Genetic and cytogenetic analysis of the fruit flyRhagoletis cerasi (Diptera: Tephritidae)

Ilias Kounatidis, Nikolaos Papadopoulos, Kostas Bourtzis, andPenelope Mavragani-Tsipidou

Abstract: The European cherry fruit fly, Rhagoletis cerasi, is a major agricultural pest for which biological, genetic, andcytogenetic information is limited. We report here a cytogenetic analysis of 4 natural Greek populations of R. cerasi, allof them infected with the endosymbiotic bacterium Wolbachia pipientis. The mitotic karyotype and detailed photographicmaps of the salivary gland polytene chromosomes of this pest species are presented here. The mitotic metaphase comple-ment consists of 6 pairs of chromosomes, including one pair of heteromorphic sex chromosomes, with the male being theheterogametic sex. The analysis of the salivary gland polytene complement has shown a total of 5 long chromosomes (10polytene arms) that correspond to the 5 autosomes of the mitotic nuclei and a heterochromatic mass corresponding to thesex chromosomes. The most prominent landmarks of each polytene chromosome, the ‘‘weak points’’, and the unusualasynapsis of homologous pairs of polytene chromosomes at certain regions of the polytene elements are also presented anddiscussed.

Key words: Rhagoletis cerasi, fruit fly, agricultural pest, polytene chromosomes, mitotic chromosomes, asynapsis, Wolba-chia pipientis.

Resume : La mouche du cerisier, Rhagoletis cerasi, est un insecte nuisible important, mais au sujet duquel peu d’informa-tions biologique, genetique et cytogenetique sont disponibles. Les auteurs rapportent ici une analyse cytogenetique de qua-tre populations grecques naturelles du R. cerasi, lesquelles sont toutes infectees par la bacterie endosymbiotiqueWolbachia pipientis. Un caryotype mitotique ainsi que des cartes photographiques detaillees des chromosomes polytenesdes glandes salivaires de cette espece sont presentes. Le complement mitotique en metaphase est constitue de six paires dechromosomes incluant une paire de chromosomes sexuels heteromorphes, le sexe male etant heterogametique. L’examendes chromosomes polytenes des glandes salivaires a revele la presence de cinq longs chromosomes (10 bras polytenes) quicorrespondent aux cinq autosomes des noyaux mitotiques et d’une masse heterochromatique correspondant aux chromoso-mes sexuels. Les points de repere les plus saillants de chaque chromosome polytene, les « points faibles » ainsi que l’asy-napsis inhabituelle de paires d’homologues de chromosomes polytenes a certains endroits des elements polytenes sontegalement presentes et discutes.

Mots-cles : Rhagoletis cerasi, mouche du cerisier, insecte nuisible, chromosomes polytenes, chromosomes mitotiques, Wol-bachia pipientis.

[Traduit par la Redaction]

Introduction

Cytogenetic analysis of Diptera has been greatly facili-tated by the existence of polytene chromosomes. Since thefirst publication of Bridges’ hand-drawn chromosome maps(Bridges 1935), polytene chromosomes have proven to beexcellent experimental material for studying chromosomestructure and function, temporal gene activities, genomicorganization, and phylogenetic relationships among species(for a recent review see Zhimulev et al. 2004 and references

therein). They also provide means for the construction ofdetailed genetic-cytogenetic maps through accurate mappingof chromosome rearrangements and precise localization ofgenes by in situ hybridization (Pardue and Gall 1975).

The European cherry fruit fly, Rhagoletis cerasi, a speciesbelonging to the family Tephritidae of the order Diptera, is amajor agricultural pest. It is considered rather oligophagous,ovipositing and breeding on fruits of sweet cherries (Prunusavium), sour cherries (P. cerasus), and honeysuckle (Loni-cera spp., mainly L. tartarica and L. xylosteum). The fly is

Received 29 August 2007. Accepted 23 November 2007. Published on the NRC Research Press Web site at genome.nrc.ca on 23 May2008.

Corresponding Editor: A. Hilliker.

I. Kounatidis and P. Mavragani-Tsipidou.1 Department of Genetics, Development and Molecular Biology, School of Biology, Facultyof Science, Aristotle University of Thessaloniki, GR-54124 Thessaloniki, Greece.N. Papadopoulos. Department of Agriculture, Crop Production and Rural Environment, University of Thessaly, Phytokou Street,38446 N. Ionia (Volos), Magnisia, Greece.K. Bourtzis. Department of Environmental and Natural Resources, University of Ioannina, 30100 Agrinio, Greece.

1Corresponding author (e-mail: mavragan@bio.auth.gr).

479

Genome 51: 479–491 (2008) doi:10.1139/G08-032 # 2008 NRC Canada

distributed in almost all European countries as well asseveral Mediterranean and some Asian countries (Fimiani1989; White and Elson-Harris 1992). It is univoltine andundergoes obligatory pupal diapause. In nature, adultsemerge at the end of spring or the beginning of summer, de-pending on the particular climate of the area. The femaleusually oviposits a single egg into each ripe or semi-ripecherry fruit, and lifetime fecundity ranges from 80 to 200eggs (Katsoyannos 1979). Adults feed on nectar, honeydew,and other sources of liquid or semiliquid food, while larvaedevelop and grow exclusively in the mesocarp of the fruits.Fruit mesocarp is destroyed by larval activity and secondaryinfections of fungi and bacteria. After completing develop-ment, larvae drop off the fruit, fall on the ground, andpupate in the soil.

Although there is considerable interest in the biology ofR. cerasi (Boller and Prokopy 1976; Boller et al. 1998;Riegler and Stauffer 2002; Schwarz et al. 2003; Kovanciand Kovanci 2006), no genetic or cytogenetic data exist onthis pest. The only report on the cytology of R. cerasi refersto the mitotic karyotype of this species (Bush and Boller1977). In the present study, we present the mitotic karyotypeand detailed photographic maps of salivary gland polytenechromosomes of R. cerasi, which could be used as referencematerial for future studies dealing with the population struc-ture of this pest and its control.

Materials and methodsRhagoletis cerasi larvae and adults are available only for

a period of approximately 2 months per year during thecherry fruit ripening period. During the rest of the year,R. cerasi undergoes obligatory pupal diapause. Moreover,artificial rearing of this insect is very difficult and there areno laboratory-adapted populations.

FliesRhagoletis cerasi larvae and pupae used in the present

study were obtained from field-infested cherries collectedfrom 4 different regions of Greece (Katerini, Kavala, Thes-saloniki, and Volos) during May and June of 2005 and 2006.

Detection of Wolbachia pipientis infectionDNA was extracted using the STE protocol (O’Neill et al.

