Drosophila: A Guide to Species Identification and Use

Preface

When you say “Drosophila” to most people, they think of Drosophila melanogaster, the laboratory workhorse that for nearly 100 years has been the premier genetic model system in biology. Many remember the smell of ether while handling flies in their undergraduate biology classes, while others recall curly wings and white eyes and Punnet squares. T. H. Morgan and his students C. Bridges, H. J. Muller, and A. H. Sturtevant pioneered the field of Drosophila genetics during the first half of the twentieth century. Their work has been carried on by countless “drosophilists” and, in some ways, culminated with the publication of the full genome sequence of Drosophila melanogaster in 2000. Now Drosophila biology is entering a new era. The genome of a second species, Drosophila pseudoobscura, has recently been completed and more are surely to come. Increasingly, when researchers state that they work on Drosophila they are met with the following question: “What species?” Scientists in the fields of ecology and evolutionary biology are beginning to co-opt various members of the genus Drosophila to serve as a model system for their own research, largely because of the ease of obtaining molecular markers from these species. As we were preparing this book for publication, the National Human Genome Research Institute approved whole genome sequencing of 10 additional Drosophila species, and the creation of BAC libraries for 20. We feel that this initiative will invigorate Drosophila research for the next 100 years. The genus Drosophila represents an unprecedented model system not only for understanding genome evolution, but also for comparative experimental research. No other group has such a well-defined phylogeny and an extensive literature on genetics, development, neurobiology and behavior, physiology, and ecology. A. H. Sturtevant, one of the pioneers of Drosophila genetics, was clearly aware of the importance of the evolutionary context in which D. melanogaster is embedded. He described many new Drosophila species, studied their behavior and genetic relationships, and published his 1921 monograph, The North American Species of Drosophila. A number of excellent resources exist in which the primary literature concerning the distribution, evolutionary relationships, and ecology for most of the known Drosophila species is summarized. In 1952, J. T. Patterson and W. Stone published their still indispensable Evolution in the Genus Drosophila, which, while drawing heavily on work generated from their own activities and those of their students, provided the first overview of the evolutionary relationships and distributions of all known Drosophila species. The international Drosophila community has since contributed its expertise to five volumes of the The Genetics and Biology of Drosophila series, edited by Ashburner, Carson and Thompson between 1981 and 1986. Michael Ashburner (1990, 2004) and Jeff Powell (1997) have both compiled encyclopedic works that effectively summarize decades of basic Drosophila research. A wide range

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Preface

of data for many Drosophila species now can be accessed through web-based resources such as Flybase and Taxodros. As the number of investigators taking advantage of Drosophila diversity grows, the need for a portable resource for identifying and using the different species becomes increasingly critical. In 2001, the Tucson Stock Center began offering the now annual Drosophila Species Identification Workshop to teach basic approaches to keying out flies and maintaining non-melanogaster species in laboratory culture. The overwhelming interest of the research community in attending this workshop and in becoming comfortable with using species in addition to D. melanogaster has led us to create this guide. We have had the amazing good fortune of having the participation in these workshops of many colleagues who were responsible for the original collections and descriptions of diverse Drosophila species: Wyatt Anderson, Michael Ashburner, Hampton Carson, David Foote, Nicolas Gompel, William Heed, Kenneth Kaneshiro, Thom Kaufman, Margaret Kidwell, Kathy Matthews, Bryant McAllister, Stephen Schaeffer, Valerie Schaworoch, Marvin Wasserman, and Marshall Wheeler. To all of them we express our gratitude for their unselfish sharing of their time and expertise with us and with the students in the workshops. Activities associated with the annual workshops, including input from the attendees, have been instrumental in shaping the contents of this guide. Space limitations prevent us from including all Drosophila species in this guide. We have chosen to focus upon the several hundred species maintained at the Tucson Stock Center, as these are readily available to the community and the demand for them is increasing. Conditions for the successful rearing of these species have been worked out and the life history differences critical to their meaningful use in comparative experimental work have been well documented for a number of them. We hope that this guide encourages and enables researchers to exploit the wealth of diversity offered by Drosophila in their investigations. Therese Ann Markow Patrick O’Grady

CHAPTER 1

Phylogenetic relationships of Drosophilidae Contents

• • • • •

The origin of the family Drosophilidae Drosophilidae: relationships among genera • Steganinae • Drosophilinae Conclusions Acknowledgements References

