involved in primary miRNA processing, among other biological processes. Amino acid identity between Dmoj\GI23123 and pasha proteins is 93.5% over a 46-aa segment. Gene Dmoj\GI23123 has an unknown molecular function but a protein domain PTHR13482 involved in nucleic acid binding was detected with Interproscan [78]. In addition it is expressed according to modENCODE D. mojavensis DB, suggesting it is fully functional. This gene arose at the time of the inversion generation as a consequence of the duplication of a 7.1-kb segment originally containing three genes: CG1792, Dvir\GJ23094 and pasha (Additional file 3). Seemingly the duplicated copies of CG1792 and Dvir\GJ23094 were partially lost by deletion whereas the duplicated copy of pasha evolved into the novel gene Dmoj\GI23123.

Another novel gene, Dmoj\GI22075, is found at the distal breakpoint (AC) of inversion 2q (Figure 6). It arose when this inversion was generated as a consequence of the duplication of a 4.3-kb segment containing a fragment of gene CG1208. This gene encodes a 508-aa protein that has glucose transmembrane transporter activity. Dmoj\GI22075 comprises three exons encoding a 153-aa protein with a 75-aa Major Facilitator Superfamily (MFS) domain [79]. The conservation of this domain indicates that it is a new functional gene and suggests that it has retained a MFS function.

Three inversions entail putative changes in gene structure and/or expression. Two GstD1 genes in opposite orientation were found at the two D. mojavensis breakpoints of inversion 2c while only one is present in the proximal breakpoint (CD) of the D. virilis chromosome (Additional file 6). In order to ascertain the origin of these two genes, a phylogeny of GstD genes in D. mojavensis and D. virilis was built (Additional file 8). The two Dmoj\GstD1 genes are co-orthologs of the Dvir\GstD1 gene and we estimated the age of the duplication event that generated them (using divergence at synonymous sites) as 16 myr. Therefore, this duplication event took place before inversion 2c and the inversion breakpoint occurred between two pre-existing duplicated GstD1 genes. GstD1 genes have been associated with the detoxification of insecticides as well as other chemical substances present at larval food sources [80]. Low et al. [81] detected that positive selection has operated on GstD1 and identified the parallel evolution of a radical glycine to lysine amino acid change (K171) in D. melanogaster, D. pseudoobscura and D. mojavensis. Matzkin [82] found additional evidence for the adaptive evolution of Dmoj\GstD1a, a gene that shows changes of expression level in response to the use of different host plants as larval substrates [83]. Inversion 2c relocated GstD1a to a new chromosomal region and left the other copy GstD1b in the original position. This might have triggered changes in their gene expression regulation and/or evolutionary constraints. The two D. mojavensis GstD1 proteins differ by 14 aa including the critical 171 residue (where GstD1a has lysine but GstD1b has glutamic acid). In addition, according to D. mojavensis modENCODE DB the relocated GstD1a gene has seemingly a much higher expression level than the gene in the original location, GstD1b. We suggest that the GstD1 duplication and subsequent separation of the two copies by inversion 2c may have had significant consequences for the adaptation of the lineage of D. mojavensis and related species of the mulleri complex to its cactophilic niche (Figure 1).

The 2r distal breakpoint was localized in D. virilis between two Hsp68 genes oriented head-to-head (Figure 5). These two genes have the same structure and size (a single exon 1,935-bp long encoding a 644-aa protein) and nearly identical sequence (8 mismatches, 99.6% identity). However, in D. mojavensis Hsp68a (661 bp) is significantly shorter than Hsp68b (1,935 bp) and posses two exons encoding a 152-aa protein (Figure 5). The two genes only show conservation of a segment encoding 90-aa corresponding to a Heat Shock Protein domain (75% aa identity). We built a phylogeny of Hsp68 in 11 Drosophila genomes (D. willistoni is the only of the 12 species lacking Hsp68, Additional file 9). While a single Hsp68 gene is present in the six melanogaster group species, two copies oriented head-to-head are found in D. pseudoobscura, D. persimilis, D. grimshawi and D. virilis. Thus, this is likely to be the ancestral state. Nonetheless the phylogenetic tree shows a high similarity between the two Hsp68 copies present within each of these four species (Additional file 9) that can be interpreted as the result of concerted evolution by recurrent gene conversion or ectopic recombination [84]. In D. mojavensis, inversion 2r relocated Hsp68b to a new chromosomal site along with its upstream regulatory sequences. A detailed sequence analysis confirms that the Dmoj\Hsp68b 5' upstream region harbors two cis-regulatory motifs called HSEs (heat shock elements) modulating the expression of this gene [85]. But we also detect a third HSE, 683 bp upstream of the Dmoj\Hsp68b 5' region, in opposite orientation to the previous two HSEs. This putative cis- regulatory motif is likely to correspond to the HSE of Dmoj\Hsp68a, apparently dragged by the inversion to the BD region upstream of Dmoj\Hsp68b. In addition, only ~2.5 kb upstream of this gene is the ~90-kb block of Histone genes and TEs (see above). Because TEs may influence chromatin organization [86] and this in turn is a significant determinant of gene expression [87, 88], the insertion of this block is likely to have altered the expression level and/or pattern of Dmoj\Hsp68b. No promoter or regulatory HSE sequences were detected upstream of Dmoj\Hsp68a but according to D. mojavensis modENCODE DB this gene is being transcribed. It may be that it has recruited a new promoter (e.g. a fragment of the transposon Galileo located 3-bp from the initial codon; see Figure 5) and acquired a new function or it is on the way to becoming a pseudogene. It must be recalled that Dmoj\Hsp68a shows an altered structure and a high rate of sequence divergence (Additional file 8). In summary, we found that inversion 2r has induced significant alterations of this gene in both structure and expression.

A footprint of a BuT5 was found in the D. mojavensis proximal breakpoint of inversion 2s, 121 bp from the start codon of CG10375 (Figure 4). We used McPromoter[89] to look for the Dmoj\CG10375 promoter. A unique putative promoter region was located 115-bp 5' from the start codon. This putative promoter region (~100-bp) includes the BuT5 footprint and has a peak with high score (0.0505) located in region B (across the breakpoint). In addition it corresponds to a model 1 promoter (DNA replication related element). These observations contrast with the promoter region of Dmel\CG10375 that is model 3 (Motif6/Motif1) and has a narrow peak with a lower score (0.03925) and imply that the 2s inversion and the BuT5 element have likely altered the expression of Dmoj\CG10375, presumably increasing it. Gene CG10375 has a single DnaJ domain and is the likely orthologous of human DNAJC8 gene, a member of the Hsp40 family.


In this study, we investigated the rapid chromosomal evolution of the D. mojavensis lineage that has fixed ten paracentric inversions since the repleta group ancestor, ~12 mya (Figure 1). Using D. buzzatii BAC-end sequences [54] and the genome sequences of D. mojavensis and D. virilis[35] we mapped, identified, annotated and analyzed all breakpoints of the seven inversions fixed in D. mojavensis chromosome 2, the most dynamic element. The results corroborated previous cytological analyses [51] and allowed us to provide significant information on the causes and consequences of these structural changes.

