extfieldsofstudy: Biology Medicine

text:
High tandem repeat content in the genome of the short-lived annual fish Nothobranchius furzeri: a new vertebrate model for aging research

A genomic analysis of the annual fish Nothobranchius furzeri, a vertebrate with the shortest known life span in captivity and which may provide a new model organism for aging research.

Background
Studies in invertebrate model organisms such as yeast and worm have identified a number of genes and pathways that regulate life span and aging [1], and some of these are conserved across taxa [2][3][4][5]. While this work has been crucial to elucidate common aging related pathways like insulin/insulin-like growth factor signaling [6][7][8], in many cases experimentally proven relevance for invertebrate genes/gene products cannot be reproduced in vertebrates. Also, the comparatively long life span of vertebrate model organisms poses a difficulty for testing of findings initially obtained in invertebrates -for example, a life expectancy of several years like that of mouse, rat, or zebrafish renders experimental analyses of potentially life extending drugs difficult.
The Turquoise killifish, Nothobranchius furzeri, might represent an alternative vertebrate model for the study of aging [9]. The fish inhabit seasonal ponds in South-East Africa and were first captured in 1968 in the Game Reserve Gona Re Zhou (GRZ) in Zimbabwe [10]. In 1975, only two pairs of the direct descendants of these fish were left. One of these pairs was then used for breeding, mainly to preserve the species for the killifish community [11]. Offspring of these fish have since been maintained by dedicated hobbyists and in the following are referred to as the GRZ strain. The maximum life span of GRZ fish was previously found to be 12-13 weeks in captivity [12][13][14]. In our facility, GRZ fish exhibit a maximum life span of 16 weeks [15]. The fish show explosive growth and early sexual maturation, and with advancing age typical aging related features such as a decline in learning/behavioral capabilities as well as expression of aging biomarkers are observed [16,17]. Furthermore, GRZ fish are susceptible to life span modulation. Maximum life span is significantly prolonged by moderately decreased water temperature and treatment with resveratrol, both of which are characterized by delayed onset of cognitive decline and expression of aging biomarkers [13,14]. In contrast to the GRZ strain established from the game reserve in Zimbabwe, recently established isolates of N. furzeri populations from southern Mozambique differ in life span and time of expression of age-related traits, which presumably reflects adaptation to the seasonal duration of their respective ponds [18]. The maximum life span of these recent isolates is 25-32 weeks. Although this is twice as long as found for the GRZ strain, it is still exceptionally short compared to other vertebrates. It also does not seem to change considerably in captivity -for example, maximum life span is 32 weeks in the first captive generation of a southern N. furzeri population collected in 2004 and remains at 31 weeks in the sixth captive generation of the derived strain [15], which is currently bred in our facility and was termed N. furzeri MZM-0403 [18]. In addition, there are several other African Nothobranchius species, which live longer than N. furzeri, including N. kunthae (37 weeks) and N. guentheri (52 weeks, reviewed in [9]), opening up the possibility to study naturally occurring aging phenotypes both in N. furzeri and across Nothobranchius species.
To date very little is known about the genetic makeup of N. furzeri and at present one cannot exclude that private mutations or inbreeding cause the exceptionally short life span of the GRZ strain. However, the presence of aging related biomarkers and changes in their expression in response to external stimuli [14,16], as well as a similarly short life span and visible symptoms of old age reported for GRZ fish 33 years ago [11], argue in favor of the action of common mechanisms of life span determination in this strain.
In order to support establishing N. furzeri as a model organism for age research, we provide here a first insight into its cytogenetic and genomic characteristics, including karyotype, genome size/composition, phylogenetic positioning and genetic variability. We compare its genomic features to those of medaka (Oryzias latipes), stickleback (Gasterosteus aculeatus), tetraodon (Tetraodon nigroviridis) and zebrafish (Danio rerio). These species serve as models in many areas of contemporary research, such as genome evolution (tetraodon), developmental biology and genetics (medaka, zebrafish), and speciation (stickleback). Genome projects are completed or well underway; whole genome analyses have been published for tetraodon and medaka [19,20], and preliminary genome assemblies have been provided for stickleback and zebrafish [21]. Clearly, the sequences facilitate large scale and systematic studies [22][23][24].
Our work is a first step towards a systematic identification of genes and biochemical pathways involved in life span determination in N. furzeri. Combined with the genetic resources for the aforementioned fish species it also forms a basis to make full use of N. furzeri as a model organism for the study of aging.

Cytogenetic characteristics
The chromosome number of N. furzeri is 2n = 38 and includes four pairs of metacentric, three pairs of acrocentric and twelve pairs of subtelo-/submetacentric chromosomes ( Figure 1a). Based on morphology, there do not seem to be clearly differentiated sex chromosomes. Specific heterochromatin staining indicates that the cytological organization of the N. furzeri genome is highly structured as is evident from the presence of large blocks of C-banding positive heterochromatin in the centromeric region of most chromosomes (Figure 1b). Accumulation of heterochromatin in other chromosomal sites cannot be detected. To evaluate the composition of heterochromatin, we performed base specific fluorochrome staining on metaphase chromosomes. Staining with the A+T specific dye DAPI resulted in poor fluorescence of centromeric regions (Figures 1a and 2a). Conversely, staining with mithramycin, which shows high affinity for G+C-rich DNA, generated bright fluorescence in most centromeric regions that are DAPI dull (Figure 1c). This indicates that constitutive heterochromatin in N. furzeri is G+C-rich. The anal-ysis of closely related African Nothobranchii, including the sympatric species N. orthonotus as well as two allopatric species from Tanzania, N. hengstleri and N. eggersi, does not indicate a comparably structured genome organization (Figure 2b-d). Thus, at the cytological level, a compartmentalization of the N. furzeri genome is apparent which is likely caused by substantial differences in DNA composition. This seems unusual since fish genomes are generally characterized by a very limited compositional heterogeneity [25,26].

Genome size
We sequenced 5.4 Mb of the short-lived strain N. furzeri GRZ, the long-lived, recently wild-derived strain N. furzeri MZM-0403, and the long-lived closely related species N. kunthae using a whole genome shotgun approach and Sanger sequencing (Table 1). To assess the genome size of N. furzeri based on the sequences, we assumed that the number of its protein coding genes does not differ significantly from that of other fish species, as was previously suggested for vertebrates [27]. We identified sequences containing protein coding information in the 5.4 Mb of both N. furzeri strains by BLASTX searches in Swiss-Prot/TrEMBL, and did the same for medaka, stickleback, tetraodon and zebrafish, for which we extracted adequate genomic samples from public databases. Based on the reported genome sizes of the latter four fish species, we then deduced the genome size of N. furzeri. In detail, 444 (8%) of the 5,540 N. furzeri GRZ sequences and 443 (8%) of the 5,686 MZM-0403 sequences show significant similarity (p < 10 -10 ) to protein coding genes. Respective sequences comprise 31%, 20%, 13% and 11% of the tetraodon, stickleback, medaka and zebrafish genomic samples, respectively, which corresponds to a genome size of 1.59-1.92 Gb for N. furzeri (Additional data files 1 and 2). For experimental confirmation, we performed flow cytometry measurements using DAPI and propidium iodide (PI). DAPI, which preferentially stains A+T rich DNA segments, yields a DNA content of 2.33 pg/diploid cell while the non-base-specific PI staining results in 3.11 pg/diploid cell (Additional data file 3). In light of the large blocks of G+C-rich heterochromatin observed in our cytogenetic studies, the value obtained with PI is likely the correct one since it is based on a dye that does not depend on DNA composition, whereas DAPI staining most probably results in an underestimation of DNA content [28]. The value obtained with PI corresponds well with our sequence based Cytogenetic features of N. furzeri Figure 1 Cytogenetic features of N. furzeri. (a) Karyotype of DAPI-stained chromosomes of a female N. furzeri of the GRZ strain. Note the absence of bright staining at the centromeric regions. Four pairs of chromosomes (1,6,9,17) are metacentric, three pairs (16,18,19) are acrocentric and the remaining 12 pairs are subtelo-/submetacentric. (b) C-banded karyotype of a female N. furzeri GRZ reveals centromeric heterochromatin in most chromosomes. (c) Mithramycin staining results in bright fluorescence of centromeres, which is due to G+C enriched heterochromatin. estimate; that is, it is equal to a genome size of approximately 1.5 Gb. The 5.4 Mb thus represents roughly 0.3-0.5% of the N. furzeri genome.