The origin of the family Drosophilidae The Drosophilidae is an acalyptrate family in the superfamily Ephydroidea (McAlpine, 1989). This superfamily contains two large families, Ephydridae and Drosophilidae, as well as several smaller families, such as Camilidae, Diastatidae, and Curtonotidae. Throckmorton (1975) suggested that Diastatidae was the closest relative of the Drosophilidae, based largely on the fact that diastatids are saprophagous in leaf mold (Oldroyd, 1964; Hennig, 1965). Even though Okada (1962) suggested that the ancestral drosophilid substrate was bleeding tree sap, Throckmorton (1975) believed that the current diversity of substrates was the result of opportunism centering on the saprophagous leaf-mold habit. Grimaldi (1990) examined the phylogenetic relationships of Ephydroidea using morphological characters. Of the three most parsimonious trees his search recovered, he selected a “preferred phylogeny”. The strict consensus of all three trees gives the more conservative hypothesis (Figure 1.1a). McAlpine (1989) presents an alternative view of evolution in the superfamily Ephydroidea (Figure 1.1b). These two phylogenetic hypotheses differ mainly in the placement of Drosophilidae. The strict consensus of Grimaldi’s (1990) trees is unable to resolve the sister group of the Drosophilidae (his preferred tree favored the Curtonotidae as the sister family of Drosophilidae). McAlpine’s (1989) phylogeny, however, suggests that the Camilidae is the sister clade of Drosophilidae. The exact placement of Drosophilidae remains an open question, as few ephydroid taxa outside of Drosophilidae and a small number of Ephydridae are well known. Throckmorton (1975) placed the origin of the Drosophilidae in the tropics, based primarily on the pan-tropical distribution of the Lissocephala, a group he considered

4

How to look at flies

Drosophilinae Steganinae Camilidae Risidae Ephydridae Diastatidae Campichoetidae (a)

Curtonotidae

(b)

Figure 1.1. (a) Phylogeny of Ephydroidea based on Grimaldi (1990); (b) Phylogeny of Ephydroidea from McAlpine (1989).

to be basal within the Drosophilidae. The fossil genus Electrophortica, described from Baltic amber (Hennig, 1965), suggests that the Drosophilidae predate the Eocene and may be 50 million years old or older.

Drosophilidae: relationships among genera Throckmorton (1962, 1975) was the first to propose a higher-level phylogenetic framework for the family Drosophilidae (Figure 1.2). He proposed a number of radiations, meant to represent multiple speciation events with subsequent diversification, based on morphological characters. There have been two major criticisms of Throckmorton’s work: first, his analyses were not based on any explicit cladistic algorithm and are therefore not repeatable; and second, he did not attempt to maintain any concept of monophyly in his study. As a result, species groups of the genus Drosophila are more closely related to other genera than they are to other species groups in their genera. In spite of these criticisms, it is still useful to review Throckmorton’s work as it agrees quite well with recent phylogenetic analyses based on molecular data (e.g. Remsen and O’Grady, 2002). The basal radiation in this family is the Steganinae radiation, which includes all the members of the subfamily Steganinae. A derivative of this radiation led to the Scaptodrosophila radiation, and the diversification of the basal Drosophilidae species. The Sophophoran radiation led to the present day subgenus Sophophora, and some related genera such as Chymomyza (Figure 1.2). The Drosophila radiation is divided into three major parts: 1. A basal radiation containing the funebris species group and related taxa 2. The virilis-repleta radiation 3. The immigrans-Hirtodrosophila radiation.

Phylogenetic relationships of Drosophilidae

5

Figure 1.2. Genus-level phylogeny of Drosophilidae based on Throckmorton (1975).

The virilis-repleta radiation includes about 20 species groups, most of which breed in rotting plant matter. The immigrans-Hirtodrosophila radiation is further divided into the Old World Hirtodrosophila and tripunctata radiations (Figure 1.2). Grimaldi (1990) proposed a cladistic reclassification of Drosophilidae based on his analysis of 218 morphological characters (Figure 1.3). His analysis suggested that,

6

How to look at flies

Stegana Rhinoleucophenga

STEGANINAE

Gitona Amiota Scaptodrosophila Chymomyza Paramycodrosophila Hirtodrosophila

Zygothrica Genus Group

Zygothrica Mycodrosophila Hawaiian Drosophila = Idiomyia crassifemur Continental Scaptomyza

Scaptomyza Genus Group

DROSOPHILIDAE

Hawaiian Scaptomyza Drosophila

Genus Drosophila

Sophophora Zaprionus Samoaia

Zaprionus Genus Group

Liodrosophila Sphaerogastrella

Styloptera Genus Group

Dettopsomyia Figure 1.3. Phylogenetic hypothesis of Drosophilidae based on Grimaldi (1990), after DeSalle and Grimaldi (1991).

in agreement with Throckmorton’s work, that some subgenera of Drosophila (Scaptodrosophila and Hirtodrosophila) should be recognized as independent genera. Grimaldi’s (1990) analysis, however, also proposed several unconventional relationships. These are the result of bias in his analytical methodology (see Remsen and O’Grady, 2002), and should perhaps be considered suspect. The most controversial of Grimaldi’s results is the placement of the endemic Hawaiian Drosophilidae – a hypothesis that is discussed at length below. A number of molecular phylogenetic studies have proposed genus-level relationships in the family Drosophilidae (DeSalle, 1992; Pelendakis and Solignac, 1993;

Phylogenetic relationships of Drosophilidae

7

Scaptomyza Hawaiian Drosophila virlis group robusta group repleta group quinaria group funebris group tripunctata group testacea group immigrans group Samoaia Hirtodrosophila Liodrosophila

Sophophora

(a)

Drosophila (Dorsilopha) Zaprionus melanogaster group obscura group willistoni group Chymomyza Scaptodrosophila

(b)

Figure 1.4. (a) Summary of molecular phylogenetic hypotheses from Kwiatowski and Ayala (1999); (b) Summary phylogeny based on Tatarenkov et al. (2001).