One hypothesis that may explain an accelerated chromosomal evolution rate is an increased mutation rate that generates more rearrangements per generation. This possibility was invoked to explain the high rate of chromosomal rearrangement between D. miranda and D. pseudoobscura[9]. Because inversions may be generated by TEs (see Introduction), one possible cause of high mutation rate is an increased transpositional activity. Therefore, it has been suggested that variation in transpositional activity of TEs might contribute to variation in rates of rearrangement fixation [12]. However, an increased mutation rate could also be due to the presence of other causes, both intrinsic and extrinsic (e.g. clastogenic chemicals or ionizing radiation). Overall, our results do not support this hypothesis because the inversions fixed in D. mojavensis seem the result of multiple generation mechanisms. We found direct evidence for the implication of transposon BuT5 in the generation of inversion 2s and only circumstantial evidence for the implication of the transposons BuT5 and Galileo in inversions 2c and 2r, respectively. Inversions 2h and 2q harbor inverted duplications of non-repetitive DNA at the two breakpoints and were likely generated by staggered single-strand breaks and repair by non-homologous end joining. Finally, no definitive conclusion can be drawn about the generation of inversions 2f and 2g. It could be argued that the latter inversions might have been generated by TEs but subsequent changes in the breakpoint regions hindered our ability to find conclusive evidence for their implication. TE copies might have excised and move to other locations after generating the inversion (a hypothesis known as "hit-and-run" [24]), or be deleted due to the high rate of loss of nonfunctional DNA in Drosophila [90, 91]. However, in the absence of supporting evidence we think that such inference is unwarranted.

In any case, the generation of inversion 2s by transposon BuT5 is a significant finding because, in Dipterans, the implication of TEs in the generation of chromosomal inversions has been demonstrated for a few polymorphic rearrangements but never for fixed inversions (see Introduction). BuT5 is a MITE with unusual features [N. Rius, A. Delprat and A. Ruiz, personal communication]. It was discovered in D. buzzatii[65] but is present in the genome of most repleta group species, implying that it was probably already present in the ancestor ~16 mya [N. Rius, A. Delprat and A. Ruiz, personal communication]. In D. mojavensis is relatively abundant and transpositionally active but copy density in the dynamic chromosome 2 is not significantly higher than in the rest of chromosomes. These observations do not support the increased mutation hypothesis.

A second explanation for accelerated chromosomal evolution is an increase of the species' population size because the rate of fixation of selectively advantageous rearrangements is a direct function of population size [26]. The high rate of chromosomal evolution of the D. yakuba lineage in comparison with the D. melanogaster lineage was attributed to differences in population size [11]. The effective population size of D. mojavensis has been estimated as ~106 yet there is variation between populations in Baja California and Mainland Sonora [92, 93]. However, there is no reason to assume that this is an unusually high figure. Population size of D. arizonae, its closest relative (Figure 1), is seemingly higher (or at least not lower) than that of D. mojavensis[92, 93]. In contrast to D. mojavensis, which is fixed for five species-specific inversions, D. arizonae has only one [51]. Therefore, population size does not provide an adequate explanation for D. mojavensis rapid chromosomal evolution.

The third hypothesis is strong natural selection in a new environment that increases the number of fixed inversions. D. mojavensis is the only mulleri complex species inhabiting the Sonoran Desert. Other species of this complex, including its closest relatives D. arizonae and D. navojoa (Figure 1), live in less harsh environments of central and southern Mexico. Thus it must be presumed that adaptation to the extreme conditions of the Sonoran desert and to the exclusive host plants exploited by D. mojavensis must have required many adaptive genetic changes. Chromosomal inversions in Drosophila have been considered for decades as adaptive devices that spread in natural populations driven by natural selection (see Introduction). In fact there is ample evidence for the adaptive significance of polymorphic inversions (those that are segregating within species) but no such evidence has been provided for fixed inversions (those that appear as interspecific differences). We have found a variety of gene alterations at the breakpoints of D. mojavensis chromosome inversions and propose that these alterations contributed to their adaptive value. Overall, strong natural selection in a new harsh environment seems the most plausible cause for D. mojavensis rapid chromosomal evolution.

The alterations associated with the breakpoints of five D. mojavensis inversions include two gene gains (Dmoj\GI23123 and Dmoj\GI22075) and three putative alterations of gene structure and/or expression regulation (Table 1). We discuss these effects in turn. In D. mojavensis two new genes were generated associated to inversions 2q and 2h. As a consequence of the generation mechanism, staggered breakage and NHEJ repair, duplications of single-copy DNA were present at the breakpoints of these inversions at the onset. In the case of inversion 2q this duplication included gene CG1208 (except for its first exon and upstream sequences, see Figure 6). The novel gene Dmoj\GI22075 is shorter than the original gene CG1208 but retains a MFS domain and could function as a sugar transmembrane transporter (if a new promoter has been recruited). In the case of inversion 2h the duplicated segment included originally three genes (see Additional file 3). Only one gene (Dmoj\GI23123) seems to have survived. This gene is related to pasha (a gene involved in primary microRNA processing and gene silencing by miRNA) and according to modENCODE data, it is expressed. We suggest that novel genes might have contributed to the adaptive value of these inversions. Novel genes are widely recognized as a source of new functions [94] but inversion-associated duplication has not been considered a molecular mechanism that can generate new genes until very recently and only in prokaryotes [20].

The two most recent inversions, 2r and 2s, that are exclusive to D. mojavensis, show putative alterations of structure and/or expression of heat shock protein (Hsp) genes. Hsp genes encode intra-cellular chaperones for other proteins and have been established as potential candidates for thermotolerance [95]. Hsp family harbors genes constitutively or inducibley expressed [96]. Heat-inducible genes are regulated by heat shock factor (HSF), which binds to HSE sequences [97] whereas other heat shock genes have an Hsf-independent regulation [98]. The distal breakpoint of inversion 2r separated two previously linked and very similar Hsp68 genes (Figure 5). One of them, Hsp68a, remained in its original location but suffered a radical change in structure and sequence. It may have acquired a new function and expression pattern or may be in the process of becoming a pseudogene. The other gene, Hsp68b, apparently kept its HSE regulatory elements but was relocated to a completely new chromatin environment and is now found near a ~90-kb block composed of Histone genes and TEs. It is difficult to imagine that the expression of this gene has not been affected by these changes. Genes of the heat- inducible Hsp70 family (to which Hsp68 belongs) are positively related to thermotolerance but overexpression has survival costs and it seems that Hsp70 concentration has an intermediate optimum [44, 99]. Some African populations of D. melanogaster with an exceptional thermotolerance show decreased levels of Hsp70 expression, caused by the insertion of TEs in one of the promoter regions of the Hsp70Ba gene [100]. In D. mojavensis an altered expression of Hsp68 genes could contribute to its exceptional thermotolerance. On the other hand, the proximal breakpoint of inversion 2s was located upstream of CG10375, a gene with a DnaJ domain that likely belongs to the constitutively expressed Hsp40 family. In D. melanogaster, hsp40 is up-regulated in mutants lacking HSF [98] and probably has an essential role in thermotolerance [101]. Thus the changes induced by inversion 2s and BuT5 insertion in the promoter of CG10375 likely conferred an adaptive advantage to D. mojavensis by increasing its thermotolerance. It can be hypothesized that the alterations of the heat inducible Hsp68 genes caused by inversion 2r and the putative positive effect on the expression level of the constitutive gene CG10375 caused by inversion 2s were in some way related and jointly contributed to the D. mojavensis unusual thermotolerance. This hypothesis might explain the rapid and exclusive fixation of both inversions in the D. mojavensis lineage.