Genome composition
The G+C content of the 5.4 Mb sample of N. furzeri GRZ is 44.9%. Interestingly, approximately 10% of the sequences have a G+C content of 60% or higher; in Figure 3a this is indicated by a second peak at approximately 62% in a plot of number of sequences against G+C content. The same unusual G+C content distribution is seen in the recently wild-derived N. furzeri strain MZM-0403 (Additional data file 4). To test whether this represents an artifact introduced by preferential propagation of G+C-rich sequences in Escherichia coli [32] during library preparation, we performed whole genome Chromosomes of four African Nothobranchius species Figure 2 Chromosomes of four African Nothobranchius species. DAPI stained chromosomes of (a) a female specimen of the N. furzeri GRZ strain, (b) a male specimen of the sympatric species N. orthonotus, and (c) a male specimen of the allopatric species N. hengstleri and (d) N. eggersi, respectively. Note the dull DAPI fluorescence at centromeric regions in N. furzeri chromosomes (indicated by arrowheads), which is indicative of the presence of G+C-rich constitutive heterochromatin and not observable in the three closely related Nothobranchius species.

(a) (b) (d) (c)
shotgun sequencing using Roche/454 Life Sciences technology, which does not involve cloning and amplification in bacterial systems [33], for N. furzeri GRZ. In this second GRZ genomic sequence sample, the G+C content is 44.3% (111.6 Mb; Table 1). Similar to the Sanger sequences, approximately 10% of the 454 sequences show a G+C content of at least 60%, which indicates that there is not a strong cloning bias in the genomic libraries we initially sequenced. Taken together, our sequence data suggest that a distinct G+C-rich fraction is present in the N. furzeri genome. A similar fraction does not seem to exist in medaka, stickleback, tetraodon and zebrafish ( Figure 3b) and is absent as well in the closely related species N. kunthae (Additional data file 4).
To assess the extent to which our sample-based estimation reflects the G+C content of the N. furzeri genome, we analyzed entire genomes as well as adequate genomic samples of medaka, stickleback, tetraodon and zebrafish. We found that the G+C content estimates of genomes and genomic samples are essentially the same ( Table 2). Our calculations are in agreement with previously published data; we estimated a G+C content of 46.4% for tetraodon and reported values are 45.5% [31] and 46.4% [20]; similarly, 40.3% was reported for medaka [20] and we found it to be 40.5%. Thus, the G+C content of N. furzeri is likely 44-45%, which is similar to stickleback (44.6%), slightly lower than in tetraodon (46.4%) and considerably higher than in medaka (40.5%) and zebrafish (36.6%). Based on genomic sequences, an inverse correlation of genome size and G+C content was recently found for the latter four fish species [34], which confirmed previous experimental results showing that small genomes are generally associated with high G+C content and vice versa [25]. N.
furzeri would seem an exception as its G+C content is nearly as high as that of tetraodon, which has a four times smaller genome. The sequence fraction in the N. furzeri genome with the high G+C content (≥60%) increases the global G+C content rather slightly -the value is 43.1% if these sequences are excluded -and it seems that these sequences occupy defined chromosomal regions (see below).
While to our knowledge there is no evidence for a direct influence of nuclear genome composition on the life span of a vertebrate species, some reports correlate life span with the composition of proteins encoded by the mitochondrial genome. For example, Rottenberg [35] found that rates of amino acid substitution per site in mitochondrial DNA of mammals are positively correlated with longevity of a genus and suggested that the evolution of longevity drove the accelerated evolution of peptides encoded by mitochondrial DNA. Moosmann and Behl [36] showed that the frequency with which cysteine is encoded by mitochondrial DNA is a specific indicator for longevity; that is, longevity is associated with a depletion of mitochondrial cysteine in aerobic species. It will be very interesting to analyze the cysteine content in mitochondrially encoded proteins in comparison to nuclear proteins in N. furzeri strains/Nothobranchius species with different life spans.

Repeats
Repetitive elements can be grouped into the main classes 'tandem repeats', 'transposon-derived interspersed repeats', 'processed pseudogenes' and 'segmental duplications'. Because our genomic samples comprise rather small, fragmented fractions of the N. furzeri genome, it is impossible to  Table  1) practically rules out a meaningful repeat analysis. We therefore concentrated on the mere identification of tandem repeats and transposon-derived interspersed repeats in the 5.4 Mb of both N. furzeri strains and N. kunthae generated by Sanger sequencing and, for comparison, analyzed our samples of medaka, stickleback, tetraodon and zebrafish genomes.
We considered as tandem repeats microsatellites, minisatellites and satellites composed of 1-5 nucleotides, 6-99 nucleotides, and over 100 nucleotides per repeat unit, respectively. About 1% of the N. furzeri DNA is composed of microsatellites, which is comparable with tetraodon (1.1%) and stickleback (0.8%), about half as much as in zebrafish (2%) and five times more than in medaka (0.2%) ( Table 2). Roest Crollius et al. [31] reported that microsatellites comprise 3.21% of the tetraodon genome. The higher number is most probably due to the different algorithms and motif sizes applied; that is, Roest Crollius et al. used a Smith and Waterman algorithmbased approach previously applied to Takifugu rubripes [37] and a motif size of 1-6 nucleotides, while we used the program Sputnik [38], a specific tool for the detection of microsatellites, and a motif size of 1-5 nucleotides.
In N. furzeri, dinucleotide repeats are the most common type of repeat (37%; Additional data file 5) and cumulatively occupy the third largest amount of sequence compared to the other tandem repeats ( Table 3). The repeat motif AC is the most frequent (26%), which is slightly less than in tetraodon and stickleback (30% each), and considerably more than in medaka (10%) and zebrafish (15%; Additional data file 5).
Minisatellites are far more abundant in N. furzeri than in the other four fish species. In particular, a 77-nucleotide minisatellite is most frequent. It comprises approximately 10% of the 5.4 Mb (Table 3) and its consensus sequence has a G+C content (63.6%) well above the genome average. Sequence conservation is high, for example, in an alignment of 189 repeat monomers; 65 of 77 positions (84%) are identical in at least 90% of monomers ( Figure 4). Using in situ hybridization we found that this minisatellite localizes to centromeric regions of many chromosomes (Figure 5a). Also, the 77-nucleotide minisatellite is N. furzeri specific as we did not detect this or similar tandem repeats in available genomic sequences of other fishes and vertebrates. We identified several other abundant and G+C-rich minisatellites (Table 3), which also localize to centromeric regions. For example, a 49-nucleotide minisatellite is also found in centromeric regions of many chromosomes, whereas a 24-nucleotide minisatellite specifically localizes to centromeres of only two chromosomes (Fig- ure 5b, c). Based on these analyses, we conclude that the large blocks of heterochromatin in centromeric regions, which are visualized at the cytological level by G+C-specific staining methods (Figure 1c), are mainly composed of N. furzeri-spe-cific and abundant G+C-rich tandem repeats. We plan to isolate additional minisatellites to be used in a more elaborate, multi-color fluorescence in situ hybridization (FISH) study to  assess chromosome specificity and derive chromosome-specific probes.
Hundreds of repetitions of the minisatellite motif 5'-TTAGGG-3' are found at telomeric ends of vertebrate chromosomes [39] and with associated proteins keep telomeres in homeostasis [40]. The telomeric repeat is not contained in the 5.4 Mb of N. furzeri. Most likely this is due to the limited sample size and so we searched for the repeat motif in the 111.6 Mb sample. We found 50 sequences, 14 of which are entirely composed of [TTAGGG] n . Correspondingly, in FISH experiments using the 5'-TTAGGG-3' motif as probe, specifically the terminal ends of chromosomes were labeled (I Nanda, unpublished results). Thus, characteristic vertebrate telomeric structures are present in N. furzeri and will be described elsewhere [15].
In addition to the G+C-rich minisatellites reported above, there is a prominent satellite repeat in N. furzeri. It comprises approximately 6% of our genomic sequence samples and has a monomer length of 348 nucleotides ( Table 3). The G+C content of its consensus sequence (41.1%) is below the genome average. This satellite accounts for a second anomaly in the G+C distribution of N. furzeri; it causes an excess of sequences at approximately 41% G+C (Figure 3a). We found that this repeat also localizes to centromeric regions of many chromosomes (Figure 5d). It is currently not clear if higher order repeat structures are formed by the two types of tandem repeats and if these are specific to certain chromosomes, or if and how this impacts on the structural organization of the N. furzeri genome [41,42]. A direct influence on N. furzeri life span seems unlikely, since the tandem repeat content of the GRZ and MZM-0403 strains is identical while their life spans differ by a factor of two. It is tempting, however, to speculate that these tandem repeats play a role in cell division, and that both the exceptional size and composition of N. furzeri centromeres might be functionally linked to the extremely fast growth of young fish. In mammals, repetitive DNA becomes demethylated over age, which might affect chromosome structure in constitutive heterochromatin composed of such repeats. Since DNA methylation is also observed in fishes [43][44][45][46], it will be interesting to analyze whether methylation occurs in the G+C-rich tandem repeats of N. furzeri and whether methylation changes with age, or differs between strains exhibiting different life spans.
In summary, tandem repeats, comprising a total of 20.6% of the genome, are exceptionally abundant in N. furzeri. They are 4-12 times more frequent than in zebrafish (5.0%), tetraodon (3.6%), stickleback (2.1%) and medaka (1.7%; Table 2). In tetraodon, two major satellites are reported [31]. One is a subtelocentric, highly variable ten-nucleotide minisatellite, which is also the most prominent tandem repeat in this fish, while the other is a centromeric, somewhat variable 118nucleotide satellite also found in Takifugu rubripes.
Interspersed transposon-derived repeats are mobile genetic elements and common to many eukaryotic genomes. Using RepeatMasker and Repbase Update, the reference library of vertebrate repeats [47], we found that 7-9% of the N. furzeri 5.4 Mb samples is composed of known interspersed repeats ( Table 2). Of these, approximately 6.3% are retrotransposons and approximately 1.5% DNA transposons. To identify novel transposon-derived repeats, we used the ab initio repeat identification program RepeatScout [48], which was recently found to be suitable for analyzing short sequences, although originally developed to analyze longer sequences [49]. Accordingly, 16-18% of our N. furzeri genomic samples are composed of novel interspersed repeats. Unfortunately, a detailed classification of the novel repeats is currently not feasible, again due to the limited/fragmented sample size. However, we assume that the fraction of these repeats identified in the 5.4 Mb closely resembles the overall fraction present in the N. furzeri genome, because our estimate for the medaka genomic sample corresponds well with the estimate given for the medaka draft genome [20]. In total, interspersed repeats comprise 24.7% of the N. furzeri genome, which isconsiderably more than in tetraodon (3.4%), stickleback (4.5%), and medaka (13.6%), and less than in zebrafish (35.4%). The overall repeat content in N. furzeri thus amounts to approximately 45%, which is the highest among the analyzed fish species. We note that this value might still be an underestimate, as we can not exclude that the interspersed repeat content might be found to be higher upon analysis of larger genomic samples since it is conceivable that we have missed 'rare' repeats in our samples.