Russo et al., 1995; Remsen and DeSalle, 1998; Kwiatowski and Ayala, 1999; Tatarenkov et al., 2001; Remsen and O’Grady, 2002). These may vary slightly from one another, but several key relationships are consistently recovered (Figures 1.4, 1.5). The main criticism of the molecular work is that the taxon sampling is much less compared to the morphological studies – a fact largely due to the rarity of some drosophilid groups. Remsen and O’Grady (2002) have greatly expanded both taxon and character sampling in this group (Figure 1.5), but much work remains to be done. Within the genus Drosophila we follow a version of Throckmorton’s (1975) radiations, modified based on results from recent phylogenetic studies and outlined in Figure 1.6. This scheme, while following the latest developments in drosophilid systematics, is somewhat confusing in that several major groups, including the genus

8

How to look at flies Chymomyza Scaptodrosophila melanogaster obscura Sophophora willistoni saltans Zaprionus Dorsilopha virilis repleta dreyfusi bromeliae virilis-repleta Clade mesophragmatica robusta melanica nannoptera Siphlodora Engiscaptomyza Hawaiian Scaptomyza Drosophilidae antopocerus Clade picture wing immigrans funebris pinicola guarani calloptera immigranscardini tripunctata bizonata Clade macroptera rubrifrons quinaria tripunctata testacea polychaeta Liodrosophila Formosa Sphaerogastrella Clade Dettopsomyia Hirtodrosophila Zygothirca Clade Mycodrosophila Paramycodrosophila Samoaia Gitona Rhinoleucophenga Steganinae Clade Stegana Amiota

Figure 1.5. Phylogenetic hypothesis for Drosophilidae presented by Remsen and O’Grady (2002).

Phylogenetic relationships of Drosophilidae

9

Drosophilidae Steganinae Drosophilinae Genus Chymomyza Genus Drosophila subgenus Dorsilopha subgenus Drosophila virilis-repleta radiation annulimana bromeliae canalinea carbonaria carsoni coffeata dreyfusi melanica mesophragmatica nannoptera peruviana polychaeta repleta robusta virilis immigrans-tripunctata radiation calloptera cardini guarani immigrans pallidipennis quinaria testacea tripunctata Hawaiian Drosophilidae Hawaiian Drosophila Scaptomyza subgenus Sophophora melanogaster obscura saltans willistoni Genus Hirtodrosophila Genus Liodrosophila Genus Samoaia Genus Scaptodrosophila Genus Zaprionus Figure 1.6. Taxonomic relationships in Drosophilidae.

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How to look at flies

Drosophila, are not monophyletic. The differences and similarities between the various hypotheses will be compared and contrasted below.

Steganinae The Steganinae is a small, poorly understood subfamily of drosophilid flies that currently contains about 400 described species (Wheeler, 1982, 1986; Baechli, 2005). Many authors consider this group to be “primitive” within the Drosophilidae, but it is actually the sister taxon of the larger, better studied Drosophilinae (Throckmorton, 1966, 1975; Wheeler, 1982; Grimaldi, 1988, 1990). Grimaldi (1988, 1990) proposed two different phylogenetic hypotheses for this group (Figures 1.7a, 1.7b). These trees mostly differed in the placement of the subgenus Amiota. A few molecular studies have included Steganine taxa, but most have not sampled this group extensively enough to propose a firm hypothesis. Remsen and O’Grady’s (2002) results, while only including four Steganine taxa, do not fully agree with either of Grimaldi’s hypotheses (Figure 1.8). Clearly, additional work expanding the number of taxa sampled with molecular characters will be required to better understand the relationships within the basal Drosophilidae. This will, in turn, enhance our knowledge of host plant adaptation in the Drosophilidae as a whole (see Powell, 1997).

Leucophenga Stegana A (Sinophthalmus) Apenthecia A (Phortica) Gitona Cacoxenus Rhinoleucophenga A (Amiota) Pseudiastata Mayagueza Acletoxenus (a)

(b)

Figure 1.7. (a) Phylogeny of Steganinae after Grimaldi (1988); (b) phylogeny of Steganinae after Grimaldi (1990).