By no means do we imply that the alterations unveiled at the breakpoints are the only cause of the D. mojavensis inversion adaptive significance. Inversions are not simple point mutations but complex structural changes involving hundreds of loci that may suffer further mutations along their evolutionary trajectory. Therefore we consider that the multiple explanations for the adaptive spread of inversions (see Introduction) are not mutually exclusive alternatives. This means that different inversions may be successful for different reasons but also that a single inversion may increase in frequency for different reasons along its trajectory. For instance, an inversion could gain an initial drive because of the alterations it causes at the breakpoints and incorporate afterwards interacting mutations that led to coadaptation or that increase local adaptation that further propel the inversion towards fixation. The molecular explanations for the role of chromosomal inversions in adaptation and speciation are only beginning to be disentangled.


The breakpoint characterization of seven inversions fixed in D. mojavensis has provided significant information on the causes and consequences of these rearrangements. Multiple generation mechanisms seem to have acted in this lineage, an observation that does not support a mutational explanation for D. mojavensis rapid chromosomal evolution. On the other hand, we have found a set of alterations at the inversion breakpoints with potential adaptive significance, including novel genes and changes in structure and/or expression of adjacent genes. Overall, our results are consistent with natural selection as an explanation for the rapid chromosomal evolution in this specialist organism living under extreme ecological conditions.


In order to map and characterize the breakpoints of D. mojavensis chromosome 2 inversions we used a three-step approach: (1) End sequencing of a set of BAC clones from D. buzzatii chromosome 2; (2) Mapping of the resulting BAC-end sequences (BES) onto the D. mojavensis genome in order to determine the number and chromosomal span of the inversions fixed during the divergence of the two lineages; (3) Identification and annotation of the breakpoint regions using the D. virilis genome as representative of the parental (non-inverted) genome. Chromosome 2 of D. mojavensis differs by 42 chromosomal inversions from the homologous element in D. virilis[6]. The use of the D. buzzatii BES allowed us to identify and characterize those inversions fixed in the D. mojavensis lineage after its divergence from the repleta group ancestor (see Figure 1).

BAC end sequencing

We selected 1,152 clones from the D. buzzatii BAC library homogenously distributed along the 28 contigs of the chromosome 2 physical map [54]. To minimize redundancy we choose overlapping clones but with different restriction patterns. This was done using the information provided by the fingerprinting analysis of BAC clones that is available at The 1,152 clones were rearrayed into 96 well plates (CHORI, Children's Hospital Oakland Research Institute) and both ends of each clone were sequenced (Macrogen Inc., Seoul, Korea) using the universal T7 primer and the modified universal SP6 primer (ATTTAGGTGACACTATAGAAGG) for PCR amplifications at the forward and reverse ends, respectively. We generated 2,127 reads over 400 bp in length, a success rate of 92.32%. Length distribution of BAC-end sequences (BES) for the two primers were similar with a pronounced mode around 700-800 bp (Additional file 10). If only high-quality BES (Q≥20) are taken into account, 80.82% of all sequences had over 400 bp in length. Our goal was to maximize the number of clones with both ends sequenced (paired BES) to increase coverage and the chances to capture all inversion breakpoints. Thus, a total of 1,004 of the original 1,152 BAC clones (87.2%) produced paired BES, whereas 119 clones (10.3%) produced a single BES. All BES were filtered with Geneious® software [102] using VecScreen database in order to identify and remove additional plasmidic sequences.

Mapping D. buzzatii BES onto the D. mojavensis genome

All D. buzzatii BES were tested for similarity to the D. mojavensis genome by BLASTN. This multiple search was carried out with the parameter set '-e' 1e-20, '-W' 7, '-r' 2, '-q' 3, '-G' 5 and '-E' 2 (e-value, word size, reward for a nucleotide match, penalty for a nucleotide mismatch, gap opening cost and gap extension cost, respectively). The values of the rest of the parameters were assigned by default. A masked CAF1 version of D. mojavensis genome, which is available at FlyBase website, was used as reference for these blast searches. The use of a masked genome based on the ReAS library [103] allowed us avoiding results with multiples hits due to repetitive sequences, such as TEs or heterochromatic fragments. Only those hits localized at chromosome 2, which is uniquely represented by scaffold 6540 [55], were considered. Of these, we only took into account the hits that had a minimum length of 50 bp (10% of sequence mean length, approximately). The rest of the hits were discarded, including multiple hits for different scaffolds (except BLAST outputs composed by multiple hits in scaffold 6540 only). All validated hits, i.e., those that met the above criteria, were reordered based on the coordinates of D. mojavensis genome.

From the initial 2,304 BES, 1,933 (83.9%) matched any region of D. mojavensis genome while 1,870 (81.2%) mapped onto chromosome 2 resulting in 2,421 hits (Additional file 10). The number of hits exceeds the number of BES because some BES yielded more than one hit. In most cases the hits produced by a single BES were concatenated, i.e. mapped at adjacent sites in the D. mojavensis genome.

We included in our study a number of BES generated in previous works [56, 59] reaching a total of 2,456 hits. Assuming that chromosome 2 is ~34 Mb long [55], we estimated an average density of one hit or marker every 13.8 kb. The distributions of hit size, e-value and percent identity are shown in (Additional file 10). Hit size was over 400 bp in 50% of all cases, and we did not obtain hits with a length lower than 50 bp due to filtering restrictions. The distribution of e-value was similar for BES from both primers, T7 and SP6, and shows a prominent peak (18.32% of all hits) at an e-value equal to 0 (Additional file 10). Finally, the distribution of percent identity between the D. buzzatii BES and the D. mojavensis genome sequences showed a bell-shaped distribution with an average value of 83.1% (Additional file 10).

A revised version of D. buzzatii physical map of chromosome 2

The published version of D. buzzatii physical map [54] comprises 28 contigs on chromosome 2. Another contig, 1031, has been anchored in chromosome 2 between contigs 1090 and 1181 in a recent mapping work [56]. Here, only four out of the 29 contigs, 1331, 987, 1330 and 1344, were not mapped to chromosome 2 and accordingly are likely to be misassembles or artifacts. We removed them from the revised version. The information provided by the comparative mapping of D. buzzatii BES onto the D. mojavensis genome allows us to assess the presence or absence of overlaps or gaps between contigs and estimate gap size. Supposing that there are no rearrangements or large indels involving contiguous sequences from adjacent contigs, we expect that contigs overlapping in D. mojavensis will also overlap in D. buzzatii, and vice versa. Based on this premise and assuming that D. buzzatii chromosome 2 has a similar size to that of D. mojavensis, we deduce that 15 of the putative gaps between contigs do not exist, i.e. we consider them closed gaps. In addition, we estimated the size of seven gaps between contigs of chromosome 2 as 20-240 kb. Finally, for the remaining two gaps, corresponding to breakpoint regions (see below), we estimated an upper bound for size. In summary, the new version of the map of D. buzzatii chromosome 2 comprises 10 contigs covering ~90% of chromosome 2 and contains 9 gaps that amount to ~5%. The remaining 5% correspond to the endmost (proximal and distal) regions that remain unmapped (see below).

Identification of syntenic segments

Each hit in the D. mojavensis genome was associated with its corresponding clone in the D. buzzatii physical map [54]. In this way, we could infer the number, arrangement and orientation of the conserved segments between D. buzzatii and D. mojavensis. With a single exception (Additional file 1), no syntenic segments were accepted with less than nine hits. Only 77 markers (3%) were not part of any syntenic segment. We guess that these markers represent common elements scattered throughout the genome, such as structural or functional domains or regulatory sequences, or represent gene transposition events. Some BES did not map to any D. mojavensis genome region. This might be caused by incompletely sequenced reads (those which were few bp long), regions with high sequence divergence between D. mojavensis and D. buzzatii or repetitive fragments. Finally, the centromere was not included in any syntenic segment owing to the lack of markers in this region (a masked D. mojavensis genome was used as reference).