Protein coding sequences
We attempted to explore the protein coding fraction contained in the 5.4 Mb of N. furzeri GRZ genomic sequence with respect to conservation in medaka, stickleback, tetraodon, zebrafish and human. As indicated above, we found that 444 of the N. furzeri GRZ sequences (473 kb of the 5.4 Mb) bear fragments of protein coding genes (Additional data file 2). Roughly one-third (152 kb) of the sequences are actually coding, and most of these (399, 94.3%) contain one to three exons. The G+C content of the 444 sequences (44.1%) fits the genome average and is considerably higher (50.3%) in the coding portions. Similar observations were made in tetraodon and medaka [19,20].
Of the 444 sequences, 310 (70%) show best matches to fishes, while 32 (7%) and 23 (5%) match best to human and mouse, respectively (Additional data file 2 In a complementary approach, we cloned complete coding sequences of five N. furzeri genes, Cdkn2b, Cdkn2d, Msra, Sirt1 and Tp53. Their gene products are involved in different aspects of aging and functionally conserved across taxa [1,50,51]. Sirtuin1 is a NAD(+)-dependent histone deacetylase considered to mediate the life span extending effects of calorie restriction [52], and its aging regulating function is conserved from yeast to mammals. Msra repairs and protects proteins from oxidation [53], and mice with mutations in it show a reduced life span [54]. Cdkn2b, Cdkn2d and Tp53 control cell cycle progression and mediate senescence [55][56][57]. A comparison of respective N. furzeri and human proteins should give an indication as to the potential use of this fish as a model for human biology. The five N. furzeri proteins are 40-65% conserved in humans (Table 4), which is in the range reported for orthologous proteins of teleost fishes and humans (60-70%) [19]. For comparison with zebrafish, which is perhaps the best established fish model for human biomedical phenotypes [58], we deduced zebrafish Cdkn2b, Cdkn2d, Msra, Sirt1 and Tp53 genes based on database searches and alignments of expressed sequence tags to genomic DNA (Additional data file 7). Conservation of inferred zebrafish proteins to human orthologs is in the same range (41-65%) as for N. furzeri and humans (Cdkn2b, 41%; Msra, 65.3%; Sirt1, 48.9%; Tp53, 48%). Conservation of the five N. furzeri aging relevant genes with respect to the other four fish species is highest in medaka, followed by stickleback, tetraodon and zebrafish (Table 4), the same as for the 180 random gene fragments described above.
Finally, we attempted to relate the 444 gene fragments and 5 aging related genes with respect to conservation in humans. We analyzed exons encompassing at least 33 amino acids, which number 366 in the former data set and 21 in the latter. In these exons, conservation with human counterparts is considerably higher for random gene fragments than for aging related genes: 67% versus 62%. Currently, we can only specu-late about the reasons for this difference. One possibility might be that it reflects a bias introduced by the sampling procedure of the 444 gene fragments. On the other hand, the difference might be related to the huge difference in life span between N. furzeri and human.

Phylogeny
In an initial attempt at phylogenetic reconstruction, we used the N. furzeri 28S rRNA gene (Additional data file 8) in comparison with 15 fish and two tetrapod sequences. However, standard parsimony analysis resulted in tree topologies much different from widely accepted taxonomic groupings (data not shown), indicating that the phylogenetic information contained in the 28S rRNA locus does not resolve the evolutionary history of fishes and tetrapods. We thus attempted to infer the phylogenetic positioning of N. furzeri with respect to the four fish model species and humans based on the 444 partial protein coding sequences and aging related genes identified in this work. A phylogenetic tree obtained by the maximum parsimony approach (Figure 6) is fully consistent with the taxonomic classification of the analyzed fish species. Accordingly, Nothobranchius is most closely related to medaka. Zebrafish and humans are both distantly related to the euteleost fish division consisting of the Atherinomorpha Nothobranchius and medaka on the one hand, and the Percomorpha stickleback and tetraodon on the other. The same tree topology was obtained using the neighbor-joining method, giving slightly lower bootstrap confidence (data not shown). We conclude that the coding fragments are sufficient to create a stable phylogeny, as previously suggested [59]. Our data are in agreement with phylogenetic analyses performed for ten fish species, which, based on expressed sequence tag analyses, found both killifish and medaka to be more distant from pufferfish than from each other [59].