1992). Infection by Wolbachia pipientis (hereafter, Wolba-chia) was detected by PCR using the primers 99F and994R, which are specific for Wolbachia 16S rDNA andyield a product of about 900 bp (O’Neill et al. 1992), andthe primers 81F and 691R, which target the wsp gene andyield a product of about 600 bp (Braig et al. 1998; Zhou etal. 1998). PCR results obtained with the 16S rDNA and wspprimers were in complete agreement. Control PCR wasperformed to test the quality of the DNA template using themitochondrial 12S rDNA primers 12SCRF and 12SCRR,which yield a 350 bp product (Klossa-Kilia et al. 2006).One microlitre of DNA extract (from a total of 50 mL) wasused as template for PCR. All PCR analyses were carriedout in 25 mL volumes and involved an initial denaturationstep at 94 8C for 5 min. This was followed by 35 cycles ofdenaturation at 94 8C for 1 min, annealing at 55 8C for1 min, and extension at 72 8C for 1 min, and a final exten-

sion at 72 8C for 10 min. The PCR mixtures included2.5 mmol/L MgCl2, all four dNTPs (each at 250 mmol/L),0.5 mmol/L of each primer, 1 unit of Taq DNA polymerase(Promega), and buffer supplied by the manufacturers. PCRproducts were visualized on 1.2% agarose gels stained withethidium bromide.

Mitotic chromosome preparationSpread chromosome preparations were made from brain

ganglia of third-instar larvae and young pupae following themethod described by Frydrychova and Marec (2002) withsome modifications. Brain tissue was dissected in Ringer’ssolution and incubated in 0.1 mmol/L colchicine for 15–25 min. The material was transferred to hypotonic solution(0.075 mol/L KCl) for 15 min and then fixed for 15 min infreshly prepared Carnoy fixative (ethanol:chloroform:aceticacid, 6:3:1). Samples were then transferred to a small dropof 60% acetic acid and macerated by moving the materialin and out of a micropipette tip several times. Finally, thematerial was placed on a warm hot plate (45 8C) for dryingand then stained in 5% Giemsa in 0.01 mol/L phosphatebuffer. The technique described by Selivon and Perondini(1997) was used for C-banding.

Polytene chromosomesPolytene chromosome preparations from salivary glands

of third-instar larvae and pupae (1–4 days old) were madefollowing the methods described by Mavragani-Tsipidou etal. (1992a) and Zambetaki et al. (1995) with some modifica-tions. The best polytene preparations were obtained frompupae. Pupae were dissected in Ringer’s solution and thepolytene tissue was first transferred to 45% acetic acid for2–3 min and then post-fixed in 1 mol/L HCl for 2 min. Thematerial was passed through lactoacetic acid (80% lacticacid : 60% acetic acid, 1:1) and stained in lactoacetic orceinfor 10–20 min. Excess stain was removed by washing thematerial in a drop of lactoacetic acid before squashing.

Construction of photographic chromosome mapsSelected photographs were taken at 100� magnification

using a computer-controlled Nikon (Eclipse 80i) phase-contrast microscope and Adobe Photoshop software (CS2).

Results

Mitotic chromosomesAs reported by Bush and Boller (1977), Rhagoletis cerasi

has 6 pairs of chromosomes, including one pair of sexchromosomes, with the male being the heterogametic sex(XY). We have confirmed the existence of 6 pairs ofchromosomes, and Fig. 1 shows the chromosomes of femaleand male animals. Following the labeling system used byRadu et al. (1975) for the Mediterranean fruit fly, Ceratitiscapitata, the sex chromosomes are labeled as the first pairof the mitotic karyotype, while the 5 autosomes are namedfrom 2 to 6 in order of descending size. Of the 5 pairs ofautosomes, the longest one (chromosome 2) is metacentric,while the rest are submetacentric. The smallest autosomes,5 and 6, are similar in length. The sex chromosomes, X andY, are very similar in length, making difficult the discrimi-nation of female and male individuals. However, in the

480 Genome Vol. 51, 2008

# 2008 NRC Canada

female mitotic complement, XX chromosomes presentsomatic pairing like the homologous autosomes (Fig. 1a),while in males, X and Y chromosomes are separated fromeach other (Figs. 1b–1d). Chromosome X appears sub-acrocentric, with a very short arm, while Y seems to beacrocentric and slightly longer than the X chromosome(Figs. 1b–1d). Both sex chromosomes are highly heterochro-matic, as shown by the C-banding technique (Fig. 1d).Somatic pairing of the homologous autosomes was detectedin the R. cerasi mitotic complement (Fig. 1).

Polytene chromosomesThe salivary gland polytene chromosomes of 1–4-day-old

pupae were analyzed. The best quality of chromosomes wasobtained when young pupae were used for preparationsinstead of larvae (see above). The pupae were obtainedfrom infested fruits collected from different regions ofGreece (see Materials and methods). The analysis of poly-tene chromosomes from the first population examined (Ka-terini) routinely showed a high number of asynapticregions. To elucidate whether this is a universal property ofthe R. cerasi genome, we expanded our study to other natu-ral populations collected from different regions of Greece(Kavala, Thessaloniki, Volos).

The polytene chromosomes of R. cerasi were not an easymaterial to work with, owing to the poor banding pattern,the tight coiling and twisting of some chromosome arms,the existence of weak points, and the inter- and intra-specificectopic pairing. Moreover, in many cases the heterochro-

matic centromeric regions of some chromosomes wereectopically paired, forming a temporal partial chromocenterthat changed the general picture and led to errors or omis-sions. Fluctuations of chromosome polytenization amongnuclei of the same fly were extremely high. For the bestanalysis of the polytene complement, we used the largestnuclei of each preparation with well-spread chromosomes.

The analysis of the salivary gland polytene chromosomesshowed that the R. cerasi polytene complement consists of atotal of 5 long chromosomes (10 polytene arms) that corre-spond to the 5 autosomes of the mitotic nuclei. There is notypical chromocenter (accumulation of heterochromatin),and this results in separation of the individual chromosomes.The centromeric region of each of the 5 individual chromo-somes is identified by a heterochromatic mass, which char-acterizes each chromosome (Figs. 2a–2e). The photographicchromosome maps, made from the Katerini population, aregiven in Figs. 3 to 7. Chromosomes were numbered from Ito V and divided into sections from 1 to 100, according totheir size. In each polytene chromosome, the longer arm isdesignated as left (L) and the shorter one as right (R). Thislabeling is arbitrary, indicating only the relative size of chro-mosomes, and does not imply any correlation to the mitoticchromosomes.