Phylogenetic relationships of Drosophilidae

11

Drosophilinae The phylogenetic relationships of the subfamily Drosophilinae have been addressed in a number of studies (Throckmorton, 1975; Okada 1989; Grimaldi, 1990; Remsen and DeSalle, 1998; Tatarenkov and Ayala, 2001; Remsen and O’Grady, 2002). No consensus view can, at this point (however, see Figures 1.4, 1.5), be proposed, so we will review the phylogenetic relationships within this subfamily sequentially below, as we treat each of the major lineages in this group (Figure 1.6; Ashburner, 1989, 2004; Powell, 1997).

Genus Chymomyza The genus Chymomyza is a genus of about 60 described species (Table 1.1), the majority of which are found in the Old and New World tropics. This genus is characterized by having the proclinate orbital setae inserted posterior to the anterior reclinate seta

A (Sinophthalmus) Stegana Gitona Rhinoleucophenga Figure 1.8. Phylogeny of some Steganinae species based on molecular characters (Remsen and O’Grady, 2002). Table 1.1.

Species placed in various genera and selected genus Drosophila groups

Genus

Species and authority

Chymomyza
Drosophila (Dorsilopha) Drosophila (Drosophila)
funebris group

amoena* Loew, 1862; procnemis* Williston, 1896 busckii* Coquillett, 1901; linearidentata Toda, 1986; neobusckii Toda, 1986 altukhovi Imasheva et al., 1994; funebris* Fabricius, 1787; macrospina* Stalker and Spencer, 1939; mutispina* Okada, 1956; pentaspina Parshad and Duggal, 1966; subfunebris* Stalker and Spencer, 1939; trispina Wheeler, 1949 biseriata* Hardy, 1965; crucigera* Grimshaw, 1901; eurypeza* Hardy, 1965; grimshawi* Oldenberg, 1914; gymnobasis* Hardy and Kaneshiro, 1971; mimica* Hardy, 1965; picticornis* Grimshaw, 1901; silvarentis* Hardy and Kaneshiro, 1968; soonae* Takada and Yoon, 1989 duncani* Sturtevant, 1918 aerea* Okada, 1956 leonensis* Wheeler and Kambysellis, 1966 brooksae* Pipkin, 1961; latifasciaformis* Duda, 1940; lebanonensis* Pipkin, 1961; pattersoni* Pipkin, 1956; stonei* Pipkin, 1956 adusta* Loew, 1862; anomala* Hardy, 1965; elmoi* Takada, 1970; palmae* Hardy, 1965 badyi* Burla, 1954a; ghesquierei* Collart, 1937; inermis* Collart, 1937; lineosa* Walker, 1860; sepsoides* Duda, 1939; tuberculatus* Malloch, 1932

Hawaiian Drosophila


Hirtodrosophila
Liodrosophila
Samoaia
Scaptodrosophila
Scaptomyza
Zaprionus


* Species in culture at the Tucson Stock Center;
All taxa in these groups were not listed.

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How to look at flies

C. mesopecta C. procnemoides C. coxata aldrichii group C. aldrichii C. bicoloripes C. laevilimbata C. bicolor C. procnemis C. lahu C. mafu procnemis group C. bambata C. cincifrons C. rufithorax C. vaidyai C. avikam C. fenestrata C. fuscimana C. distincta fuscimana group C. olympia C. wirthi C. amoena C. obscura obscura group C. formosana C. obscuroides C. costata C. mexicana C. caudaluta costata group C. japonica C. atrimana C. leucopoda Figure 1.9. Subdivision and phylogenetic relationships within the genus Chymomyza, after Okada (1976).

(see Chapter 2 for detailed discussion of all morphological characters). Most members of this group also possess femoral spines (see Grimaldi, 1986: 356, 1990:65). Okada (1976) used morphological characters to divide Chymomyza into five species groups: aldrichii, costata, fuscimana, obscura, and procnemis (Figure 1.9). Grimaldi (1986) described seven new species in the aldrichii species group, all of which are found in the Neotropics, and presented a phylogeny of this group (Figure 1.10) that greatly improves the resolution in this clade over Okada’s tree. The aldrichii species group is notable because at least five species placed in it are hypercephalic.

Phylogenetic relationships of Drosophilidae

13

C. microdiopsis C. exophthalma C. mycopelates C. bicoloripes C. procnemoides C. aldrichii C. procnemolita C. diatropa C. guyanensis Figure 1.10. Phylogeny of the aldrichii species subgroup (genus Chymomyza), after Grimaldi (1988).

Many members of the genus Chymomyza are attracted to cut wood, and this may serve as a lek or oviposition site for these species. Grimaldi (1986) considered the use of this substrate to be a specialization, possibly derived from a polyphagous habit. A number of Neotropical Chymomyza species display elaborate courtship behaviors, including male display and aggression (Grimaldi, 1986; Grimaldi and Fenster, 1989).