Genomic distance

Once established the order and orientation of all syntenic segments in chromosome 2 between D. buzzatii and D. mojavensis, we estimated the genomic distance between the two species using GRIMM software [58]. The genomic distance is the minimum number of chromosomal rearrangements that differentiate two species [104]. The number of rearrangements estimated in this way was the sum of all inversions that had been fixed in the two lineages, D. mojavensi s and D. buzzatii, since their divergence from Primitive I [12]. The three inversions, 2m, 2n and 2z7, fixed in the D. buzzatii lineage have been previously identified [12] and their breakpoints characterized at the molecular level [58]. This allowed us to subtract them from the total and infer the inversions fixed in the other lineage, i.e. from Primitive I to D. mojavensis.

Breakpoint analysis

We identified the breakpoint regions as the D. mojavensis genome sequences located between each pair of adjacent syntenic segments and estimated their size as the distance from the final marker in one syntenic segment to the initial marker in the next syntenic segment. The two breakpoints belonging to the same inversion were associated with the aid of GRIMM results. Once the two breakpoints of each inversion were identified, we proceeded to confirm these results by comparing the breakpoint regions sequences with D. virilis genome using FlyBase GBrowse

D. virilis is the phylogenetically closest species to D. mojavensis whose genome has been sequenced [35]. For this reason it was used as reference for the breakpoint comparative analysis. In order to narrow down the breakpoint regions, we blasted the breakpoint sequences against the D. virilis genome CAF1 masked version, also available at FlyBase website A threshold e-value of 1e-3 was set to take into account the phylogenetic distance between D. virilis and D. mojavensis (Figure 1). All the BLASTN searches were performed with the parameters '-W' 7, '-r' 2, '-q' 3, '-G' 5 and '-E' 2. All hits for each breakpoint sequence were ordered according to D. mojavensis coordinates and the coordinates defined by the similarity loss between D. mojavensis and D. virilis were the new breakpoint limits. A final refinement of the breakpoint regions was carried out comparing the structures of those genes adjacent to the breakpoints in D. mojavensis with their respective orthologs in D. virilis (annotations extracted from FlyFase [105]. If the exon number and gene size were the same or very similar in the two orthologs, coding sequences still present in the D. mojavensis breakpoint regions were excluded from them, confining breakpoints to the intergenic space.

To detect orthologs in the D. virilis genome we downloaded from FlyBase [105] all the nucleotide sequences corresponding to the pair of genes adjacent to each D. mojavensis breakpoint, and then we used them as queries for BLASTN searches against D. virilis genome. We considered as ortholog that gene whose sequence in D. virilis was covered by the most significant hit of that search. To ensure the results, each BLAST search was repeated by exchanging the reference genome with D. mojavensi s using as queries those D. virilis genes sequences putatively identified as orthologs in the first BLAST results (the Reciprocal Best Hit method, [106]). The function of genes adjacent to the breakpoints was inferred from the function of D. melanogaster orthologs in FlyBase and, for those genes without D. melanogaster orthologs, by searching for conserved domains using Interproscan [78] or NCBI Conserved Domain Database [107].

Search of transposable elements

We generated a database with all the D. mojavensis breakpoint sequences. Then we identified all the TEs present at the breakpoint regions by a set of BLASTN searches against DPDB [108], non redundant nucleotide database [109] and RepBase update [110]. We also used RepeatMasker [111] to detect repeats and TEs. Finally, we performed a set of BLAST searches against the breakpoint database using as queries a group of known TEs: Galileo from D. mojavensis (BK006357.1) [112], Newton-1, Newton-2, Kepler-1 and Kepler-5 from D. buzzatii[113], and BuT5 from D. mojavensis [N. Rius, A. Delprat and A. Ruiz, personal communication].

Detection of tandemly arranged duplicated genes

A number of the characterized inversion breakpoints were located between tandemly arranged duplicated genes in the parental (D. virilis genome). In order to test whether this number was expected under a random breakage model, we analyzed all the intergenic regions between duplicated genes in D. mojavensis chromosome 2. We first downloaded a database of predicted proteins for this species available at FlyBase website (version r1.3 of Feb 18, 2010). We extracted from this database all the proteins encoded by genes in scaffold 6540 (chromosome 2) and reordered them based on their gene position on this chromosome. Then, we carried out a search for pairs of similar proteins encoded by adjacent genes using BLAST 2 sequences (bl2seq) [64] with a cutoff e-value of 1e-30. Based on the characteristics of duplicated genes found at breakpoint regions we considered that a pair of proteins was encoded by duplicated genes when the sequence identity between them was over 33% and at least one of the hits was longer than 57% of the shortest query length. Finally we counted the number of intergenic regions located between duplicated genes according to bl2seq results.



We thank Esther Betrán, Andrew Clark, Alejandra Delprat, Bill Etges, Nuria Rius and two anonymous reviewers for useful comments on a previous version of the manuscript. This work was supported by a grant (BFU2008-04988) to AR and a FPI doctoral fellowship (BES-2009-021560) to YG from the Ministerio de Ciencia e Innovación, Spain.