Genetic variability
The N. furzeri GRZ strain studied in this work most probably originates from a single pair, which stems from wild fish captured in 1968, and was used for breeding more than 30 years ago [11]. If we assume a generation time of 6 months -comprising 12-16 weeks following hatching and about the same time for embryonic development -GRZ fish would have been inbred for approximately 80 generations until today. Thus, it was generally assumed that GRZ fish are highly homogeneous [17], although experimental proof was lacking.
To assess the genetic variability of the GRZ strain, we genotyped eight microsatellites (four di-, three tri-and one tetranucleotide repeat, identified in the 5.4 Mb), in 20 GRZ specimens. All microsatellites are monomorphic; we observed only one allele in the 20 specimens, respectively (Additional data file 9). To extend the analysis to gene associated markers, we resequenced parts of 19 protein coding genes encompassing 4 exonic and 15 intronic regions (10.2 kb in total). Again, there was no sequence variation in the 20 GRZ fish. In a third approach, we performed multi-locus fingerprinting in ten GRZ specimens using the 5'-GGAT-3' repeat as a marker [60]. As for the other markers, all specimens were homozygous at all 5'-GGAT-3' marker loci. Thus, we provide here the first experimental confirmation that the GRZ strain is highly inbred. In contrast, N. furzeri strain MZM-0403, bred in the sixth captive generation in our facility, exhibits 2-6 alleles in the microsatellites and has single nucleotide variations in 14 of the 19 genes. Also, a laboratory strain of the closely related species N. kunthae is highly heterogeneous in all markers that could be analyzed (Additional data files 9 and 10). While the inbreeding status of the GRZ strain is advantageous for further genetic analyses, it is currently unclear to what extent its short life span results from inbreeding depression [61]. A reduction of fitness can be caused by the accumulation of deleterious alleles [62], and reduced genetic diversity often affects birth weight, survival, reproduction and resistance to disease [63]. In this respect is noteworthy that the N. furzeri GRZ strain is derived from one 'normal' breeding pair that, 35 years ago, lived "a good 4 and a half months" (16-18 weeks), and that "healthy young fish" were used for spawning [11]. In our facility, the 10% survivorship (that is, the maximum life span) of direct descendants of these fish is 16 weeks and the fish can live up to 21 weeks [15], which is well in the range reported for the founders. Furthermore, there are differences in age-related histological damage between GRZ and age matched MZM-0403, which argue for accelerated aging in the GRZ strain rather than inbreedingrelated early death. For example, measurement of lipofuscin, which is an autofluorescing pigment that accumulates over age in a number of species [64], shows accelerated aging in brain and liver of the GRZ strain compared to age matched MZM-0403 [18]. To assess the degree of inbreeding depression on the life span of N. furzeri GRZ, access to new N. furzeri wild isolates from the GRZ game reserve in Zimbabwe or a nearby location is highly desirable. These fish could be cross-bred with the inbred GRZ strain and traits related to fitness, such as growth rate, reproduction, and survival, could be recorded and compared to the parental GRZ strain/wild isolate. On the other hand, it has to be noted that inbred lines are extremely powerful tools for biomedical research; if not subjected to severe inbreeding depression, they are valuable in many respects as they provide a uniform and stable genetic background and respective phenotype(s). Our results prove Evolutionary relationships of N. furzeri to medaka, stickleback, tetraodon, zebrafish and human Figure 6 Evolutionary relationships of N. furzeri to medaka, stickleback, tetraodon, zebrafish and human. The evolutionary history based on the maximum parsimony (MP) method [79] and MEGA4 [78] is shown and the most parsimonious tree given (length = 3,287). The tree topology is fully consistent with the taxonomic classification of the analyzed fish species. Accordingly, Nothobranchius is most closely related to medaka, while both zebrafish and human are distantly related to the euteleost fish division.

Conclusion
Our analyses provide a first glimpse into the genome of the short-lived annual fish N. furzeri, which we would like to support as a new vertebrate model for age research, as has been suggested previously [9,12,17].
At an estimated 1.6-1.9 Gb, N. furzeri has the largest genome compared with medaka, stickleback, tetraodon and zebrafish, which serve as models in many areas of contemporary research and for which genome projects have been completed or are underway. At the cytological level, a compartmentalization of the N. furzeri genome is apparent, which is caused by an accumulation of G+C-rich heterochromatin in centromeric regions of most chromosomes. This is mirrored in the composition of N. furzeri repeats, which among the analyzed fishes occupy the largest genomic fraction and are distinguished by an unmatched portion of G+C-rich and -poor tandem repeats. The most abundant G+C-rich species-specific minisatellites localize to centromeric regions and cause an unusual second peak in the G+C content distribution of the N. furzeri genome. The exceptionally high tandem repeat and G+C content is neither an inbreeding artifact of the GRZ strain nor can it be directly linked to life span, since genome structure and repeat content do not differ between the N. furzeri GRZ and MZM-0403 strains, although the former is highly inbred and the latter is a recently wild-derived strain and they differ in life span by a factor of two. The high tandem repeat and G+C content rather represent an N. furzeri-specific feature since this is much less pronounced in the closely related species N. kunthae, and has so far not been observed in vertebrates.
The N. furzeri and N. kunthae genomic sequences are a source of markers that can be used to build a first generation linkage map as well as to assess the genetic variability of Nothobranchius laboratory strains and natural populations with different life spans. An initial marker set provided the first molecular confirmation that the N. furzeri strain GRZ is highly inbred in contrast to the recently wild-derived strain MZM-0403 as well as the N. kunthae laboratory strain, which were found to be very heterogeneous.
Cross-breeding of N. furzeri strains with different life spans is currently being performed in our laboratory and should enable the identification of quantitative trait loci and facilitate cloning of aging-relevant genetic determinants. The present study illustrates the challenges that will have to be addressed in an N. furzeri genome project that we would like to establish in order to make maximal use of this fish species as a vertebrate model for aging research.

Chromosome preparations and banding analyses
To obtain mitotic chromosomes, healthy fish were exposed to a 0.03% colchicine solution for 8-12 h in a well ventilated container. Then fish were sedated on ice, killed by caput dislocation, and gills, spleen and kidney carefully removed. Preparation of cell suspension, hypotonic treatment, fixation of cells, and preparation of slides were as described [65]. Conventional chromosome staining, fluorescence staining with DAPI (4',6-diamidino-2-phenylindole) and mithramycin were as described [66]. To enhance the mithramycin fluorescence, slides were first pre-incubated with distamycin A. The location of constitutive heterochromatin on chromosomes was visualized as reported [67].

Flow cytometry measurements
Fin clips were taken from three N. furzeri GRZ females and at least 10,000 cells per sample measured on a Cell Analyzer CAII (Partec, Muenster, Germany). As reference, similar numbers of erythrocytes of a female chicken were used. Preparation of cell suspensions, fixation, DAPI staining and measurements were as described [28]. PI staining was performed at a final concentration of 50 μg/ml for 30 minutes at 37°C as described [60]. For each dye, three independent measurements were performed.

Fluorescence in situ hybridization
FISH mapping of repetitive DNA on N. furzeri GRZ chromosomes was performed as described [68]. Briefly, a biotin labeled plasmid clone bearing a specific tandem repeat was denatured and hybridized overnight at 37°C to denatured metaphase chromosomes. After post-hybridization washes at low (2× SSC, 50°C) and moderately (1× SSC, 60°C) stringent conditions, hybridization sites were detected with fuorescein isothiocyanate (FITC) conjugated avidin (Vector Laboratories, Burlingame, CA, USA) followed by signal amplification through incubation of slides with biotinylated anti-avidin and FITC conjugated avidin. Slides were examined on a microscope with a CCD camera (Zeiss, Goettingen, Germany); chromosomes showing specific FITC signals were displayed on counter-stained DAPI metaphases using appropriate software (Applied Spectral Imaging, Neckarhausen, Germany).

DNA/RNA
For genomic library construction, DNA of one male specimen of each of N. furzeri GRZ, N. furzeri MZM-0403, and N. kunthae, was isolated from 30 mg frozen tissue with the Blood and Cell Culture DNA Mini kit (Qiagen, Hilden, Germany). For marker validation, genomic DNA from 20 N. furzeri GRZ, 10 MZM-0403 and 10 N. kunthae specimens was isolated using 30 mg frozen tissue and AquaGenomicSolution (MobiTec, Goettingen, Germany), respectively. For GRZ, 50 ng DNA from ten specimens each were combined in a pool. For cloning of known aging related genes, RNA was isolated from one N. furzeri GRZ male using the RNeasy Mini kit (Qiagen). Primer sequences are given in Additional data file 11.

Sequencing
For shotgun sequencing using Sanger technology, randomly sheared,  (Table 1). These sequences were not assembled prior to analyses and submitted to the trace archive at NCBI.

PCR
Primers were designed using the GAP4 module of the Staden Sequence Analysis Package [72] and ordered from Metabion (Martinsried, Germany). For PCR, 50 ng DNA, 10 pmol of each primer and PuReTaq Ready-To-Go PCR beads (GE Healthcare, Munich, Germany) were used; initial denaturation was at 94°C for 30 s followed by 35 cycles of 60 s at 94°C, 60 s at primer annealing temperature, 60 s extension at 72°C, and a final extension at 72°C for 300 s. PCR products were sequenced as described above. Sequences were visualized, edited and assembled in GAP4.

Marker generation and experimental validation
Microsatellites were identified in the 5.4 Mb of N. furzeri GRZ using Sputnik (see Repeat identification). Flanking primers were designed in sequences with: one microsatellite; at least 20 repeat units for dinucleotide repeats, or at least 10 repeat units for tri-and tetranucleotide repeats; at least 100 nucleotides of flanking sequence; perfect repeats. PCR was performed in two pools of GRZ DNA (see DNA/RNA) with one primer being 6-FAM-labeled. For genotyping, ABI 3730xl sequencers and GeneMapper software Version 4 was used.
Allele calling was performed independently by two individuals. Discrepancies were resolved by retyping. For resequencing of gene-associated markers, primers were designed in 19 sequences with significant homology to known genes in at least one of the four fish reference genomes. PCR and sequencing was performed in two DNA pools of GRZ, and ten individual N. kunthae fish as described above.