Genus Drosophila The genus Drosophila is a very large group of well over 1500 described species. Sturtevant (1939, 1942) divided Drosophila into a number of subgenera. Currently, Drosophila is divided into ten subgenera (Ashburner, 1989, 2004), the largest of which is undoubtedly the subgenus Drosophila. The subgenus Sophophora, with over 300 described species, is the second largest. Together, the subgenera Drosophila and Sophophora account for roughly 90 per cent of the diversity in the genus Drosophila. Although no single study has extensively sampled this entire group, several studies have treated parts of this genus. What is clear at this time is that the genus Drosophila is not monophyletic and should probably be divided into several clades (see sections below). Throckmorton’s (1975) review of phylogeny in the family Drosophilidae included a number of “radiations”, or assemblages of closely related species groups (Figure 1.2). This higher-level taxonomic group, fitting between subgenus and species group, is unique to the Drosophilidae. We have adopted its use, particularly within the large, diverse subgenus Drosophila, as a way to organize our discussions of phylogeny in this family. Further phylogenetic analyses employing greatly expanded taxon and character sampling will lead to a firmer understanding of this group, and will serve as the groundwork for further revisionary studies of this family.

Genus Drosophila: subgenus Dorsilopha The subgenus Dorsilopha was described by Sturtevant (1942) as a subgenus of Drosophila containing a single species, D. busckii. Recently, two additional species have been described from Burma (Toda, 1986), bringing the total number of taxa in this subgenus to three (Table 1.1). Remsen and O’Grady (2002) placed this subgenus

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How to look at flies

close to the genus Zaprionus (Figure 1.5), similar to its placement in other molecular phylogenies (Figures 1.4a, 1.4b).

Genus Drosophila: subgenus Drosophila Below, we follow a version of Throckmorton’s radiations, modified after a number of recent molecular and morphological analyses (Tatarenkov and Ayala, 2001; Remsen and O’Grady, 2002). The major groups covered (Figure 1.6) include (1) the virilis-repleta radiation; (2) the immigrans-tripunctata radiation; and (3) the Hawaiian Drosophilidae, a group containing the Hawaiian Drosophila and the genus Scaptomyza.

• funebris

species group. Throckmorton (1975) considered the funebris species group to be basal within the Drosophila radiation because of their distribution and ecological habits. It currently contains seven species (Table 1.1). It is uncertain where this group is placed within the Drosophilidae, although some studies consider this group a part of the immigrans-tripunctata radiation (Figure 1.5; Remsen and O’Grady, 2002). Additional work will be necessary to determine the phylogenetic affinities of this unique clade.

Genus Drosophila: virilis-repleta radiation Throckmorton’s (1975) concept of the virilis-repleta radiation consists of a basal radiation with the virilis, melanica, annulimana, carsoni, bromeliae, peruviana, nannoptera, carbonaria, tumiditarsus, polychaeta, and robusta species groups, as well as the subgenera Phloridosa and Sordophila, and the genus Dettopsomyia. The subgenus Sordophila has been synonymized with the nannoptera species group (Ward and Heed, 1970). Throckmorton (1975) places the ancestor of this radiation in the Old World tropics, although several groups are now Holarctic in distribution. A secondary radiation took place in the New World tropics, and includes the repleta, canalinea, dreyfusi, coffeata, and mesophragmatica species groups. Tatarenkov and Ayala (2001) recently examined the phylogenetic relationships within this radiation using molecular characters from the Ddc and amd loci. Their results are significant in that they call into question the monophyly of the virilisrepleta radiation by placing D. repletoides outside this group. They suggest that this might be the result of insufficient character sampling, and propose gathering additional data to resolve this issue, as the bootstrap support for the placement of this taxon does not preclude its inclusion at the base of this radiation. Including representatives of Phloridosa and Dettopsomyia would also be advisable to fully test the monophyly of the virilis-repleta radiation. Within the remainder of the virilis-repleta radiation, Tatarenkov and Ayala’s (2001) results are highly congruent with those of Throckmorton (1975). The basal members of this group form a heterogeneous radiation that is not monophyletic (Figure 1.11). The New World repleta radiation does consist of four of the five groups that Throckmorton (1975) included (the coffeata group was not sampled). The repleta group is also supported as monophyletic. This is in contrast to the molecular results of Durando et al. (2000) who suggested that the repleta group might not be monophyletic with respect to the other members of the repleta radiation (Figure 1.11, see repleta discussion below).