Authors’ Affiliations

Departament de Genètica i de Microbiologia, Universitat Autònoma de Barcelona


  1. White MJD: Animal Cytology and Evolution. 1973, London: Cambridge University Press, 3Google Scholar
  2. Coghlan A, Eichler EE, Oliver SG, Paterson AH, Stein L: Chromosome evolution in eukaryotes: a multi-kingdom perspective. TRENDS in Genetics. 2005, 21: 673-682. 10.1016/j.tig.2005.09.009.PubMedGoogle Scholar
  3. Hoffmann A, Rieseberg LH: Revisiting the Impact of Inversions in Evolution: From Population Genetic Markers to Drivers of Adaptive Shifts and Speciation?. Annual review of ecology, evolution, and systematics. 2008, 39: 21-42. 10.1146/annurev.ecolsys.39.110707.173532.PubMed CentralPubMedGoogle Scholar
  4. Kirkpatrick M: How and why chromosome inversions evolve. PLoS Biology. 2010, 8: e1000501-10.1371/journal.pbio.1000501.PubMed CentralPubMedGoogle Scholar
  5. Murphy WJ, Larkin DM, Everts-van der Wind A, Bourque G, Tesler G, Auvil L, Beever JE, Chowdhary BP, Galibert F, Gatzke L, Hitte C, Meyers SN, Milan D, Ostrander EA, Pape G, Parker HG, Raudsepp T, Rogatcheva MB, Schook LB, Skow LC, Welge M, Womack JE, O'brien SJ, Pevzner PA, Lewin HA: Dynamics of mammalian chromosome evolution inferred from multispecies comparative maps. Science. 2005, 309: 613-617. 10.1126/science.1111387.PubMedGoogle Scholar
  6. Bhutkar A, Schaeffer SW, Russo SM, Xu M, Smith TF, Gelbart WM: Chromosomal rearrangement inferred from comparisons of 12 Drosophila genomes. Genetics. 2008, 179: 1657-1680. 10.1534/genetics.107.086108.PubMed CentralPubMedGoogle Scholar
  7. Ranz JM, Casals F, Ruiz A: How malleable is the eukaryotic genome? Extreme rate of chromosomal rearrangement in the genus Drosophila. Genome Research. 2001, 11: 230-239. 10.1101/gr.162901.PubMed CentralPubMedGoogle Scholar
  8. Grotthuss M von, Ashburner M, Ranz J: Fragile regions and not functional constraints predominate in shaping gene organization in the genus Drosophila. Genome Research. 2010, 20: 1084-1096. 10.1101/gr.103713.109.Google Scholar
  9. Bartolomé C, Charlesworth B: Rates and patterns of chromosomal evolution in Drosophila pseudoobscura and D. miranda. Genetics. 2006, 173: 779-791. 10.1534/genetics.105.054585.PubMed CentralPubMedGoogle Scholar
  10. Papaceit M, Aguadé M, Segarra C: Chromosomal evolution of elements B and C in the Sophophora subgenus of Drosophila: evolutionary rate and polymorphism. Evolution; International Journal of Organic Evolution. 2006, 60: 768-781.PubMedGoogle Scholar
  11. Ranz JM, Maurin D, Chan YS, Grotthuss M von, Hillier LW, Roote J, Ashburner M, Bergman CM: Principles of genome evolution in the Drosophila melanogaster species group. PLoS Biology. 2007, 5: e152-10.1371/journal.pbio.0050152.PubMed CentralPubMedGoogle Scholar
  12. González J, Casals F, Ruiz A: Testing chromosomal phylogenies and inversion breakpoint reuse in Drosophila. Genetics. 2007, 175: 167-177.PubMed CentralPubMedGoogle Scholar
  13. Kupiec M, Petes TD: Allelic and ectopic recombination between Ty elements in yeast. Genetics. 1988, 119: 549-559.PubMed CentralPubMedGoogle Scholar
  14. Lim JK, Simmons MJ: Gross chromosome rearrangements mediated by transposable elements in Drosophila melanogaster. BioEssays: News and Reviews in Molecular, Cellular and Developmental Biology. 1994, 16: 269-275.Google Scholar
  15. Delprat A, Negre B, Puig M, Ruiz A: The transposon Galileo generates natural chromosomal inversions in Drosophila by ectopic recombination. PloS One. 2009, 4: e7883-10.1371/journal.pone.0007883.PubMed CentralPubMedGoogle Scholar
  16. Coulibaly MB, Lobo NF, Fitzpatrick MC, Kern M, Grushko O, Thaner DV, Traoré SF, Collins FH, Besansky NJ: Segmental duplication implicated in the genesis of inversion 2Rj of Anopheles gambiae. PloS One. 2007, 2: e84910-Google Scholar
  17. Cáceres M, Sullivan RT, Thomas JW: A recurrent inversion on the eutherian X chromosome. Proceedings of the National Academy of Sciences of the United States of America. 2007, 104: 18571-18576. 10.1073/pnas.0706604104.PubMed CentralPubMedGoogle Scholar
  18. Richards S, Liu Y, Bettencourt BR, Hradecky P, Letovsky S, Nielsen R, Thornton K, Hubisz MJ, Chen R, Meisel RP, Couronne O, Hua S, Smith MA, Zhang P, Liu J, Bussemaker HJ, Batenburg MF van, Howells SL, Scherer SE, Sodergren E, Matthews BB, Crosby MA, Schroeder AJ, Ortiz-Barrientos D, Rives CM, Metzker ML, Muzny DM, Scott G, Steffen D, Wheeler DA, Worley KC, Havlak P, Durbin KJ, Egan A, Gill R, Hume J, Morgan MB, Miner G, Hamilton C, Huang Y, Waldron L, Verduzco D, Clerc-Blankenburg KP, Dubchak I, Noor MA, Anderson W, White KP, Clark AG, Schaeffer SW, Gelbart W, Weinstock GM, Gibbs RA: Comparative genome sequencing of Drosophila pseudoobscura: chromosomal, gene, and cis-element evolution. Genome Research. 2005, 15: 1-18. 10.1101/gr.3059305.PubMed CentralPubMedGoogle Scholar
  19. Sonoda E, Hochegger H, Saberi A, Taniguchi Y, Takeda S: Differential usage of non-homologous end-joining and homologous recombination in double strand break repair. DNA Repair. 2006, 5: 1021-1029. 10.1016/j.dnarep.2006.05.022.PubMedGoogle Scholar
  20. Furuta Y, Kawai M, Yahara K, Takahashi N, Handa N, Tsuru T, Oshima K, Yoshida M, Azuma T, Hattori M, Uchiyama I, Kobayashi I: Birth and death of genes linked to chromosomal inversion. Proceedings of the National Academy of Sciences of the United States of America. 2011, 108: 1501-1506. 10.1073/pnas.1012579108.PubMed CentralPubMedGoogle Scholar
  21. Cáceres M, Ranz JM, Barbadilla A, Long M, Ruiz A: Generation of a widespread Drosophila inversion by a transposable element. Science. 1999, 285: 415-418. 10.1126/science.285.5426.415.PubMedGoogle Scholar
  22. Casals F, Cáceres M, Ruiz A: The foldback-like transposon Galileo is involved in the generation of two different natural chromosomal inversions of Drosophila buzzatii. Molecular biology and evolution. 2003, 20: 674-685. 10.1093/molbev/msg070.PubMedGoogle Scholar
  23. Evans AL, Mena PA, McAllister BF: Positive selection near an inversion breakpoint on the neo-X chromosome of Drosophila americana. Genetics. 2007, 177: 1303-1319. 10.1534/genetics.107.073932.PubMed CentralPubMedGoogle Scholar
  24. Bergman CM, Pfeiffer BD, Rincón-Limas DE, Hoskins RA, Gnirke A, Mungall CJ, Wang AM, Kronmiller B, Pacleb J, Park S, Stapleton M, Wan K, George RA, de Jong PJ, Botas J, Rubin GMCS: Assessing the impact of comparative genomic sequence data on the functional annotation of the Drosophila genome. Genome Biology. 2002, 3: 1-20.Google Scholar