In all populations of R. cerasi studied, a very high num-ber of asynaptic regions were routinely observed. Most ofthese regions were asynaptic in all polytene complements,and this is the reason they are incorporated in the photo-graphic maps (Figs. 3–7). In the maps, as well as the text,

Fig. 1. Mitotic metaphase chromosomes from nerve ganglia of Rhagoletis cerasi. (a) Female larva, G-banding. (b, c) Male larvae, G-banding.(d) Male larva, C-banding.

Kounatidis et al. 481

# 2008 NRC Canada

we refer to each asynaptic region as As (asynapsis), fol-lowed by the chromosome (I–V) where it is observed, fol-lowed by a letter code (a, b, c, etc.) to denote the different

asynaptic regions observed in each chromosome. The rela-tive degree of asynapsis of these chromosomal regions inthe different populations studied is shown in Table 1. A

Fig. 2. Centromeric regions of the five polytene elements of Rhagoletis cerasi. (a) Chromosome I. (b) Chromosome II. (c) Chromosome III.(d) Chromosome IV. (e) Chromosome V. (f) The heterochromatic mass observed in polytene nuclei. Scale bar represents 5 mm.

Fig. 3. Photographic map of salivary gland polytene chromosome I of Rhagoletis cerasi. L and R indicate the left and right arms of thechromosome and C indicates the centromere. Arrowheads indicate the weak points. Arrows indicate the limits of inverted tandem duplica-tions. As indicates the regions where asynapsis is observed. Scale bar represents 10 mm.

482 Genome Vol. 51, 2008

# 2008 NRC Canada

Fig. 4. Photographic map of salivary gland polytene chromosome II of Rhagoletis cerasi. L and R indicate the left and right arms of thechromosome and C indicates the centromere. Arrowheads indicate the weak points. Arrows indicate the limits of inverted tandem duplica-tions. As indicates the regions where asynapsis is observed. Scale bar represents 10 mm.

Fig. 5. Photographic map of salivary gland polytene chromosome III of Rhagoletis cerasi. L and R indicate the left and right arms of thechromosome and C indicates the centromere. Arrowheads indicate the weak points. As indicates the regions where asynapsis is observed.Scale bar represents 10 mm.

Fig. 6. Photographic map of salivary gland polytene chromosome IV of Rhagoletis cerasi. L and R indicate the left and right arms of thechromosome and C indicates the centromere. Arrowheads indicate the weak points. As indicates the regions where asynapsis is observed.Scale bar represents 10 mm.

Kounatidis et al. 483

# 2008 NRC Canada

number of weak points, which are usually present in thepolytene complement, and the most prominent reverse tan-dem duplications are also shown in the R. cerasi photo-graphic maps (Figs. 3–7).

Chromosome I (sections 1–23) (Fig. 3)Chromosome I is the longest chromosome in the polytene

complement, and it is considered to be the most ‘‘difficult’’one of the entire set. This is mainly due to the very poorbanding pattern, the poor stretching, and the frequency ofbreakages. The entire element is characterized by the hugeand compact heterochromatic mass of the centromeric re-gion (Fig. 2a). Arm IL is recognized by its asynaptic region

1 with the prominent double band at region 1A. This asyn-apsis, As(I)a, seems to include region 1B (Fig. 3). In thesame arm another asynapsis, As(I)b, was found at region7A. The right arm of the chromosome is characterized by anarrow, swollen tip (region 23) followed by a conspicuouspuff in region 22. The most important landmark of this armis region 14, close to the centromere, which in all prepara-tions was found to be asynaptic (As(I)c). It consists of adark band and a faint band in region 14B followed by a suc-cession of many thin dark bands in region 14C (Fig. 3).Since the analysis of the whole chromosome I is very diffi-cult, the frequency of the three asynapses (As(I)a, As(I)b,and As(I)c) found in the different populations studied could

Fig. 7. Photographic map of salivary gland polytene chromosome V of Rhagoletis cerasi. L and R indicate the left and right arms of thechromosome and C indicates the centromere. Arrowheads indicate the weak points. As indicates the regions where asynapsis is observed.Scale bar represents 10 mm.

Table 1. Percentage of individuals in each of four natural Rhagoletis cerasi popula-tions that displayed asynapses in specific polytene chromosome regions.

Percentage of individuals

Asynapsis Chromosome region Katerini Kavala Thessaloniki VolosAs(I)a 1 * * * *As(I)b 7 * * * *As(I)c 14 * * * *As(II)a 24 100 100 100 100As(II)b 26 87.5 66.6 57.1 75As(II)c 27 100 100 100 100As(II)d 41 75 66.6 100 75As(II)e 44 100 100 100 100As(III)a 58 87.5 33.3 71.4 25As(III)b 62 87.5 100 85.7 100As(III)c 64 * * * *As(IV)a 80 100 100 100 100As(IV)b 81 100 100 100 100As(IV)c 83 100 100 100 100As(V)a 84 100 100 100 100As(V)b 98 100 100 100 100As(V)c 100 100 100 100 100n 24 18 21 24

Note: Each asynaptic region is referred to as As followed by the chromosome (I–V) where itis observed and a letter code (a, b, c, etc.) to denote the different asynaptic regions observed ineach chromosome. n, number of individuals tested. *, percentage could not be estimated; fordetails see text.

484 Genome Vol. 51, 2008

# 2008 NRC Canada

not be estimated. Therefore, they are not given in Table 1.Regions 16 and 19–20 of the right arm were frequentlyfound in a paired condition (Fig. 3), which constitutes evi-dence for the existence of inverted tandem duplications.

Chromosome II (sections 24–44) (Fig. 4)Chromosome II is easily identified because of its charac-

teristic tips. The telomeric regions 24 and 44 of arms IILand IIR, respectively, are always asynaptic with an easilyrecognizable banding pattern (Figs. 4 and 8). Three asynap-tic regions were found on the left arm (As(II)a at region 24,As(II)b at region 26, and As(II)c at region 27) and two werefound on the right arm (As(II)d at region 41 and As(II)e atregion 44) (Figs. 4 and 8). The relative frequencies of thefive asynaptic regions in the different populations studiedare given in Table 1. The frequency of three of them(As(II)a, As(II)c, and As(II)e) was 100% in all populationsstudied. Even though the homology of the maternal and pa-ternal strands seems to be evident in some asynaptic regions(As(II)e and As(II)c) (Figs. 4, 8e–8h), differences betweenthe banding patterns of the parent strands were observed atAs(II)a (Figs. 8a–8d). Indeed, although the length and thebanding pattern of parent strands seem to be the same in75% of the preparations (Fig. 8a), in 25% of the prepara-

tions differences regarding the length and the banding pat-tern of maternal and paternal strands were evident(Figs. 8b–8d). Region 34 of arm IIL was frequently foundin a paired condition (Fig. 4), which constitutes evidencefor the existence of inverted tandem duplication.