Phylogenetic relationships of Drosophilidae

15

D. hydei D. eohydei D. mulleri D. buzzatii repleta D. canapalpa D. repleta D. mercatorum repleta radiation

D. ellisoni mesophragmatica

D. gaucha D. camargoi

dreyfusi

D. canalinea

canalinea

D. bromeliae

bromeliae

D. nannoptera

nannoptera

D. aracatacas

annulimana

D. robusta robusta D. sordidula melanica

D. melanica D. virilis

virilis

D. polychaeta Hawaiian Drosophila Figure 1.11. Phylogeny of the virilis-repleta radiation based on Tatarenkov and Ayala (2001).

polychaeta

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How to look at flies

Remsen and O’Grady (2002) have recently examined a number of molecular and morphological characters for the entire subfamily Drosophilinae (see above). Their taxon sampling strategy was slightly different from that of Tatarenkov and Ayala (2001) in that they included Dettopsomyia and several other genera and subgenera within Drosophilidae, but did not sample as intensively within the virilis-repleta radiation itself (Figures 1.5, 1.11). Their results also suggest that the virilis-repleta radiation (sensu Throckmorton, 1975) is not monophyletic (Figure 1.5). The genus Dettopsomyia is placed in a clade at the base of the Drosophilinae, along with the genera Sphaerogastrella and Liodrosophila. This placement is in agreement with that of Okada (1974) and Grimaldi (1990). Although they did not sample the tumiditarsus species group and cannot address its placement, the polychaeta species group is outside of the virilis-repleta radiation in their analysis (Remsen and O’Grady, 2002). Furthermore, they include the subgenus Siphlodora within the virilis-repleta radiation (Figure 1.5), contrary to the results of Throckmorton (1975). This suggests that both the virilis-repleta radiation and the genus Drosophila are not monophyletic. Even though Throckmorton’s (1975) concept of the virilis-repleta radiation is not exactly recovered in recent analyses (Figure 1.2), some of the relationships that he proposed are quite well supported. The close association of the species groups in the repleta radiation is supported by molecular and morphological analyses (Durando et al., 2000; Tatarenkov and Ayala, 2001; Remsen and O’Grady, 2002). The basal members of the virilis-repleta radiation are also supported as being closely related to the New World repleta radiation, even though support for some of these relationships is not high (Tatarenkov and Ayala, 2001; Remsen and O’Grady, 2002). A more complete analysis, with expanded taxon and character sampling, of the virilis-repleta radiation, is needed to better resolve the relationships of these taxa and their affinities within the genus Drosophila.

• annulimana species group. The annulimana species group contains a total of 15

•

described species, all of which are Neotropical in distribution (Table 1.2; Baechli, 2005; Wheeler, 1982, 1986). Throckmorton (1975) considered this group to be a member of the basal virilis-repleta radiation. Tatarenkov and Ayala (2001) also placed this group in a basal position as the sister taxon of the repleta, mesophragmatica, dreyfusi, canalinea, bromeliae, and nannoptera species groups (Figure 1.11). A phylogeny of this group, based on the study by Tosi and Pereira (1993), is shown in Figure 1.12. This clade falls into three main subgroups: (1) schineriararama-pseudotalamanca-gibberosa; (2) arapuan-arassari; and (3) annulimanaaragua-aracataca-arauna. bromeliae species group. The bromeliae species group is a small, poorly known group of Neotropical flower feeding species (Table 1.2; Wheeler, 1982, 1986). Currently, this group contains four described species (Val and Marques, 1996; Baechli, 2005). Tatarenkov and Ayala (2001) considered the bromeliae group to be the sister taxon of the nannoptera species group (Figure 1.11), while Remsen and O’Grady (2002) suggested that the mesophragmatica group was more closely related to the bromeliae species (Figure 1.5). The former placement is better supported, both in terms of character and morphology (both groups have prescutellar setae). Throckmorton (1975) considered the nannoptera, bromeliae, and peruviana

Phylogenetic relationships of Drosophilidae Table 1.2.