Reference data sets
In order to compare our genomic data of N. furzeri strains GRZ and MZM-0403, and N. kunthae, which we obtained by random genome-wide sample sequencing, we simulated this approach in silico for the other four fish species. That is, we extracted random genomic data from the publicly available genome assemblies for each. In detail, these were 20 samples of random genomic sequence sets of medaka, stickleback, tetraodon and zebrafish from genome assemblies at Ensembl [21]

BLAST analyses
To determine the coding fraction, a BLASTX search in Swiss-Prot/TrEMBL (version 50.6) was performed. Hits with p < 10 -10 were regarded significant. Reference data sets of the four fish species were analyzed in the same way. Then, N. furzeri GRZ and MZM-0403 sequences containing coding portions were used for TBLASTX searches in medaka, stickleback, tetraodon and zebrafish cDNA collections downloaded from Ensembl [21] (see 'Reference data sets' above for release dates). A p-value < 10 -10 was set as the threshold for significance.

Repeat identification
First, tandem repeats were identified and masked (replaced by Ns) using Tandem [74], with standard parameters. Third, for de novo repeat identification, RepeatScout, version 1.0.2 [48] with standard parameters, except for a minimum repeat copy number of 3, was used on the pre-masked sequences. Finally, a species specific repeat consensus library was built using both the identified known repeat families and the previously unknown repeat consensus sequences.

Phylogenetic analyses
Of the 444 gene fragments of N. furzeri GRZ, we first identified 44 for which we could unambiguously assign human orthologs based on BLASTX searches in Swiss-Prot. Based on these assignments, we searched for orthologous regions in medaka, stickleback, tetraodon, and zebrafish genome assemblies at University of California Santa Cruz (UCSC) [75] (oryLat1, tetNig1, gasAcu1, danRer5, hg18) by inspection of MAF tracks. We extracted exons, which we regarded as true orthologs if: no duplicated gene was detected in the respective species; splice sites were conserved; and no frameshifts were detected if consecutive exons were concatenated. In total, this resulted in 71 orthologous exons, identified in all six species (average length is 142 bp for human exons), which represent fragments of 26 protein coding genes (sequences are listed in Additional data file 12, including exons of N. furzeri GRZ). Five of the gene fragments represent one exon, while the others span at least one splice site. For further analyses, sequences were concatenated, translated, aligned by Clus-talW v1.81 with standard parameters [76] and trimmed at the ends if necessary. Coding sequences of the aging related genes Msra, Sirt1, and Tp53 were also translated and aligned. All alignments were concatenated, resulting in 4,828 amino acid positions, and analyzed by MEGA4 [77] with different methods (alignment available on request). Positions containing gaps and missing data were eliminated, leaving a total of 4,245 positions in the final dataset, of which 508 were parsimony informative. A maximum parsimony tree was obtained using the Close-Neighbor-Interchange algorithm in which the initial trees were obtained with the random addition of sequences (100 replicates).

Sequence comparison
Nucleotide sequences of putative orthologs of the four analyzed fish species were identified as described above and are listed in Additional data file 7. Nucleotide and translated sequences were aligned using needle (EMBOSS, version 3.0.0 [78]).


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dRecQ4 Is Required for DNA Synthesis and Essential for Cell Proliferation in Drosophila

Background The family of RecQ DNA helicases plays an important role in the maintenance of genomic integrity. Mutations in three of the five known RecQ family members in humans, BLM, WRN and RecQ4, lead to disorders that are characterized by predisposition to cancer and premature aging. Methodology/Principal Findings To address the in vivo functions of Drosophila RecQ4 (dRecQ4), we generated mutant alleles of dRecQ4 using the targeted gene knock-out technique. Our data show that dRecQ4 mutants are homozygous lethal with defects in DNA replication, cell cycle progression and cell proliferation. Two sets of experiments suggest that dRecQ4 also plays a role in DNA double strand break repair. First, mutant animals exhibit sensitivity to gamma irradiation. Second, the efficiency of DsRed reconstitution via single strand annealing repair is significantly reduced in the dRecQ4 mutant animals. Rescue experiments further show that both the N-terminal domain and the helicase domain are essential to dRecQ4 function in vivo. The N-terminal domain is sufficient for the DNA repair function of dRecQ4. Conclusions/Significance Together, our results show that dRecQ4 is an essential gene that plays an important role in not only DNA replication but also DNA repair and cell cycle progression in vivo.

Cancer predisposition of either human patients or mice models with RecQ4 mutations represent another unresolved issue (for review, see [2]). Mutations in the human RecQ4 gene have been found to contribute to three rare syndromes: Rothmund-Thomson syndrome [5,[30][31][32], RAPADILINO syndrome [22,25] and Baller-Gerold syndrome [2,23]. Currently there is no common conclusion on whether these three syndromes are independent disorders or represent one syndrome with different symptoms. Several labs have developed mice models with different RecQ4 mutations, but these mice show different phenotypes that range from embryonic lethality to defects restricted to adult mice, some of which resemble the symptoms of human patients [19,28,33].
Several recent studies have revealed new insights concerning the role of RecQ4 in DNA replication initiation [18,20,21,24]. Cut5, the metazoan homolog of Saccharomyces cerevisiae Dpb11, which is required for loading DNA polymerases onto chromatin, was shown to interact with the Xenopus RecQ4 (xRecQ4) both in vitro and in vivo [20,21]. Purified N-terminal fragments of xRTS/ xRecQ4 were able to rescue the DNA replication defects of xRecQ4-depleted Xenopus egg extracts [20]. In mammalian cells, RecQ4 has been shown to interact with RAD51 and PARP1, suggesting that it may also participate in DNA repair [2,10,34,35]. However, the role of RecQ4 in DNA repair has not been fully characterized, particularly in the context of an in vivo system. Unlike in mammals, the fruit fly genome encodes three complete RecQ helicases, namely dBLM, dRecQ4 and dRecQ5 [24,[36][37][38][39][40][41][42][43]. In addition, DmWRNexo was recently identified as the Drosophila homologue of human WRN exonuclease domain [44,45]. In order to develop a model system more amenable to genetic analysis of RecQ4 function in vivo which would also help to clarify, at least, some of the controversies about RecQ4, we set out to characterize RecQ4 in Drosophila.
In this report, we describe the generation and phenotypic analyses of dRecQ4 mutants in Drosophila. Our results show that the dRecQ4 mutants exhibit defects in DNA replication. They are also selectively sensitive to paraquat and gamma irradiation. Mutant animals exhibit lower efficiency of double strand break (DSB) repair as assayed by reconstitution of the DsRed transgene in vivo [46]. Rescue experiments with various truncated dRecQ4 proteins suggest that the N-terminal domain of dRecQ4 is essential for DSB repair, whereas both the N-terminal domain and the helicase domain are indispensable for DNA replication and animal viability.