Chromosome III (sections 45–64) (Fig. 5)The analysis of the left arm of this chromosome is very

difficult, mainly due to the poor banding pattern, the tightcoiling, and the breakages, especially at the acrocentric area(regions 45–48). The right arm, however, is easily recog-nized by its characteristic tip, the well-banded pattern, andthe characteristic puff at region 59 followed by many darkbands in region 58. Three asynaptic regions (As(III)a,As(III)b, and As(III)c) were found on this arm, at regions58, 62, and 64, respectively, and their relative frequenciesin the different R. cerasi populations are given in Table 1.Asynapsis As(III)a seems to include the tandem dark bandsin region 58 (Figs. 9b, 9c). However, in most of the prepara-tions, this asynapsis covers the whole region 58–59(Fig. 9d). Asynapsis As(III)c seems to include the telomericregion 64 (Figs. 5 and 10b–10d), while As(III)b includes re-gion 62 (Figs. 5, 10a, 10e, 10h). However, in most prepara-tions, the whole area was found to be asynaptic (Figs. 10c,

Fig. 8. Variations in the appearance of the telomeric unsynapsed regions of chromosome II. (a–d) As(II)a at section 24 of arm IIL. (e–h)As(II)e at section 44 of arm IIR. Scale bar represents 5 mm.

Kounatidis et al. 485

# 2008 NRC Canada

10g). It is difficult to determine the relative frequency of theAs(III)c asynapsis in the different populations studied, sincethe asynapsis in region 62 may cover the whole segment (re-gions 62–64) (Figs. 10c, 10g). In region 62 (As(III)b), an ap-parent deficiency of a segment was observed. Indeed, inabout 25% of the pupae analyzed, a homozygous deficiencyof a segment of many small bands was detected in region62A–62B (Figs. 10c, 10d). In all populations studied, thefrequency of the asynapsis As(III)b was 100% (Table 1).The different puffing patterns of the sister chromatids in re-gion 63B (Figs. 10g, 10h) seem to be interesting.

Chromosome IV (sections 65–83) (Fig. 6)Chromosome IV is characterized by a large amount of

heterochromatin in the centromeric area (Fig. 2d). IVL is adifficult arm, mainly because of the very poor banding pat-tern of regions 65–67. The right arm is easily identified byits flared asynaptic tip at region 83. Three asynaptic regionsare observed in this arm: As(IV)a, As(IV)b, and As(IV)c atregions 80, 81, and 83, respectively. In all populationsstudied, the frequency of these asynaptic regions was 100%(Table 1).

Chromosome V (sections 84–100) (Fig. 7)This chromosome is the most distinctive of the polytene

complement, because of the characteristic tips. Both tips arealways found asynaptic (Fig. 7). The VL tip is easily identi-fied by a pair of dark bands in 84C, followed by two puffs(84C and 84D). The right free end has a very characteristictip with thin bands followed by a bundle of many darkbands (100A). Three asynaptic regions were found, one onarm VL (As(V)a) in region 84 and two on arm VR, As(V)bin region 98 and As(V)c in region 100 (Fig. 7). In all popu-lations studied, the frequency of the three asynapses was100% (Table 1).

Presence of Wolbachia in R. cerasiAll 4 natural populations of R. cerasi used in the present

study were found to be infected with Wolbachia, based on abacteria-specific PCR assay (Fig. 11).

DiscussionThe first cytological data for Rhagoletis cerasi were re-

ported by Bush and Boller (1977). They reported a total of6 pairs of mitotic chromosomes, including a heteromorphicsex chromosome pair, which is consistent with our findings.However, there are differences in chromosome lengths andarm ratios between our data and those of Bush and Boller(1977).

Rhagoletis cerasi has the same number of chromosomes(2n = 12) found in 4 other Rhagoletis species, namelyR. berberidis, R. nova, R. conversa, and R. brincidi (Bushand Boller 1977; Frias 2004), while for R. meigeni a differ-ent number of chromosomes (2n = 8+X) was reported (Bushand Boller 1977). The total number of chromosomes foundin R. cerasi is consistent with the modal number of chromo-some pairs of most of the calyptrate Diptera (Boyes and vanBrink 1965). Indeed, Ceratitis capitata (Bedo 1986;Zacharopoulou 1987), Dacus curcubitae (Singh and Gupta1984), Lucilia cuprina (Foster et al. 1980), and many spe-cies of the genus Bactrocera (Mavragani-Tsipidou et al.1992a; Hunwattanakul and Baimai 1994; Baimai et al.1995, 1996; Zhao et al. 1998; Shahjahan and Yesmin 2002)have a similar number of mitotic chromosomes. However,the relative length and the arm ratio of both autosomes andsex chromosomes are quite different among species, evenamong those that are phylogenetically very close. Indeed,contrary to most of the above species, in which the Y chro-mosome was reported as a very small, dot-like chromosome,in R. cerasi the Y chromosome was found to be longer thanthe X chromosome (Bush and Boller 1977; present study)(Fig. 1). Great variation in the number, morphology, andlength of sex chromosomes has been reported in Rhagoletisspecies (Bush and Boller 1977; Frias 2004).

Somatic pairing of homologous chromosomes was de-tected in the R. cerasi mitotic complement (Fig. 1). In mostorganisms at metaphase of mitosis the chromosomes tend tobe evenly distributed over the plate. In Diptera, however,metaphase mitotic chromosomes tend to adjoin each other(Metz 1916; Frias 2004). Although somatic pairing has beenobserved in other organisms (Jones 1941), the tendency ofsomatic pairing of homologues is characteristic of Diptera.Regarding the R. cerasi mitotic complement, the somatic

Fig. 9. Sections 58–59 of chromosome arm IIIR, where the asynap-sis As(III)a was detected. (a) Tightly synapsed. (b, c) Unsynapsed.(d) Coverage of the whole region. Scale bar represents 5 mm.

486 Genome Vol. 51, 2008

# 2008 NRC Canada

pairing of XX chromosomes facilitated the discrimination offemales and males.

As described in the Results, the R. cerasi polytene com-plement consists of a total of 5 long banded chromosomes(10 polytene arms) that correspond to the 5 autosomes ofthe mitotic nuclei (Figs. 3–7). Chromosome I and the leftarms of chromosomes III and IV were the most ‘‘difficult’’ones of the whole complement. These arms are the onlyones that do not possess extensive asynaptic regions, a veryprominent and characteristic feature of R. cerasi polytenenuclei.