17

Species placed in the virilis-repleta radiation

Species group

Species and authority

annulimana

annulimana Duda, 1927; aracataca* Vilela and Val, 1983; aragua Vilela and Pereira, 1982; araicas Pavan and Nacrur, 1950; arapuan Cunha and Pavan, in Pavan and Cunha, 1947; ararama Pavan and Cunha, 1947; arassari Cunha and Frota-Pessoa, in Pavan and Cunha, 1947; arauna Pavan and Nacrur, 1950; breuerae Rocha, 1971; gibberosa* Patterson and Mainland, in Patterson, 1943; paratarsata Vilela, 1985; pseudotalamancana Pereira and Vilela, 1987; schineri Pereira and Vilela, 1987; talamancana* Wheeler, 1968; tarsata Schiner, 1868 aguape Val and Marques, 1996; bromeliae* Sturtevant, 1921 Pavan and Cunha, 1947; florae Sturtevant, 1916 albomarginata Duda, 1927; annularis Sturtevant, 1916; annulosa Vilela and Baechli, 1990; canalinea* Patterson and Mainland, 1944; canalinioides Wheeler, 1957; davidgrimaldii Vilela and Baechli, 1990; hendeli Vilela and Baechli, 1990; melanoptera Duda, 1927; panamensis Malloch, 1926; paracanalinea Wheeler, 1957; parannularis Vilela and Baechli, 1990 carbonaria Patterson and Wheeler, 1942 carsoni Wheeler, 1957 coffeata Williston, 1896; fuscolineata Duda, 1925; pagliolii Cordiero, 1963; pedroi Vilela, 1984 boliviana Duda, 1927; briegeri Pavan and Breuer, 1954; camargoi* Dobzhansky and Pavan, in Pavan, 1950; decemseriata Hendel, 1936; dreyfusi Dobzhansky and Pavan, 1943; fuscipennis Duda, 1927; krugi Pavan and Breuer, 1954; lugubripennis Duda, 1927; wingei Cordiero, 1964 afer Tan, Hsu, and Sheng, 1949; bisetata Toda, 1988; euronotus* Patterson and Ward, 1952; longiserrata Toda, 1988; melanica* Sturtevant, 1916; melanissima Sturtevant, 1916; melanura Miller, 1944; micromelanica* Patterson, in Sturtevant and Novitski, 1941; nigromelanica Patterson and Wheeler, 1942; paramelanica* Patterson, 1943; tsigana Burla and Gloor, 1952 altiplanica Brncic and Koref-Santibanez, 1957; brncici Hunter and Hunter, 1964; canaescens Duda, 1927; gasici Brncic, 1957; gaucha* Jaeger and Salzano, 1953; mesophragmatica Duda, 1927; orkui Brncic and Koref-Santibanez, 1957; pavani* Brncic, 1957; varacochi Brncic and Koref-Santibanez, 1957 acanthoptera* Wheeler, 1949; nannoptera* Wheeler, 1949; pachea Patterson and Wheeler, 1942; wassermani* Pitnick and Heed, 1994 peruviana Duda, 1927 asper* Lin and Tseng, 1971; bivibrissae Toda, 1988; daruma Okada, 1956; fraburu* Burla, 1954; hirtipes* Lamb, 1914; illota Williston, 1896; latifshahi Gupta and Ray-Chaudhuri, 1970; polychaeta* Patterson and Wheeler, 1942 aldrichi* Patterson, in Patterson and Crow, 1940; anceps Patterson and Mainland, 1944; antonietae Tidon-Sklorz and Sene, 2001; arizonae* Ruiz, Heed and Wasserman, 1990; bifurca* Patterson and Wheeler, 1942; borborema* Vilela and Sene, 1977; brevicarinata Patterson and Wheeler, 1942; buzzatii* Patterson and Wheeler, 1942; californica Sturtevant, 1923; carolinae Vilela, 1983; carcinophila Wheeler, 1960; coroica Wasserman, 1962; desertorum Wasserman, 1962; eleonorae Tosi et al., 1990; ellisoni* Vilela, 1983; eohydei* Wasserman, 1962; eremophila* Wasserman, 1962; fasciola Williston, 1896; fascioloides Dobzhansky and Pavan, 1943; fulvalineata Patterson and Wheeler, 1942; fulvimacula* Patterson and Mainland, 1944; fulvimaculoides* Wasserman and Wilson, 1957; gouveai Tidon-Sklorz and Sene, 2001; guayllabambae* Rafael and Arcos, 1988; hamatofila* Patterson and Wheeler, 1942; hermioneae Vilela, 1983; hexastigma* Patterson and Mainland, 1944; huayla Pilar, Pilares and Vasquez, 1988; huaylasi* Pla and Fontdevila, in Fontdevila et al., 1990; huancavilcae Rafael and Arcos, 1989; huckinsi Etges and Heed, in Etges et al., 2001; huichole Etges and Heed, in Etges et al., 2001; hydei* Sturtevant, 1921; icteroscuta Wheeler, 1949; inca Dobzhansky and Pavan, 1943; (Continued)

bromeliae bromelioides canalinea

carbonaria carsoni coffeata dreyfusi

melanica

mesophragmatica

nannoptera peruviana polychaeta

repleta

18 Table 1.2.

How to look at flies (Continued )

Species group

Species and authority

repleta (continued )