  25. Prazeres da Costa O, González J, Ruiz A: Cloning and sequencing of the breakpoint regions of inversion 5g fixed in Drosophila buzzatii. Chromosoma. 2009, 118: 349-360. 10.1007/s00412-008-0201-5.PubMedGoogle Scholar
  26. Lande R: The Expected Fixation Rate of Chromosomal Inversions. Evolution. 1984, 38: 743-752. 10.2307/2408386.Google Scholar
  27. Charlesworth B: Fundamental concepts in genetics: effective population size and patterns of molecular evolution and variation. Nature Reviews Genetics. 2009, 10: 195-205.PubMedGoogle Scholar
  28. Navarro A, Betrán E, Barbadilla A, Ruiz A: Recombination and gene flux caused by gene conversion and crossing over in inversion heterokaryotypes. Genetics. 1997, 146: 695-709.PubMed CentralPubMedGoogle Scholar
  29. Dobzhansky TG: Genetics of the evolutionary process. 1970, New York: Columbia Univ PrGoogle Scholar
  30. Kirkpatrick M, Barton N: Chromosome inversions, local adaptation and speciation. Genetics. 2006, 173: 419-434. 10.1534/genetics.105.047985.PubMed CentralPubMedGoogle Scholar
  31. Sperlich D, Pfreim P: Cromosomal polymorphism in natural and experimental populations. Edited by: Ashburner M, Carson HL. 1986, Thompson JNJ London, 3: 257-309.Google Scholar
  32. Puig M, Cáceres M, Ruiz A: Silencing of a gene adjacent to the breakpoint of a widespread Drosophila inversion by a transposon-induced antisense RNA. Proceedings of the National Academy of Sciences of the United States of America. 2004, 101: 9013-9018. 10.1073/pnas.0403090101.PubMed CentralPubMedGoogle Scholar
  33. Celniker SERG: The Drosophila melanogaster Genome. Annual Review of Genomics and Human Genetics. 2003, 4: 89-117. 10.1146/annurev.genom.4.070802.110323.PubMedGoogle Scholar
  34. modENCODE Consortium: Identification of functional elements and regulatory circuits by Drosophila modENCODE. Science. 2010, 330: 1787-1797.Google Scholar
  35. Drosophila 12 Genomes Consortium: Evolution of genes and genomes on the Drosophila phylogeny. Nature. 2007, 450: 203-218. 10.1038/nature06341.Google Scholar
  36. Barker JSF: Population genetics of Opuntia breeding Drosophila in Australia. Ecological genetics and evolution. 1982, Academic Press, Australia, 209-224.Google Scholar
  37. Barker J, Starmer W, MacIntyre R: Ecological and evolutionary genetics of Drosophila. 1990, New York: Plenum Publishing CorporationGoogle Scholar
  38. Markow T, O'Grady PM: Drosophila: a guide to species identification and use. 2005, Academic PressGoogle Scholar
  39. Markow TA, O'Grady PM: Drosophila biology in the genomic age. Genetics. 2007, 177: 1269-1276. 10.1534/genetics.107.074112.PubMed CentralPubMedGoogle Scholar
  40. Fellows DP, Heed WB: Factors affecting host plant selection in desert adapted cactophilic Drosophila. Ecology. 1972, 53: 850-858. 10.2307/1934300.Google Scholar
  41. Heed WB, Mangan RL: Community ecology of the Sonoran Desert Drosophila. Edited by: Ashburner M, Carson HL, Thompson JNJ. 1986, New York: Academic Press, 3e: 311-345.Google Scholar
  42. Ruiz A, Heed W: Host-Plant Specificity in the Cactophilic Drosophila mulleri Species Complex. Journal of Animal Ecology. 1988, 57: 237-249. 10.2307/4775.Google Scholar
  43. McKnight T, Hess D: Climate Zones and Types: The Köppen System. 2000, Upper Saddle River, NJ: Prentice Hall, 200-1.Google Scholar
  44. Krebs RA: A comparison of Hsp70 expression and thermotolerance in adults and larvae of three Drosophila species. Cell Stress & Chaperones. 1999, 4: 243-249. 10.1379/1466-1268(1999)004<0243:ACOHEA>2.3.CO;2.Google Scholar
  45. Stratman R, Markow TA: Resistance to thermal stress in desert Drosophila. Functional Ecology. 1998, 12: 965-970. 10.1046/j.1365-2435.1998.00270.x.Google Scholar
  46. Gibbs AG, Fukuzato F, Matzkin LM: Evolution of water conservation mechanisms in Drosophila. Journal of experimental biology. 2003, 206: 1183-1192. 10.1242/jeb.00233.PubMedGoogle Scholar
  47. Matzkin LM, Markow T: Transcriptional regulation of metabolism associated with the increased desiccation resistance of the cactophilic Drosophila mojavensis. Genetics. 2009, 182: 1279-1288. 10.1534/genetics.109.104927.PubMed CentralPubMedGoogle Scholar
  48. Kircher HW: Chemical composition of cacti and its relationship to sonoran desert Drosophila. Edited by: Barker JSF, Starmer WT. 1982, New York: Academic Press, 143-158.Google Scholar
  49. Fogleman JC, Danielson PB: Chemical interactions in the cactus-microorganism-Drosophila model system of the Sonoran Desert. American Zoologist. 2001, 41: 877-889. 10.1668/0003-1569(2001)041[0877:CIITCM]2.0.CO;2.Google Scholar
  50. Wasserman M: Cytological studies of the repleta group of the genus Drosophila. V. The mulleri subgroup. Univ Texas Publ. 1962, 6205: 85-118.Google Scholar
  51. Ruiz A, Heed WB, Wasserman M: Evolution of the mojavensis cluster of cactophilic Drosophila with descriptions of two new species. The Journal of Heredity. 1990, 81: 30-42.PubMedGoogle Scholar
  52. Wasserman M: Cytological evolution of the Drosophila repleta species group. Edited by: Powell JR, Krimbas CB. 1992, Boca Raton, Florida: CRC Press, 455-541.Google Scholar
  53. Runcie DE, Noor MAF: Sequence signatures of a recent chromsomal rearrangement in Drosophila mojavensis. Genetica. 2009, 136: 5-11. 10.1007/s10709-008-9296-0.PubMed CentralPubMedGoogle Scholar
  54. González J, Nefedov M, Bosdet I, Casals F, Calvete O, Delprat A, Shin H, Chiu R, Mathewson C, Wye N, Hoskins R, Schein J, de Jong P, Ruiz A: A BAC-based physical map of the Drosophila buzzatii genome. Genome Research. 2005, 15: 885-892. 10.1101/gr.3263105.PubMed CentralPubMedGoogle Scholar
  55. Schaeffer SW, Bhutkar A, McAllister BF, Matsuda M, Matzkin LM, O'Grady PM, Rohde C, Valente VLS, Aguadé M, Anderson WW, Edwards K, Garcia ACL, Goodman J, Hartigan J, Kataoka E, Lapoint RT, Lozovsky ER, Machado CA, Noor MAF, Papaceit M, Reed LK, Richards S, Rieger TT, Russo SM, Sato H, Segarra C, Smith DR, Smith TF, Strelets V, Tobari YN, Tomimura Y, Wasserman M, Watts T, Wilson R, Yoshida K, Markow TA, Gelbart WM, Kaufman TC: Polytene chromosomal maps of 11 Drosophila species: the order of genomic scaffolds inferred from genetic and physical maps. Genetics. 2008, 179: 1601-1655. 10.1534/genetics.107.086074.PubMed CentralPubMedGoogle Scholar
  56. Prada C: Evolución cromosómica del cluster Drosophila martensis: origen de las inversiones y reuso de puntos de rotura. PhD thesis. 2010, Universitat Autònoma de BarcelonaGoogle Scholar
  57. Pevzner P, Tesler G: Genome rearrangements in mammalian evolution: lessons from human and mouse genomes. Genome Research. 2003, 13: 37-45. 10.1101/gr.757503.PubMed CentralPubMedGoogle Scholar