dRecQ4 is essential for development
Prior to the report of Wu et al., there were no transposable elements inserted within or nearby the dRecQ4 locus [24]. We took advantage of the targeted knockout technique to generate dRecQ4 mutants through the replacement of the endogenous locus with an engineered mutant form via homologous recombination. Specifically, an 8 kb genomic fragment was modified by replacing the start codon ATG with CCTAGGGTCGACCCGCG and inserting an I-SceI recognition site into the second exon of dRecQ4 ( Figure 1A; see Materials and methods for details). Targeting of the dRecQ4 locus was achieved by a modified procedure described by Rong and Golic [47] and Egli and colleagues [48,49]. Four mutant candidates were obtained and confirmed by restriction enzyme digestions; all four alleles showed similar phenotypes in the viability test (see below). One of these alleles, dRecQ4 14 , was further confirmed by DNA sequencing ( Figure 1B) revealing that the start codon mutation and open reading frame shift are as designed. This allele, which can be fully rescued by a genomic rescue transgene as judged by adult flies' viability (see below), was used for detailed phenotypic analysis throughout this study. dRecQ4 14 mutants are homozygous lethal, indicating that dRecQ4 is an essential gene. Using a GFP marked balancer chromosome, we separated the homozygous from the heterozygous dRecQ4 14 animals. Nearly all the homozygous mutants survive for up to 8 days under normal culture conditions. However, they exhibit developmental delays when compared with heterozygous siblings or y w wild type flies and eventually die at early pupal stage ( Figure 1C). The lethal phenotype of the dRecQ4 14 mutants can be fully rescued by either a genomic fragment of dRecQ4 or a UAS mediated dRecQ4 expression (data not shown), further indicating that the lethality phenotype was a direct consequence of the dRecQ4 mutation.
To determine the role of maternal contributions to development, we set out to generate dRecQ4 germline clones in females that carry FRT combined dRecQ4 mutation and ovo D chromosomes (Table 1). However, when these females were crossed to y w; dRecQ4 14 /TM3, Kr-GFP males, no eggs were obtained ( Table 1). The oocytes from dRecQ4 14 FRT2A/FRT2A ovo D females failed to go beyond stage 6 of oogenesis (data not shown). These results suggest that dRecQ4 is essential for oogenesis, further supporting the conclusion that dRecQ4 is an essential gene critical to cell viability.
dRecQ4 loss-of-function affects endogenous DNA replication both in the salivary glands and the late larval brain It has been shown in Xenopus egg extracts that DNA replication is blocked when RecQ4 is depleted [20,21]. To specifically determine whether dRecQ4 is essential for DNA replication in vivo, we measured both DNA content and cell numbers of salivary glands from wild type and dRecQ4 mutant animals. Figure 2B shows that salivary glands from wild type and mutants have similar cell numbers at third instar larval stage (5 days AED). However, the total amount of DNA from each salivary gland at this stage differs significantly between wild type and mutants ( Figure 2A). The amount of DNA normalized by cell number is much lower in mutant cells than in wild type cells (,0.18 ng/cell and ,0.92 ng/ cell, respectively), indicating a defect in DNA accumulation, presumably reflecting an under replication of DNA. Salivary gland increases its DNA content through cell cycle independent endoreplication. Inefficient DNA endoreplication of salivary gland cells is also consistent with the finding of small cells and nuclei in this tissue of mutant animals ( Figure 2C).
To directly investigate whether dRecQ4 is required for DNA replication, particularly in the non-endoreplicating cells, we performed BrdU labeling followed by anti-BrdU immuno-staining on the larval brain. Figure 3 shows that at the stage of four days after egg deposition (AED), wild type and dRecQ4 mutants have comparable BrdU incorporation likely reflecting maternal contributions of dRecQ4 protein in the mutants. However, at five days AED the mutant brain incorporates much less BrdU than the wild type, which is consistent with the observed reduction of DNA accumulation in salivary glands (see above). Our data are consistent with those of Wu ) and together they demonstrate that dRecQ4 is involved in DNA replication in Drosophila.
dRecQ4 is involved in double strand breaks repair Inactivation of RecQ4 in mouse results in defective sister chromatid cohesion and aneuploidy [50]. To determine whether dRecQ4 deficiency causes a similar effect on chromosomal behavior in Drosophila cells, we analyzed metaphase spreads from wild type and mutant brain cells. In dRecQ4 mutants, the spreads' patterns fall into three major categories ( Figure 4A-C): normal pattern ( Figure 4A), segregated ( Figure 4B) and fragmented ( Figure 4C) aberrant patterns. The segregated patterns are indicative of a failure of sister chromatids association (arrows in Figure 4B). The fragmented patterns have broken chromosomes, often with the broken ends fused together (arrow heads in Figure 4C). Statistical analysis shows that the frequency of aberrant patterns in dRecQ4 14 mutants is much higher than in wild type control. Specifically, mutant cells have less than 10% of normal patterns, with the aberrant segregated and fragmented patterns representing the majority of the mitotic cell population. In wild type cells, over 80% of the cells have normal patterns, with only less than 20% being the mildly abnormal segregated patterns. These results suggest that dRecQ4 is important for maintaining genome integrity.
To investigate DNA repair pathways in which dRecQ4 participates, we treated mutants with various DNA-damaging mutagens including hydroxyurea (HU), methyl methane sulfonate (MMS), paraquat and gamma irradiation ( Table 2). These mutagens exert their effects through distinct mechanisms and, thus, can provide insights into DNA repair defects in dRecQ4 mutants. Our results show sensitivity of dRecQ4 14 mutants to paraquat and gamma irradiation ( Table 2). Paraquat mainly causes single-base damage which is corrected through base excision repair pathway. Using T7 phage display screen, human RecQ4 was found to interact with poly(ADP-ribose) polymerase-1 (PARP-1), an enzyme that maintains genome stability through its involvement in base excision repair pathway ( [34]). Together with our in vivo data, they demonstrate that RecQ4 is involved in the base excision repair pathway. DNA double strand break (DSB) is the major type of DNA damage after gamma irradiation, the sensitivity of dRecQ4 14 mutants to gamma irradiation suggests that dRecQ4 is also involved in the DSB repair pathways.
To demonstrate more directly that dRecQ4 participates in DSB repair, we employed the in vivo inducible DSB break-repair system in Drosophila (Fig. 5A; [46]). In this system, the reporter construct, Rr3, consists of a DsRed gene interrupted by the recognition sequence for the rare-cutting endonuclease, I-SceI. UIE is a transgene that expresses the enzyme I-SceI under the control of the ubiquitin gene promoter. The intact Rr3 element does not express a functional DsRed gene product owing to the presence of the cutting site. However, when a DSB is formed at the I-SceI cutting site, repair via the single-strand annealing pathway results in a functional DsRed gene ( Figure 5B a and c). We tested the repair efficiency of the induced DSBs both in the presence and in the absence of dRecQ4 14 mutation. When Rr3 is not cut, the heterozygous control and homozygous mutant animals have similar survival ratios ( Figure 5C. category b and d), which serves as a system control. However, when Rr3 is cut, the survival ratio is significantly reduced in dRecQ4 homozygous mutant background compared with heterozygous animals ( Figure 5C. category a and c), indicating a reduced efficiency of DSB repair in the absence of dRecQ4 gene function. These results provide additional support to our conclusion that dRecQ4 is involved in DSB repair in vivo.
dRecQ4 loss-of-function leads to mitotic (M) phase arrest and reduction of cell proliferation in late wing and eye imaginal discs Cell cycle progression is strictly controlled by a strong checkpoint system that arrests progression of the cell cycle until either DNA replication is completed or DNA damage is repaired [51]. To test whether DNA replication and/or repair defects in dRecQ4 14 mutants lead to cell cycle arrest in vivo, wing imaginal discs from wild type and mutant larvae were dissected, trypsinized and stained with propidium iodide (PI), followed by fluorescence activated cell sorting (FACS). Compared with wild type, more cells from dRecQ4 14 mutants are accumulated at G2/M phase at the expense of G1/G0 and S phase cells ( Figure 6A). It is notable that  the second peak is shifted with its position closer to the G1 peak in the dRecQ4 mutant cells comparing with that in the WT cells ( Figure 6A). A possible explanation could be that in the absence of dRecQ4, the S-M checkpoint becomes defective, which allows the mutant cells to enter mitosis with incompletely replicated DNA. Immunostaining using antibody against phospho-histone H3, which serves as M phase marker, shows a significant increase of M phase cells in the wing discs of dRecQ4 14 larvae compared with that of wild type ( Figure 6B).
To test whether the cell cycle aberration in the absence of dRecQ4 function leads to cell proliferation defect, two strategies were employed. First, we conducted the tissue specific knock-out of dRecQ4 function using tissue specific flipase that acts on a FRTflanking genomic rescue transgene. We generated a transgenic line, pTARG-dRecQ4, which harbors a dRecQ4 genomic segment flanked by two FRTs (see Materials and methods for details). This transgene is able to completely rescue the dRecQ4 mutant animals to adulthood without any observable defects ( Figure 7B). Taking    advantage of such rescued flies, we used ey-FLP to specifically delete the rescuing genomic fragment in the eye-antenna primordia. Figure 7C shows that tissue-specific knockout of dRecQ4 in the eye leads to small rough eyes with disorganized and fewer ommatidia, phenotypes that are indicative of reduced cell proliferation.
In a second strategy to determine whether cell proliferation is affected in the absence of dRecQ4, normal somatic clone analysis was employed (see Materials and methods for details). As shown in Figure 7D, dRecQ4 mutant clones can be detected adjacent to their twin clones (lower row) 72 hours after the induction of FLP expression by heat shock. Similar to the wild type (GFP 2 ) clones (upper row), the total number of dRecQ4 clones (GFP 2 , lower row) is similar to their twin clones (2XGFP). However, the dRecQ4 mutant clones (GFP 2 , lower row) are smaller than the 2XGFP twin clones (lower row), while the wild type (GFP 2 , upper row) clones are similar in size to their 2XGFP twin clones (upper row). The dRecQ4 mutant clones on average occupy about 20% of the territory that their twin clones occupy (2XGFP); in wild type clones, the size of GFP 2 clones is similar to that of their twin clones ( Figure 7E). Taken together, these data suggest that dRecQ4 mutant cells are aberrant in cell cycle progression, which may have resulted in cell proliferation defects.
Both the N-terminal domain and the helicase domain of dRecQ4 are essential to its in vivo function Our studies described thus far suggest that dRecQ4 plays a role in multiple processes including DNA replication, DNA repair and cell cycle progression. To clarify which function is primarily responsible for its essentiality and the protein domain for such function, we performed dRecQ4 functional domain dissection experiments. The dRecQ4 protein consists of 1579 amino acids including a helicase domain extending from aa 867 to 1208. A series of deletion mutants of the dRecQ4 coding sequence were generated, and the resulting truncated proteins are as shown in Figure 8. The corresponding transgenic flies were generated and analyzed for their ability to rescue the mutants' phenotypes including lethality, BrdU incorporation and sensitivity to DNA damaging reagents. The full length dRecQ4 is able to fully rescue dRecQ4 14 mutant to adulthood without any obvious defects. The C-terminal deletion form, dRecQ4[D1234-1579], can also rescue dRecQ4 14 animal to adulthood, but only at an efficiency of about 10%. Nevertheless, it can fully rescue the BrdU incorporation defects and the sensitivity to gamma irradiation of the mutant. Neither of the other truncations, lacking either the N terminal or the helicase domain or both, exhibited any ability to rescue the mutant animals to eclosed flies ( Figure 8). However, gamma irradiation sensitivity can be rescued fully by truncated forms that contain the N-teminal domain, namely dRecQ4D868-1579, dRecQ4D1234-1579 and dRecQ4D868-1207, in addition to the full-length protein. These results indicate that both the N-terminal and helicase domain of dRecQ4 are indispensable for animal viability, although the N-terminal domain alone is sufficient to rescue the mutants' sensitivity to gamma irradiation.