No typical chromocenter like that of Drosophila mela-nogaster exists in the R. cerasi polytene complement. Thisis also the case for C. capitata (Bedo 1986; Zacharopoulou1987, 1990), L. cuprina (Foster et al. 1980), Bactroceraoleae (Mavragani-Tsipidou et al. 1992a), B. tryoni (Zhao etal. 1998), and B. curcubitae (Shahjahan and Yesmin 2002).However, the centromeric regions of R. cerasi chromosomesare easily identified, being a prominent landmark of each in-dividual chromosome. Thus, chromosomes I, IV, and V arecharacterized by large heterochromatic centromeric masses,while the centromeric masses of II and III are of limited

size (Fig. 2). Characteristic centromeric masses, which areprominent landmarks of each polytene chromosome, havebeen found in other Diptera as well (Mavragani-Tsipidou etal. 1992a; Zhao et al. 1998).

The analysis of the polytene nuclei of males and femalesshowed no difference in the number and quality of the poly-tene elements in the different sexes, indicating the absenceof banded sex chromosomes. This observation is consistentwith previous reports on C. capitata (Bedo 1986, 1987;Zacharopoulou 1987, 1990; Bedo and Zacharopoulou 1988),L. cuprina (Childress 1969; Foster et al. 1980; Bedo 1982),Chrysomya bezziana (Bedo 1992), B. oleae (Mavragani-Tsipidou et al. 1992a; Zambetaki et al. 1995), B. tryoni(Zhao et al. 1998), and B. curcubitae (Shahjahan and Yes-min 2002). The heterochromatic mass found in all polytenenuclei of R. cerasi individuals (Fig. 2f) may represent theunderreplicated XX and XY sex chromosomes, as proposedfor other Diptera (Childress 1969; Bedo 1982, 1987; Bedoand Zacharopoulou 1988; Mavragani-Tsipidou et al. 1992a;Shahjahan and Yesmin 2002). This is consistent with thehighly heterochromatic nature of both R. cerasi sex chro-mosomes, as shown by the C-banding technique (Fig. 1d).

Fig. 10. Sections 62–64 of chromosome arm IIIR, where the asynapses As(III)b and As(III)c were detected. (a–h) Variations in the appear-ance of the asynapses As(III)b and As(III)c. (c, d) Deficiency of a segment of many thin bands. (g, h) Different puffing patterns of the sisterchromatids (for details see text). Scale bar represents 5 mm.

Kounatidis et al. 487

# 2008 NRC Canada

No prominent differences in size or density of this masswere detected among the different R. cerasi individuals.This observation could be explained by the equal sizes ofthe sex chromosomes found in the mitotic complement ofthis insect (Fig. 1). However, further analysis of X-linkedsequences and the localization by in situ hybridization ofgenes on both mitotic and polytene chromosomes ofR. cerasi will permit establishment of correlations betweenthese two sets of chromosomal elements and will provideinformation on the fate of the sex chromosomes of themitotic complement.

The study of R. cerasi polytene chromosomes revealed ahigh number of weak points distributed among the chromo-somes (Figs. 3–7). Most of them were found in the ‘‘dif-ficult’’ chromosome arms of R. cerasi. Weak points are theresult of local underreplication of the chromosomes duringpolytenization, which at the cytological level shows up aschromosome breaks (Painter 1934; Lamb and Laird 1987;Moshkin et al. 2001; Belyaeva et al. 2006). Moreover, 3inverted tandem duplications were detected in a paired con-dition in chromosome arms IR and IIL. A similar situationseems to exist in B. oleae (Mavragani-Tsipidou et al.1992a; Zambetaki et al. 1995), Culex pipiens (Zambetaki etal. 1998), and many Drosophila species (Kastritsis et al.1986; Mavragani-Tsipidou et al. 1990, 1992b). In general,fixed duplications in the genome, regardless of their origin,may be important chromosomal aberrations in the context ofevolution (Ohno 1970).

One of the most striking features of the R. cerasi polytenechromosomes is the asynapsis detected in several regions ofmost chromosome arms (Figs. 4–7). As mentioned in theResults, the analysis of polytene nuclei from differentR. cerasi populations was undertaken to find the extent ofasynapsis in different populations. Seventeen prominentasynapses were detected in the 4 populations studied(Figs. 4–7). Nine of these unsynapsed regions were found ata frequency of 100% in all populations studied, while for therest of them, the frequency varied among the differentpopulations (Table 1).

The phenomenon of somatic synapsis in the polytenechromosomes of Diptera is an excellent tool for cytologicalanalysis. When synapsis is complete, both homologous chro-

matids synapse band by band, resulting in a single chromo-some. When synapsis is incomplete, however, the contact isrestricted to some of the regions, and the homologues arepartly separated. It has been reported in different studiesthat during normal development of D. melanogaster, the fre-quency of salivary gland nuclei showing partial synapsis inany of the chromosomes varies between 6.5% and 45%(Zhimulev 1996; Zhimulev et al. 2004). However, this isnot the case for the R. cerasi polytene complement, sinceunsynapsed regions were detected in all nuclei of the samepreparation in relaxed as well as greatly stretched chromo-somes.

Asynapsis is frequently observed in hybrid chromosomesof some inter- and intra-specific crosses (e.g., Machado etal. 2006), as well as in the presence of rearrangements suchas deletions, duplications, inversions, and translocations(Lefevre 1976). In the case of R. cerasi, no heterozygoticinversions or translocations were found in the polytenechromosome slides obtained from the 4 populationsstudied. The possibility of very short inversions that couldexplain the existence of asynapsis in certain regions can beexcluded, since we never found these regions tightly syn-apsed (Lefevre 1976). On the other hand, short deficienciesor additions of genetic material in one of the parentalstrands may explain the constant appearance of these asyn-aptic regions in R. cerasi. The homology of the maternaland paternal strands in many of the constantly observedasynapses (such as As(II)b, As(II)c, As(II)e, As(IV)c,As(V)b, and As(V)c) is evident (Figs. 4, 6, 7, 8e–8h). How-ever, even in these cases, differences in the length of theinterbands or even in the density of bands are obvious(Figs. 8, 9, 10), suggesting the existence of microdeletionsor microduplications, which are virtually undetectable byconventional cytology. In other cases, such as asynapsesAs(II)a (Figs. 8a–8d) and As(III)b (Fig. 10), the differencesare very prominent. In the case of As(II)a, one sister chro-matid is longer, containing a number of additional thinbands at the telomeric region (Figs. 8b–8d), while in armIIIR, a deficiency of at least 4 thin bands was observed(Figs. 10c–10d).

The prominent difference observed in the puffing patternsof the sister chromatids in the asynaptic area 62–64(Figs. 10g–10h) suggests differences in the functions of thesister chromatids. It is generally accepted that the puffs ofpolytene chromosomes are indicators of gene activity(Clever 1964; Ashburner et al. 1974; Lezzi and Richards1989; see also the review by Zhimulev et al. 2004 and refer-ences therein). This correlation between puffs and gene acti-vation was an early demonstration of the link betweentranscription and changes in chromatin structure (Udvardyet al. 1985). Since a puff may be the result of even a singleband, an insertion or deletion of one or more bands in oneof the sister chromatids may explain the differences in thepuffing patterns of the R. cerasi parent strands.