  58. Tesler G: GRIMM: genome rearrangements web server. Bioinformatics (Oxford, England). 2002, 18: 492-493. 10.1093/bioinformatics/18.3.492.Google Scholar
  59. Calvete Torres O: Dinámica evolutiva de las reordenaciones cromosómicas y coincidencia de los puntos de rotura: Análisis molecular de las inversiones fijadas en el cromosoma 2 de Drosophila buzzatii. PhD thesis. 2010, Universitat Autònoma de BarcelonaGoogle Scholar
  60. Froenicke L, Caldés MG, Graphodatsky A, Müller S, Lyons LA, Robinson TJ, Volleth M, Yang F, Wienberg J: Are molecular cytogenetics and bioinformatics suggesting diverging models of ancestral mammalian genomes?. Genome Research. 2006, 16: 306-310. 10.1101/gr.3955206.PubMed CentralPubMedGoogle Scholar
  61. Bourque G, Tesler G, Pevzner PA: The convergence of cytogenetics and rearrangement-based models for ancestral genome reconstruction. Genome research. 2006, 16: 311-313. 10.1101/gr.4631806.PubMed CentralPubMedGoogle Scholar
  62. Ranz J, Gonzalez J, Casals F, Ruiz A: Low occurrence of gene transposition events during the evolution of the genus Drosophila. Evolution; International Journal of Organic Evolution. 2003, 57: 1325-1335.PubMedGoogle Scholar
  63. Prada C, Delprat A, Ruiz A: Testing chromosomal phylogenies and inversion breakpoint reuse in Drosophila. The martensis cluster revisited. Chromosome Research. 2011, 19: 251-265. 10.1007/s10577-011-9195-6.PubMedGoogle Scholar
  64. Tatusova TA, Madden TL: BLAST 2 Sequences, a new tool for comparing protein and nucleotide sequences. FEMS Microbiology Letters. 1999, 174: 247-250. 10.1111/j.1574-6968.1999.tb13575.x.PubMedGoogle Scholar
  65. Cáceres M, Puig M, Ruiz A: Molecular characterization of two natural hotspots in the Drosophila buzzatii genome induced by transposon insertions. Genome research. 2001, 11: 1353-1364. 10.1101/gr.174001.PubMed CentralPubMedGoogle Scholar
  66. Arca B, Zabalou S, Loukeris TG, Savakis C: Mobilization of a Minos transposon in Drosophila melanogaster chromosomes and chromatid repair by heteroduplex formation. Genetics. 1997, 145: 267-279.PubMed CentralPubMedGoogle Scholar
  67. Beall EL, Rio DC: Drosophila P-element transposase is a novel site-specific endonuclease. Genes & Development. 1997, 11: 2137-2151. 10.1101/gad.11.16.2137.Google Scholar
  68. Tamura K, Subramanian S, Kumar S: Temporal patterns of fruit fly (Drosophila) evolution revealed by mutation clocks. Molecular Biology and Evolution. 2004, 21: 36-44.PubMedGoogle Scholar
  69. Dehal P, Predki P, Olsen AS, Kobayashi A, Folta P, Lucas S, Land M, Terry A, Ecale Zhou CL, Rash S, Zhang Q, Gordon L, Kim J, Elkin C, Pollard MJ, Richardson P, Rokhsar D, Uberbacher E, Hawkins T, Branscomb E, Stubbs L: Human chromosome 19 and related regions in mouse: conservative and lineage-specific evolution. Science. 2001, 293: 104-111. 10.1126/science.1060310.PubMedGoogle Scholar
  70. Carbone L, Harris RA, Vessere GM, Mootnick AR, Humphray S, Rogers J, Kim SK, Wall JD, Martin D, Jurka J, Milosavljevic A, Jong PJ de: Evolutionary breakpoints in the gibbon suggest association between cytosine methylation and karyotype evolution. PLoS Genetics. 2009, 5: e1000538-10.1371/journal.pgen.1000538.PubMed CentralPubMedGoogle Scholar
  71. Voineagu I, Narayanan V, Lobachev KS, Mirkin SM: Replication stalling at unstable inverted repeats: Interplay between DNA hairpins and fork stabilizing proteins. 2008, 105: 9936-9941.Google Scholar
  72. Bai Y, Casola C, Feschotte C, Betrán E: Comparative genomics reveals a constant rate of origination and convergent acquisition of functional retrogenes in Drosophila. Genome Biology. 2007, 8: R11-10.1186/gb-2007-8-1-r11.PubMed CentralPubMedGoogle Scholar
  73. Bhutkar A, Russo SM, Smith TF, Gelbart WM: Genome-scale analysis of positionally relocated genes. Genome Research. 2007, 17: 1880-1887. 10.1101/gr.7062307.PubMed CentralPubMedGoogle Scholar
  74. Vibranovski MD, Zhang Y, Long M: General gene movement off the X chromosome in the Drosophila genus. Genome Research. 2009, 19: 897-903. 10.1101/gr.088609.108.PubMed CentralPubMedGoogle Scholar
  75. Lifton RP, Goldberg ML, Karp RW, Hogness DS: The organization of the histone genes in Drosophila melanogaster: functional and evolutionary implications. Cold Spring Harbor Symposia on Quantitative Biology. 1978, 1047-1051. 42 Pt 2
  76. Kremer H, Hennig W: Isolation and characterization of a Drosophila hydei histone DNA repeat unit. Nucleic Acids Research. 1990, 18: 1573-1580. 10.1093/nar/18.6.1573.PubMed CentralPubMedGoogle Scholar
  77. Cohen S, Agmon N, Yacobi K, Mislovati M, Segal D: Evidence for rolling circle replication of tandem genes in Drosophila. Nucleic acids research. 2005, 33: 4519-4526. 10.1093/nar/gki764.PubMed CentralPubMedGoogle Scholar
  78. Zdobnov EM, Apweiler R: InterProScan--an integration platform for the signature-recognition methods in InterPro. Bioinformatics (Oxford, England). 2001, 17: 847-848. 10.1093/bioinformatics/17.9.847.Google Scholar
  79. Marger MD, Saier MH: A major superfamily of transmembrane facilitators that catalyse uniport, symport and antiport. Trends in Biochemical Sciences. 1993, 18: 13-20. 10.1016/0968-0004(93)90081-W.PubMedGoogle Scholar
  80. Salinas AE, Wong MG: Glutathione S-transferases--a review. Current Medicinal Chemistry. 1999, 6: 279-309.PubMedGoogle Scholar
  81. Low WY, Ng HL, Morton CJ, Parker MW, Batterham P, Robin C: Molecular evolution of glutathione S-transferases in the genus Drosophila. Genetics. 2007, 177: 1363-1375. 10.1534/genetics.107.075838.PubMed CentralPubMedGoogle Scholar
  82. Matzkin LM: The molecular basis of host adaptation in cactophilic Drosophila: molecular evolution of a glutathione S-transferase gene (GstD1) in Drosophila mojavensis. Genetics. 2008, 178: 1073-1083. 10.1534/genetics.107.083287.PubMed CentralPubMedGoogle Scholar
  83. Matzkin LM, Watts TD, Bitler BG, Machado CA, Markow T: Functional genomics of cactus host shifts in Drosophila mojavensis. Molecular Ecology. 2006, 15: 4635-4643. 10.1111/j.1365-294X.2006.03102.x.PubMedGoogle Scholar
  84. Bettencourt BR, Feder ME: Rapid concerted evolution via gene conversion at the Drosophila hsp70 genes. Journal of Molecular Evolution. 2002, 54: 569-586. 10.1007/s00239-001-0044-7.PubMedGoogle Scholar
  85. Tian S, Haney RA, Feder ME: Phylogeny Disambiguates the Evolution of Heat-Shock cis-Regulatory Elements in Drosophila. PLoS One. 2010, 5: e10669-10.1371/journal.pone.0010669.PubMed CentralPubMedGoogle Scholar
  86. Zhang Z, Pugh BF: Genomic Organization of H2Av Containing Nucleosomes in Drosophila Heterochromatin. PloS One. 2011, 6: e20511-10.1371/journal.pone.0020511.PubMed CentralPubMedGoogle Scholar