Discussion
The studies described in this report demonstrate that dRecQ4 is essential for Drosophila development. Loss-of-function of dRecQ4 leads to a failure of proper DNA replication and inappropriate cell cycle progression. The dRecQ4 mutant animals show a preferential sensitivity to gamma irradiation. Because gamma irradiation mainly causes DNA double strand breaks, this suggests an in vivo role of dRecQ4 in DSB repair, which is further supported by DsRed reconstitution experiment in vivo. While the C-terminal domain of dRecQ4 is not essential to its function, the N-terminal domain and the helicase domain are indispensable in vivo. Considering our combined results and what is known up-to-date about RecQ4, the following issues are worth further discussion.
(1) dRecQ4 is the only essential RecQ helicase in flies. The three Drosophila helicase genes, dBLM, dRecQ4 and dRecQ5, as well as the Drosophila homologue of human WRN exonuclease gene, dWRNexo, have now all been studied genetically [37,38,40,45] (Chen and Jiao, unpublished). Interestingly, among all the mutants of these dRecQ genes, only dRecQ4 mutants exhibit a homozygous lethal phenotype; mutants of the other dRecQ family members only show either fertility defects or various defects in DNA repair, including recombinational repair. There are several possibilities why dRecQ4 mutants show a more severe phenotype (i.e., lethality) than the other dRecQ mutants. First, dRecQ4 mutants clearly have defects in DNA replication. The defects occur not only in multiploid cells during endoreplication, but also in normal diploid cells. Interestingly, endoreplication defects do not result in total cell number changes although it does affect cell growth, while replication defects in normal diploid cells lead to cell proliferation abnormality. The simplest explanation for this observation is that, at late embryonic stages/early larval stage, the maternal contribution of dRecQ4 helps the mutant animal acquire proper cell number of the salivary glands before loss of zygotic dRecQ4 takes effect. However, by the time of second and third instar larval stages when the cells in the wing imaginal discs proliferate extensively, the maternal dRecQ4 has been diluted and/or degraded to a level that is insufficient to promote cell cycle progression ( Figure 6A). The failure of proper cell cycle progression and proliferation is likely to be the cause of lethality.
If the DNA replication defects were the primary cause of animal lethality, one would expect that a dRecQ4 domain that is responsible for DNA replication should rescue the lethality. However, our results show that the N-terminal domain of dRecQ4, dRecQ4g868-1579 (Figure 8), which is homologous to the xRecQ4 N-terminal domain, necessary and sufficient for DNA replication in Xenopus [20,21], does not rescue the animal's lethality. As shown in this study, dRecQ4 also plays an important role in DSB repair ( Figure 5). If the DNA repair defects were the primary cause for lethality of dRecQ4 mutants, one might expect that the other fly RecQ mutants that have DSB repair defects, such as dBLM, would also be lethal, which in fact is not the case [38]. Together, these considerations suggest that the lethality of the dRecQ4 mutants is likely a consequence of loss of both of its primary functions, namely DNA replication and DNA repair.
(2) RecQ4 deficient cells and animals show differential sensitivity to genotoxic agents. It has been controversial in the literature regarding the mutagen sensitivity for RecQ4 deficient cells and/or model organisms [4,15,[26][27][28]34,[52][53][54]. For example, it has been shown by two independent groups that RecQ4 is involved in UVinduced damage repair in human cells [15,52]. However, using fibroblasts derived from different RTS patients, Cabral et al. found that these cells are not sensitive to a wide variety of genotoxic agents including ionizing or UV irradiation, H2O2 and HU [53]. Jin [34]. At the animal level, the mouse model generated by Hoki et al. which bears in frame deletion of exon 13 RecQ4 shows normal sensitivity to IR and UV irradiation [28]. However, our Drosophila RecQ4 mutants are strongly sensitive to ionizing radiation, which suggests a role for dRecQ4 in DSB repair that is in agreement with what has been found in Xenopus [54]. The possible explanations for the conflicting sensitivity results of RecQ4 mutants reported thus far could be as follows: (i) it is possible that RecQ4 plays a more important role in DSB repair pathways in flies than in humans. There are five RecQ helicases in humans while there are only three in flies. The functions for RecQ members may be more specialized in humans than in flies. For example, in humans BLM and WRN are primarily involved in different DSB repair pathways with BLM more in homologous recombinational repair and WRN more in non-homologous end joining repair [4,[6][7][8]. Since there is only dWRNexo in flies, the homologous function of human WRN in flies may have been incorporated in dRecQ4 protein; (ii) more likely, cells derived from different human patients with different mutations or the same mutations in different genetic backgrounds could be also the causes of differential sensitivities. Xu and Liu recently found that the N terminal region and helicase domain of human RecQ4 both possess helicase activity [55], which argues for the possibility that human patients or mutant mouse with intact N terminal region have less severe sensitivity to mutagens. This is very well evidenced by our domain dissection study in Drosophila. The N terminal domain is sufficient to rescue dRecQ4 mutant's sensitivity to gamma irradiation.
(3) Although we expected dRecQ4 14 allele to be null based on the designed mutations of the translation start codon ATG and the frame shift of the coding sequence, the phenotypes are generally weaker than Wu's null mutant; for example, our mutants die at early pupal stage, but theirs at early larval stage. It is possible that our mutant allele is not completely null that may express a truncated form of dRecQ4; we note that the 1017th codon (for Figure 8. Functional dissection of dRecQ4 protein in vivo. Constructs that express full-length and different truncated forms of the dRecQ4 protein are shown. Full length dRecQ4 protein that consists of 1579 amino acids including a helicase domain extending from aa 867 to 1208 can rescue 100% of dRecQ4 14 mutants to adulthood. The C terminal deletion form (dRecQ4D1234-1579) of dRecQ4 can rescue dRecQ4 14 animals to adulthood with an efficiency of only 10%, but fully rescue the BrdU incorporation defects and gamma-irradiation sensitivity of the mutants. None of the other deletion forms can rescue any dRecQ4 mutant animals to adulthood. However, gamma-irradiation sensitivity can be rescued fully by truncated forms that contain the N-teminal domain, namely dRecQ4D868-1579, dRecQ4D1234-1579 and dRecQ4D868-1207, in addition to the fulllength protein. doi:10.1371/journal.pone.0006107.g008 methionine) is followed by a 562 amino acids in-frame reading frame of dRecQ4 (there is currently no appropriate antibodies available to detect this possible truncated protein of dRecQ4). Unlike the null allele of Wu that could not generate mutant somatic clones in the wing discs, our mutant is capable to support limited cell proliferation ( Figure 7D). Our mutant may thus represent a potentially useful tool in further mechanistic studies of DNA repair in vivo.
(4) Functional domain dissection combined with rescue experiments suggests that the essential functions of the dRecQ4 protein reside in the N-terminal and the helicase domain. A very recent report by Xu and Liu [56] has shown for the first time that human RecQ4 exhibits dual DNA helicase activity. Two distinct regions of the protein, the conserved helicase motifs and the Sld2like N-terminal domain, display independent ATP-dependent DNA unwinding activity. Although the N-terminal domain of RecQ4 is sufficient for DNA replication initiation in Xenopus ( [20]), our in vivo data clearly suggests the helicase domain is required for proper DNA replication in Drosophila. The C-terminal domain of dRecQ4 is dispensable for its essentiality, but the rescue efficiency of the truncated protein that lacks the C-terminal is only about 10% compared with the full length dRecQ4 protein. It is possible that the C-terminal domain modulates the protein activity of dRecQ4, possibly via amino acids modifications and/or interactions with other proteins.