The 4 natural populations of R. cerasi used in the presentstudy were found to be infected with the endosymbiotic bac-terium Wolbachia (Fig. 11). Could Wolbachia infections beassociated with the phenomenon of asynapsis? Wolbachia isa group of gram-negative, obligatory intracellular, andmaternally transmitted endosymbionts that have beendetected in a wide range of arthropod species including all

Fig. 11. PCR detection of Wolbachia pipientis, using primers (99Fand 994R) specific for Wolbachia 16S rDNA, in five male and fivefemale Rhagoletis cerasi individuals from the Volos population. m,male individuals; f, female individuals.

488 Genome Vol. 51, 2008

# 2008 NRC Canada

major orders of insects (Bourtzis and O’Neill 1998; Bourtzisand Braig 1999). They are usually present in the reproduc-tive tissues of their hosts and have been associated withreproductive alterations such as cytoplasmic incompatibility,parthenogenesis, feminization, and male killing (Bourtzisand Miller 2003, 2006; Veneti et al. 2004). Their presencein developing gametes is the reason for lateral gene transferof bacterial genes to insect host genomes (Kurland 2000).Wolbachia–host transfer events were first described in thebean beetle Callosobruchus chinensis and the filarial nema-tode Oncocerca sp. (Kondo et al. 2002; Fenn et al. 2006).Recently, additional evidence was reported demonstratingthat Wolbachia DNA insertions are present in the genomesof the filarial nematodes Brugia malayi, B. timori,B. pahangi, and Dirofilaria immitis; the parasitoid waspsNasonia vitripennis, N. giraulti, and N. longicornis; themosquito species Culex pipiens; and Drosophila simulans,D. sechellia, and D. ananassae (Dunning Hotopp et al.2007). In one case, nearly the entire bacterial genome(>1 Mbp) was transferred from a Wolbachia strain to thehost, D. ananassae (Dunning Hotopp et al. 2007). Based onthis evidence, one could speculate that similar transferevents could have taken place in the Wolbachia–R. cerasisymbiotic association. Such lateral gene transfers couldcause the large number of asynaptic phenomena observed inthe polytene chromosomes of R. cerasi. It is also noteworthythat similar asynaptic phenomena were observed in the poly-tene chromosomes of Culex species, where Wolbachia genetransfer events are also documented (Zambetaki et al. 1998;Dunning Hotopp et al. 2007). Whether gene transfer eventshave occurred between Wolbachia and the cherry fruit flyand are associated with asynaptic phenomena is currentlyunder investigation.

AcknowledgmentsThis study was co-funded by the European Social and

National Resources – EPEAEK II – Pythagoras, and theGreek ministry of education. The present paper is dedicatedto the memory of Professor Costas D. Kastritsis.

ReferencesAshburner, M., Chihara, C., Meltzer, P., and Richards, G. 1974.

Temporal control of puffing activity in polytene chromosomes.Cold Spring Harb. Symp. Quant. Biol. 38: 655–662. PMID:4208797.

Baimai, V., Trinachartvanit, W., Tigvattananont, S., Grote, P.J.,Poramarcom, R., and Kijechalao, U. 1995. Metaphase karyo-types of fruit flies of Thailand. I. Five sibling species of theBactrocera dorsalis complex (Diptera: Tephritidae). Genome,38: 1015–1022. doi:10.1139/g95-134. PMID:8536995.

Baimai, V., Trinachartvanit, W., Tigvattananont, S., and Grote, P.J.1996. Metaphase karyotypes of fruit flies of Thailand. II. Fivesibling species in four subgenera of the Bactrocera dorsaliscomplex. J. Sci. Soc. Thail. 22: 97–104.

Bedo, D.G. 1982. Differential sex chromosome replication and do-sage compensation in polytene trichogen cells of Lucilia cuprina(Diptera: Calliphoridae). Chromosoma, 87: 21–32. doi:10.1007/BF00333507. PMID:7160277.

Bedo, D.G. 1986. Polytene and mitotic chromosome analysis inCeratitis capitata (Diptera; Tephritidae). Can. J. Genet. Cytol.28: 180–188. doi:10.1139/g86-025.

Bedo, D.G. 1987. Polytene chromosome mapping in Ceratitis capi-tata (Diptera: Tephritidae). Genome, 29: 598–611. doi:10.1139/g87-101.

Bedo, D.G. 1992. Polytene chromosomes of the Old World screw-worm fly (Chrysomya bezziana) and its evolutionary relation-ships with Lucilia cuprina and Cochiliomyia hominivorax(Diptera: Calliphoridae). Genome, 35: 294–303. doi:10.1139/g92-045. PMID:1618389.

Bedo, D.G., and Zacharopoulou, A. 1988. Intertissue variability ofpolytene chromosome banding patterns. Trends Genet. 4: 90–91.doi:10.1016/0168-9525(88)90093-5. PMID:3238761.

Belyaeva, E.S., Demakov, S.A., Pokholkova, G.V., Alekseyenko,A.A., Kolesnikova, T.D., and Zhimulev, I.F. 2006. DNA under-replication in intercalary heterochromatin regions in polytenechromosomes of Drosophila melanogaster correlates with the for-mation of partial chromosomal aberrations and ectopic pairing.Chromosoma, 115: 355–366. doi:10.1007/s00412-006-0063-7.PMID:16583218.

Boller, E.F., and Prokopy, R.J. 1976. Bionomics and managementof Rhagoletis. Annu. Rev. Entomol. 21: 223–246. doi:10.1146/annurev.en.21.010176.001255.

Boller, E.F., Katsoyannos, B.I., and Hippe, C. 1998. Host races ofRhagoletis cerasi L. (Dipt., Tephritidae): effect of prior adultexperience on oviposition site preference. J. Appl. Entomol.122: 231–237.

Bourtzis, K., and Braig, H.R. 1999. The many faces of Wolbachia.In Rickettsiae and rickettsial diseases at the turn of the thirdmillennium. Edited by D. Raoult and P. Brouqui. Elsevier, Paris.pp. 199–219.

Bourtzis, K., and Miller, T.A. (Editors). 2003. Insect symbiosis.CRC Press, Boca Raton, Fla.

Bourtzis, K., and Miller, T.A. (Editors). 2006. Insect symbiosis.Vol. 2. CRC Press, Boca Raton, Fla.

Bourtzis, K., and O’Neill, S.L. 1998. Wolbachia infections andtheir influence on arthropod reproduction. Bioscience, 48: 287–293. doi:10.2307/1313355.