  87. Bell O, Tiwari VK, Thomä NH, Schübeler D: Determinants and dynamics of genome accessibility. Nature Reviews Genetics. 2011, 12: 554-564. 10.1038/nrg3017.PubMedGoogle Scholar
  88. Li G, Reinberg D: Chromatin higher-order structures and gene regulation. Current Opinion in Genetics & Development. 2011, 21: 175-186. 10.1016/j.gde.2011.01.022.Google Scholar
  89. Ohler U: Identification of core promoter modules in Drosophila and their application in accurate transcription start site prediction. Nucleic Acids Research. 2006, 34: 5943-5950. 10.1093/nar/gkl608.PubMed CentralPubMedGoogle Scholar
  90. Petrov DA, Lozovskaya ER, Hartl DL: High intrinsic rate of DNA loss in Drosophila. Nature. 1996, 384: 346-349. 10.1038/384346a0.PubMedGoogle Scholar
  91. Petrov DA, Hartl DL: High rate of DNA loss in the Drosophila melanogaster and Drosophila virilis species groups. Molecular Biology and Evolution. 1998, 15: 293-302.PubMedGoogle Scholar
  92. Matzkin LM, Eanes WF: Sequence variation of alcohol dehydrogenase (Adh) paralogs in cactophilic Drosophila. Genetics. 2003, 163: 181-194.PubMed CentralPubMedGoogle Scholar
  93. Matzkin LM: Population genetics and geographic variation of alcohol dehydrogenase (Adh) paralogs and glucose-6-phosphate dehydrogenase (G6pd) in Drosophila mojavensis. Molecular Biology and Evolution. 2004, 21: 276-285.PubMedGoogle Scholar
  94. Kaessmann H: Origins, evolution, and phenotypic impact of new genes. Genome Research. 2010, 20: 1313-1326. 10.1101/gr.101386.109.PubMed CentralPubMedGoogle Scholar
  95. Hoffmann A, Sørensen JG, Loeschcke V: Adaptation of Drosophila to temperature extremes: bringing together quantitative and molecular approaches. Journal of Thermal Biology. 2003, 28: 175-216. 10.1016/S0306-4565(02)00057-8.Google Scholar
  96. McColl G, Hoffmann A, McKechnie SW: Response of two heat shock genes to selection for knockdown heat resistance in Drosophila melanogaster. Genetics. 1996, 143: 1615-1627.PubMed CentralPubMedGoogle Scholar
  97. Parker CS, Topol J: A Drosophila RNA polymerase II transcription factor binds to the regulatory site of an hsp 70 gene. Cell. 1984, 37: 273-283. 10.1016/0092-8674(84)90323-4.PubMedGoogle Scholar
  98. Neal SJ, Karunanithi S, Best A, So AK-C, Tanguay RM, Atwood HL, Westwood JT: Thermoprotection of synaptic transmission in a Drosophila heat shock factor mutant is accompanied by increased expression of Hsp83 and DnaJ-1. Physiological Genomics. 2006, 25: 493-501. 10.1152/physiolgenomics.00195.2005.PubMedGoogle Scholar
  99. Krebs RA, Feder ME: Hsp70 and larval thermotolerance in Drosophila melanogaster: how much is enough and when is more too much?. Journal of Insect Physiology. 1998, 44: 1091-1101. 10.1016/S0022-1910(98)00059-6.PubMedGoogle Scholar
  100. Lerman DN, Feder ME: Naturally occurring transposable elements disrupt hsp70 promoter function in Drosophila melanogaster. Molecular Biology and Evolution. 2005, 22: 776-783.PubMedGoogle Scholar
  101. Carmel J, Rashkovetsky E, Nevo E, Korol A: Differential Expression of Small Heat Shock Protein Genes in Fruit Flies (Drosophila melanogaster) along a Microclimatic Gradient. Journal of Heredity. 2011, 10.1093/jhered/esr027Google Scholar
  102. Drummond A, Ashton B, Cheung M, Cooper A, Heled J, Kearse M, Moir R, Stones-Havas S, Sturrock S, Thierer TWA: Geneious v5.1. 2010, Available from http://www.geneious.comGoogle Scholar
  103. Li , Ye J, Li S, Wang J, Han Y, Ye C, Wang J, Yang H, Yu J, Wong GK-S, Wang J: ReAS: Recovery of ancestral sequences for transposable elements from the unassembled reads of a whole genome shotgun. PLoS Computational Biology. 2005, 1: e43-10.1371/journal.pcbi.0010043.PubMed CentralPubMedGoogle Scholar
  104. Hannenhalli S, Pevzner P: Transforming men into mice (polynomial algorithm for genomic distance problem). Proceedings of the 36th Annual IEEE Symposium on Foundations of Computer Science. 1995, 581-592.Google Scholar
  105. Drysdale R: FlyBase: a database for the Drosophila research community. Methods in Molecular Biology. 2008, 420: 45-59. 10.1007/978-1-59745-583-1_3.PubMedGoogle Scholar
  106. Moreno-Hagelsieb G, Latimer K: Choosing BLAST options for better detection of orthologs as reciprocal best hits. Bioinformatics. 2008, 24: 319-324. 10.1093/bioinformatics/btm585.PubMedGoogle Scholar
  107. Marchler-Bauer A, Lu S, Anderson JB, Chitsaz F, Derbyshire MK, DeWeese-Scott C, Fong JH, Geer LY, Geer RC, Gonzales NR, Gwadz M, Hurwitz DI, Jackson JD, Ke Z, Lanczycki CJ, Lu F, Marchler GH, Mullokandov M, Omelchenko MV, Robertson CL, Song JS, Thanki N, Yamashita RA, Zhang D, Zhang N, Zheng C, Bryant SH: CDD: a Conserved Domain Database for the functional annotation of proteins. Nucleic Acids Research. 2011, 39: D225-229. 10.1093/nar/gkq1189.PubMed CentralPubMedGoogle Scholar
  108. Casillas S, Egea R, Petit N, Bergman CM, Barbadilla A: Drosophila polymorphism database (DPDB): a portal for nucleotide polymorphism in Drosophila. Fly. 2007, 1: 205-211.PubMedGoogle Scholar
  109. Benson DA, Karsch-Mizrachi I, Lipman DJ, Ostell J, Sayers EW: GenBank. Nucleic Acids Research. 2011, 39: D32-37. 10.1093/nar/gkq1079.PubMed CentralPubMedGoogle Scholar
  110. Jurka J, Kapitonov VV, Pavlicek A, Klonowski P, Kohany O, Walichiewicz J: Repbase Update, a database of eukaryotic repetitive elements. Cytogenetic and Genome Research. 2005, 110: 462-467. 10.1159/000084979.PubMedGoogle Scholar
  111. Chen N: Using RepeatMasker to identify repetitive elements in genomic sequences. Current Protocols in Bioinformatics. 2004, Chapter 4: Unit 4.10Google Scholar
  112. Marzo M, Puig M, Ruiz A: The Foldback-like element Galileo belongs to the P superfamily of DNA transposons and is widespread within the Drosophila genus. Proceedings of the National Academy of Sciences of the United States of America. 2008, 105: 2957-2962. 10.1073/pnas.0712110105.PubMed CentralPubMedGoogle Scholar
  113. Casals F, Cáceres M, Manfrin MH, González J, Ruiz A: Molecular characterization and chromosomal distribution of Galileo, Kepler and Newton, three foldback transposable elements of the Drosophila buzzatii species complex. Genetics. 2005, 169: 2047-2059. 10.1534/genetics.104.035048.PubMed CentralPubMedGoogle Scholar
  114. Tamura K, Dudley J, Nei M, Kumar S: MEGA4: Molecular Evolutionary Genetics Analysis (MEGA) software version 4.0. Molecular Biology and Evolution. 2007, 24: 1596-1599. 10.1093/molbev/msm092.PubMedGoogle Scholar


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