DNA constructs
5 kb dRecQ4 genomic fragment (coding region of the gene) and 3 kb 59 regulatory sequences with intended modifications were cloned in the pTARG vector [49] to make the gene targeting construct,pTARG-dRecQ4. Changes were introduced by PCR with the following oligos (altered bases for either restriction sites and/or mutations are highlighted by underlining). The primers used to amplify the 5 kb dRecQ4 genomic sequence were 59-TCC-CCGCGGACGATTCGGTGTTCAAGCTAAAAT-39 and 59-GGACTAGTGCAGGATGCGATTGAAATCCACTT-39. The primers for amplifying the upstream 3 kb fragment were 59-AT-AAGAATGCGGCCGCGCTCTCCATCGTGATGGGCCT-39 and 59-GGCC TAGGGTCGGCGGCTGTCTTTAATTGTC-AATA -39. Mutation of ATGG to CCTAGGGTCGACCCGCGG generates a new restriction site (SacII) for identification of mutant DNA. Oligos used to introduce the I-SceI cleavage sequence at the MfeI cutting site were 59-AATTTAGGGATAACAGGGTAAT-39and 59-AATTATTACCCTGTTATCCCTA-39.
For constructing UAS-dRecQ4, primers 59-ATAAGAATGCGG-CCGCACATGGACGATTCGGTGTTC-39 and 59-GGGGT AC-CTCACGTACGCCTCTTGATAA39 were used to PCR the genomic DNA that spans from the start to the stop codons of dRecQ4 gene's coding region for putting into pUAST vector at Not I and Kpn I sites (start and stop codons are underlined).
pTARG-dRecQ4[rescue] construct contains 2.1 kb upstream of the ATG and 1.5 kb downstream after stop codon sequence in addition to the entire coding region. It was constructed by putting two PCR products into the pTARG vector at Not I and Avr II sites. The two pairs of primers used for PCR were as follows: 59-AATGAATTGCGGCCGCGTCGGGAACTACAGTCCAACCT-39/59-CGAAACCGGTTGGCTTAGGGAAGCTTCG-39 and 59-GCCAACCGGTTTCGCAAGAGAAAGCAGC-39/59-TAGACC-TAGGATGAAGGAGCACGGCCAAATGCCAG-39. Restriction sites for cloning are highlighted in italics.
For primers that are used for generating domain dissection constructs, please see Table 3. Detailed cloning strategies are available upon request.

Fly stocks and genetics
Flies were cultured at 25uC for all experiments. For generation of germline clones (GLCs), we used the FLP-DFS system as described [57]. Briefly, the dRecQ4 14 mutation was recombined with the third chromosomal FRT insertion 2A, balanced with TM6B, Tb balancer and crossed with ovo D , FRT2A males. Offspring of this cross were given a heat shock (37uC, 2 hrs) at late third instar larval stage and virgin females with correct genotypes (Table 1) were crossed with heterozygous mutant males. Table 3. Forward and reverse primers used to generate pUAST-dRecQ4g constructs that produce truncated proteins used in Fig. 8

Generation of dRecQ4 14 mutant
For generation of the dRecQ4 mutant, we used the ends-in gene targeting method [47,48]. Donor transgenic flies that bear the targeting construct on the second chromosome were crossed to flies that contain hs-I SceI and hs-FLP transgenes. Three heat shocks (38uC, 1 hr each) were applied on days 2, 3 and 4 after egg laying. Heat-shocked virgins were singly crossed to y w; ey-FLP; MKRS/TM2, y+ males, and females were screened for targeted integration of targeting construct indicated by the w + marker. Reduction of two dRecQ4 copies (one wild type and one mutant copy) by I-CreI was performed by crossing the targeted alleles to w1118; hs-I-CreI, Sb/TM6. The offspring were given a single heat shock (36uC, 1 hr) at the third instar larval stage. w 2 males were crossed individually to y w; actin-Gal4/TM3, Ser to make stocks. The allele, we designated dRecQ4 14 , was further characterized by DNA sequencing for the intended mutations, the primers for PCR were 59-TCCCAGCATGTGATAGTCTG-39 and 59-TCCTC-AAGATTACCAG AGCTC-39. The resulting data has been deposited in GenBank (accession number GQ128383).

Generation of somatic clones
Loss-of-function somatic clones were induced using FLP/FRT mediated mitotic recombination [58]. To induce the clones, first instar larvae with correct genotypes were heat shocked for 1 hour at 38uC and then dissected at third instar larval stage. Mutant clones and twin spot areas were measured with confocal images using the histogram function of Adobe Photoshop.

Immunohistochemistry
For BrdU labeling, wild type, mutant or rescued mutant larvae were dissected in PBS and then incubated in PBS containing 1 mg/ml BrdU (Sigma B-5002) for 30 min at 25uC. After three rinses with PBS, samples were fixed for 30 min in 4% paraformaldehyde followed by washing 3 times in PBST (PBS, 0.1% Triton X-100) and then treatment with 2M HCl for 30 min. After three washes in PBST, samples were incubated with mouse anti-BrdU (1:100, ZYMED). TRITC-conjugated anti-mouse secondary antibody (Jackson ImmunoResearch Laboratories) was used with dilution of 1:100 for 2 hrs at room temperature. Images were taken under a Leica DM6000 confocal microscope. More than 10 brains were examined per genotype. Rabbit anti-phospho histone H3 (Ser 10) antibody (1:100) for detecting mitotic phase was purchased from Millipore. FITC-conjugated anti-rabbit secondary antibody (1:100) was from Jackson ImmunoResearch Laboratories. Over 10 discs were analyzed for each genotype.
Fluorescence activated cell sorting 80-100 wing discs of the same genotypes (the mutant or wild type) were dissected in PBS and digested with trypsin-EDTA (Sigma, T-4174). Cells were dissociated for 5 hours by gentle shaking. The dissociated cells were fixed in 75% ethanol, stained with propidium iodide and analyzed with a Becton Dickinson Vantage Fluorescence activated cell sorter. The events of either the wild type or the mutant were 10000. The experiment was repeated 3 times.

DNA content measurement
Genomic DNA from 100 salivary glands of wild type or dRecQ4 14 mutant larvae (5 days old) was extracted, followed by A260/280 measurement. Two sample independent t-test was used to determine statistical significance.

Chromosome spreads
Brains from wild type Canton S and dRecQ4 14 homozygous third instar larvae were dissected in 0.7% saline, treated with colchicine and hypotonic solution, fixed in acetic acid/methanol/H2O (11:11:2) and stained for 5 min in 0.2 mg/ml DAPI. The preparations were examined under Leica DM6000 fluorescent microscope.

DNA damage sensitivity tests
Four mutagens were used in this experiment (hydroxyurea, methyl methane sulfonate, paraquat and c-irradiation, see Table 2). All chemicals were purchased from Sigma. Chemical mutagens were added to fly food at final concentrations that have been used for DNA damage assay in Drosophila (see also Table 2). 20 females and 10 males of y w; dRecQ4 14 /TM3, Sb,Kr-GFP flies were put in the vial that contained different mutagens for 6 h before the parents were discarded. At the 6 th day, the heterozygous (GFP + ) and homozygous (GFP 2 ) mutants were scored. The survival ratio was measured as the number of GFP 2 to half the number of GFP + . For c-irradiation, eggs from y w; dRecQ4 14 /TM3, Kr-GFP flies were collected for 5 hrs and allowed to develop for 12 hrs before being exposed to 9 Gy c-irradiation with a 60 Co source. Only gamma irradiation was used to test the sensitivity of the rescued mutants to DNA damaging reagent. The same dosage was applied as above.

DsRed repair assay
DsRed DSB repair model was carried out essentially according to the Preston method [46]. Since nearly all the dRecQ4 14 homozygous mutants can live to early pupal stage, the relative survival ratio from early instar larvae to early pupae stage was used to indicate the repair efficiency indirectly, assuming the lethality is caused by unrepaired DSBs. The cross was made as follows: First instar larvae and early pupae representing each categories were counted for calculation of the relative survival ratio.