Boyes, J.W., and van Brink, J.M. 1965. Chromosomes of calyptrateDiptera. Can. J. Genet. Cytol. 7: 537–550. doi:10.1139/g65-073.

Braig, H.R., Zhou, W., Dobson, S.L., and O’Neill, S.L. 1998. Clon-ing and characterization of a gene encoding the major surfaceprotein of the bacterial endosymbiont Wolbachia pipientis. J.Bacteriol. 180: 2373–2378. PMID:9573188.

Bridges, C.B. 1935. Salivary chromosome maps with a key to thebanding of the chromosomes of Drosophila melanogaster. J.Hered. 26: 60–64.

Bush, G.L., and Boller, F. 1977. Chromosome morphology ofRhagoletis cerasi species complex (Diptera, Tephritidae). Ann.Entomol. Soc. Am. 70: 316–318.

Childress, D. 1969. Polytene chromosomes and linkage group-chromosome correlations in the Australian sheep blowflyLucilia cuprina (Diptera: Calliphoridae). Chromosoma, 26:208–214. doi:10.1007/BF00326456. PMID:5800665.

Clever, U. 1964. Actinomycin and puromycin: effect on sequentialgene activation by ecdysone. Science (Washington, D.C.), 146:794–795. doi:10.1126/science.146.3645.794. PMID:14197575.

Dunning Hotopp, J.C., Clark, M.E., Oliveira, D.C.S.G., Foster,J.M., Fischer, P., Munoz Torres, M.C., et al. 2007. Widespreadlateral gene transfer from intracellular bacteria to multicellulareukaryotes. Science (Washington, D.C.), 317: 1753–1756.doi:10.1126/science.1142490. PMID:17761848.

Fenn, K., Conlon, C., Jones, M., Quail, M.A., Holroyd, N.E., Park-hill, J., and Blaxter, M. 2006. Phylogenetic relationships of theWolbachia of nematodes and arthropods. PLoS Pathog. 2: e94.doi:10.1371/journal.ppat.0020094. PMID:17040125.

Kounatidis et al. 489

# 2008 NRC Canada

Fimiani, P. 1989. Mediterranean region. In Fruit flies: their biol-ogy, natural enemies and control. Edited by A.S. Robinson andG. Hopper. Elsevier, Amsterdam, the Netherlands. pp. 39–50.

Foster, G.G., Whitten, M.J., Konovalov, C., Bedo, D.G., Maddern,R.H., and Boon, D.T. 1980. Cytogenetic studies of Lucilia cu-prina dorsalis R.-D. (Diptera: Calliphoridae). Polytene chromo-some maps of the autosomes and cytogenetic localization ofvisible genetic markers. Chromosoma, 81: 151–168. doi:10.1007/BF00285946.

Frias, D. 2004. Importance of larval morphology and heterochro-matic variation in the identification and evolution of sibling spe-cies in the genus Rhagoletis (Diptera: Tephritidae) in Chile. InProceedings of the 6th International Symposium on Fruit Fliesof Economic Importance, Stellenbosch, South Africa, 6–10 May2002. Edited by B.N. Barnes. Isteg Scientific Publications, Irene,South Africa. pp. 267–276.

Frydrychova, R., and Marec, F. 2002. Repeated losses of TTAGGtelomere repeats in evolution of beetles (Coleoptera). Genetica,115: 179–187. doi:10.1023/A:1020175912128. PMID:12403172.

Hunwattanakul, N., and Baimai, V. 1994. Mitotic karyotype of fourspecies of fruit flies (Bactrocera) in Thailand. Kasetsart J. (Nat.Sci.), 28: 142–148.

Jones, D.F. 1941. Somatic segregation. Bot. Rev. 7: 291–307.Kastritsis, C.D., Scouras, Z.G., and Ashburner, M. 1986. Duplica-

tions in the polytene chromosomes of Drosophila auraria. Chro-mosoma, 93: 381–385. doi:10.1007/BF00327598.

Katsoyannos, B.I. 1979. Zum Reproduktions und Wirtswahlverhal-ten der Kirschenfliege, Rhagoletis cerasi L. (Diptera: Tephriti-dae). Doctoral dissertation, Eth Zurich, Nr. 6409.

Klossa-Kilia, E., Kilias, G., Tryfonopoulos, G., Koukou, K.,Sfenthourakis, S., and Parmakelis, A. 2006. Molecular phylo-geny of the Greek populations of the genus Ligidium (Isopoda,Oniscidae) using three mtDNA gene segments. Zool. Scr. 35:459–472. doi:10.1111/j.1463-6409.2006.00243.x.

Kondo, N., Nikoh, N., Ijichi, N., Shimada, M., and Fukatsu, T. 2002.Genome fragment of Wolbachia endosymbiont transferred to Xchromosome of host insect. Proc. Natl. Acad. Sci. U.S.A. 99:14280–14285. doi:10.1073/pnas.222228199. PMID:12386340.

Kovanci, O.B., and Kovanci, B. 2006. Effect of altitude on seaso-nal flight activity of Rhagoletis cerasi flies (Diptera: Tephriti-dae). Bull. Entomol. Res. 96: 345–351. PMID:16923201.

Kurland, C.G. 2000. Something for everyone. Horizontal genetransfer in evolution. EMBO Rep. 1: 92–95. doi:10.1093/embo-reports/kvd042. PMID:11265763.

Lamb, M.M., and Laird, C.D. 1987. Three euchromatic DNA se-quences under-replicated in polytene chromosomes of Drosophilaare localized in constrictions and ectopic fibers. Chromosoma, 95:227–235. doi:10.1007/BF00294779. PMID:3113852.

Lefevre, G. 1976. A photographic representation and interpretationof the polytene chromosomes of Drosophila melanogaster sali-vary glands. In The genetics and biology of Drosophila.Vol. 1a. Edited by M. Ashburner and E. Novitski. AcademicPress, London. pp. 31–66.

Lezzi, M., and Richards, G. 1989. Salivary glands. In Ecdysone:from chemistry to mode of action. Edited by J. Koolman. GeorgThieme Verlag, Stuttgart. pp. 393–406.

Machado, L.P.B., Madi-Ravazzi, L., and Tadei, W.J. 2006. Repro-ductive relationships and degree of synapsis in the polytenechromosomes of the Drosophila buzzatii species cluster. Braz.J. Biol. 66(1b): 279–293. PMID:16710520.

Mavragani-Tsipidou, P., Kyrpides, N., and Scouras, Z.G. 1990.Evolutionary implications of duplications and Balbiani rings inDrosophila. A study of Drosophila serrata. Genome, 33: 478–485. doi:10.1139/g90-071. PMID:2227403.

Mavragani-Tsipidou, P., Karamanlidou, G., Zacharopoulou, A.,Koliais, S., and Kastritsis, C.D. 1992a. Mitotic and polytenechromosome analysis in Dacus oleae (Diptera: Tephritidae).Genome, 35: 373–378. doi:10.1139/g92-056. PMID:1624130.

Mavragani-Tsipidou, P., Scouras, Z.G., and Natsiou-Voziki, A.1992b. The Balbiani ring and the polytene chromosomes of Dro-sophila bicornuta. Genome, 35: 64–67. doi:10.1139/g92-011.PMID:1572529.

Metz, C.W. 1916. Chromosome studies on Diptera. II. The pairedassociation of chromosomes in Diptera and its significance. J.Exp. Zool. 21: 213–280. doi:10.1002/jez.1400210204.

Moshkin, Y.M., Alekseyenko, A., Semeshin, F.V., Spierer, A.,Spierer, P., Makarevich, G.F., et al. 2001. The Bithorax Com-plex of Drosophila melanogaster: underreplication and morphol-ogy in polytene chromosomes. Proc. Natl. Acad. Sci. U.S.A. 98:570–574. doi:10.1073/pnas.021353598. PMID:11136231.

Ohno, S. 1970. Evolution by gene duplication. Springer-Verlag,New York.

O’Neill, S.L., Giordano, R., Colbert, A.M.E., Karr, T.L., andRobertson, H.M. 1992. 16S rRNA phylogenetic analysis of thebacterial endosymbionts associated with cytoplasmic incom-patibility in insects. Proc. Natl. Acad. Sci. U.S.A. 89: 2699–2702. doi:10.1073/pnas.89.7.2699. PMID:1557375.

Painter, T.S. 1934. Salivary gland chromosomes and the attack onthe gene. J. Hered. 25: 465–476.

Pardue, M.L., and Gall, J.G. 1975. Nucleic acid hybridization to theDNA of cytological preparations. Methods Cell Biol. 10: 1–16.PMID:1102840.

Radu, M., Rossler, Y., and Koltin, Y. 1975. The chromosomes ofthe Mediterranean fruit fly Ceratitis capitata (Wied): karyotypeand chromosomal organization. Cytologia (Tokyo), 40: 823–828.

Riegler, M., and Stauffer, C. 2002. Wolbachia infections and super-infections in cytoplasmically incompatible populations of theEuropean cherry fruit fly Rhagoletis cerasi (Diptera, Tephriti-dae). Mol. Ecol. 11: 2425–2434. doi:10.1046/j.1365-294X.2002.01614.x. PMID:12406252.

Schwarz, D., McPheron, B.A., Hartl, G.B., Boller, E.F., and Hoff-meister, T.S. 2003. A second case of genetic host races in Rha-goletis? A population genetic comparison of sympatric hostpopulations in the European cherry fruit fly, Rhagoletis cerasi.Entomol. Exp. Appl. 108: 11–17. doi:10.1046/j.1570-7458.2003.00065.x.

Selivon, D., and Perondini, A.L.P. 1997. Evaluation of techniquesfor C and ASG banding of the mitotic chromosomes of Anastre-pha species (Diptera, Tephritidae). Braz. J. Genet. 20: 651–653.

Shahjahan, R.M., and Yesmin, F. 2002. Polytene chromosome mapsof the melon fly Bactrocera curcubitae (Diptera, Tephritidae).Genome, 45: 1167–1174. doi:10.1139/g02-081. PMID:12502263.

Singh, O.P., and Gupta, J.P. 1984. Studies on mitotic and salivarychromosomes of Dacus curcubitae Coquilett (Diptera, Tephriti-dae). Genetica, 62: 217–221. doi:10.1007/BF00056439.

Udvardy, A., Maine, E., and Schedl, P. 1985. The 87A7 chromo-mere. Identification of novel chromatin structures flanking theheat shock locus that may define the boundaries of higher orderdomains. J. Mol. Biol. 185: 341–358. doi:10.1016/0022-2836(85)90408-5. PMID:2997449.

Veneti, Z., Clark, M.E., Karr, T.L., Savakis, C., and Bourtzis, K.2004. Heads or tails: host–parasite interactions in theDrosophila–Wolbachia system. Appl. Environ. Microbiol. 70:5366–5372. doi:10.1128/AEM.70.9.5366-5372.2004. PMID:15345422.

White, I.M., and Elson-Harris, M.M. 1992. Fruit flies of economicsignificance: their identification and bionomics. CAB Interna-tional, Wallingford, UK.

490 Genome Vol. 51, 2008

# 2008 NRC Canada

Zacharopoulou, A. 1987. Cytogenetic analysis of mitotic and sali-vary gland chromosomes in the medfly Ceratitis capitata. Gen-ome, 29: 67–71. doi:10.1139/g87-011.

Zacharopoulou, A. 1990. Polytene chromosome maps in the medflyCeratitis capitata. Genome, 33: 184–197. doi:10.1139/g90-030.

Zambetaki, A., Kleanthous, K., and Mavragani-Tsipidou, P. 1995.Cytogenetic analysis of Malpighian tubule and salivary glandpolytene chromosomes of Bactrocera oleae (Dacus oleae) (Dip-tera: Tephritidae). Genome, 38: 1070–1081. doi:10.1139/g95-143.

Zambetaki, A., Pasteur, N., and Mavragani-Tsipidou, P. 1998. Cy-togenetic analysis of Malpighian tubule polytene chromosomesof Culex pipiens (Diptera: Culicidae). Genome, 41: 751–755.doi:10.1139/gen-41-6-751.

Zhao, J.T., Frommer, M., Sved, J.A., and Zacharopoulou, A. 1998.

Mitotic and polytene analyses in the Queensland fruit fly, Bac-trocera tryoni (Diptera: Tephritidae). Genome, 41: 510–526.doi:10.1139/gen-41-4-510. PMID:9796100.

Zhimulev, I.F. 1996. Morphology and structure of polytene chro-mosomes. Adv. Genet. 34: 1–497. PMID:9348397.

Zhimulev, I.F., Belayaeva, E.S., Semeshin, V.F., Koryakov, D.E.,Demakov, S.A., Demakova, O.V., et al. 2004. Polytene chromo-somes: 70 years of genetic research. Int. Rev. Cytol. 241: 203–275. PMID:15548421.

Zhou, W., Rousset, F., and O’Neill, S. 1998. Phylogeny and PCR-based classification of Wolbachia strains using wsp gene se-quences. Proc. R. Soc. Lond. B Biol. Sci. 265: 509–515.PMID:9569669.

Kounatidis et al. 491

# 2008 NRC Canada