ivai Vilela, 1983; koepferae* Fontdevila and Wasserman, in Fontdevila et al., 1988; leonis* Patterson and Wheeler, 1942; limensis* Pavan and Patterson, in Pavan and Cunha, 1947; linearepleta Patterson and Wheeler, 1942; longicornis* Patterson and Wheeler, 1942; mainlandi* Patterson, 1943; mapiriensis Vilela and Bachli, 1990; mariettae Vilela, 1983; martensis* Wasserman and Wilson, 1957; mathisi Vilela, 1983; mayaguana* Vilela, 1983; mercatorum* Patterson and Wheeler, 1942; meridiana* Patterson and Wheeler, 1942; meridionalis* Wasserman, 1962; mettleri* Heed, 1977; micromettleri* Heed, 1989; mojavensis* Patterson, in Patterson and Crow, 1940; moju Pavan, 1950; mojuoides Wasserman, 1962; mulleri* Sturtevant, 1921; navojoa* Ruiz, Heed and Wasserman, in Ruiz et al., 1990; neohydei Wasserman, 1962; neorepleta* Patterson and Wheeler, 1942; nigricruria* Patterson and Mainland, in Patterson, 1943; nigrodumosa* Wasserman and Fontdevila, in Fontdevila et al., 1990; nigrohydei* Patterson and Wheeler, 1942; nigrospiracula* Patterson and Wheeler, 1942; novemaristata Dobzhansky and Pavan, 1943; onca Dobzhansky and Pavan, 1943; pachuca* Wasserman, 1962; papei Baechli and Vilela, 2002; paraguttata Thompson, in Wheeler, 1957; paranaensis* Barros, 1950; parisiena* Heed and Grimaldi, 1991; pegasa* Wasserman, 1962; peninsularis* Patterson and Wheeler, 1942; pictilis Wasserman, 1962; pictura Wasserman, 1962; promeridiana Wasserman, 1962; propachuca* Wasserman, 1962; prorepleta Duda, 1925; pseudorepleta Vilela and Bachli, 1990; querubimae Vilela, 1983; racemova Patterson and Mainland, 1944; ramsdeni Sturtevant, 1916; repleta* Wollaston, 1858; richardsoni* Vilela, 1983; ritae* Patterson and Wheeler, 1942; rosinae Vilela, 1983; senei Vilela, 1983; serenensis Brncic, 1957; serido* Vilela and Sene, 1977; seriema Tidon-Sklorz and Sene, 1995; spenceri* Patterson, 1943; stalkeri* Wheeler, 1954; starmeri* Wasserman, Koepfer and Ward, 1973; straubae* Heed and Grimaldi, 1991; subviridis Patterson and Mainland, in Patterson, 1943; uniseta Wasserman, Koepfer and Ward, 1973; venezolana* Wasserman, Fontdevila, and Ruiz, 1983; vicentinae Vilela, 1983; wheeleri* Patterson and Alexander, 1952; zottii Vilela, 1983 bai Watabe and Liang, in Watabe et al., 1990; cheda Tan, Hsu, and Sheng; colorata Walker, 1849; gani Liang and Zhang, in Watabe et al., 1990; lacertosa* Okada, 1956; medioconstricta Watabe et al., in Watabe et al., 1990; moriwakii Okada and Kurokawa, 1957; neokadai Kaneko and Takada, 1966; nigrescens Chen and Watabe, 1993; okadai Takada, 1956; pseudosordidula Kaneko, Tokumitsu, and Takada, 1964; robusta* Sturtevant, 1916; seyanii Chassagnard and Tsacas, in Chassagnard et al., 1997; sordidula* Kikkawa and Peng, 1938; unimaculata Strobl, 1893; yunnanensis Watabe and Liang, in Watabe et al., 1990; zonata Chen and Watabe, 1993 repletoides Hsu, 1943 americana Spencer, 1938; borealis Patterson, 1952; canadiana Takada and Yoon, 1989; ezoana Takada and Okada, 1958; flavomontana Patterson, 1952; kanekoi Watabe and Higuchi, 1979; lacicola Patterson, 1944; littoralis Meigen, 1830; lummei Hackman, 1972; montana Patterson and Wheeler, 1942; novamexicana Patterson, 1941; virilis Sturtevant, 1916

robusta

tumiditarsus virilis

•

groups to be close relatives on the basis of internal anatomy. Members of the flower breeding peruviana and flavopilosa species groups should be examined to more clearly understand the evolution of this group. canalinea species group. The canalinea species group consists of a total of eleven Neotropical species (Table 1.2; Wheeler, 1982, 1986; Baechli, 2005). Both morphology (Throckmorton, 1975) and molecular (Tatarenkov and Ayala, 2001)

Phylogenetic relationships of Drosophilidae

19

D. aragua D. aracataca D. annulimana D. arauua D. arapuan D. arassai D. schnerei D. pseudotalamanca D. ararama D. gibberosa Figure 1.12. Phylogeny of the annulimana species group (Tosi and Pereira, 1993).

•

•

•

•

•

place the canalinea group in the derived repleta radiation, along with the repleta, dreyfusi, and mesophragmatica species groups (Figures 1.1, 1.11). carbonaria species group. The carbonaria species group is monotypic, containing only the species D. carbonaria (Table 1.2). Although Throckmorton (1975) placed this group in the basal virilis-repleta radiation (Figure 1.2), the placement of this species has never been examined using cladistic methods. Drosophila carbonaria has been bred from mesquite fluxes, and its distribution mirrors that of some Prosopis species from Mexico into the southwestern United States (Patterson and Stone, 1952). carsoni species group. The carsoni species group is another monotypic group found in the United States (Table 1.2). Throckmorton (1975) reported that the eastern and western populations of D. carsoni might actually represent more than one species. Although this species can be reared in the laboratory it is quite rare in the wild, and molecular, chromosomal, and breeding experiments have never been performed. coffeata species group. This Neotropical group is comprised of four known species (Table 1.2; Baechli, 2005). Vilela (1984) reports that this group is synonymous with the castanea species group (sensu Throckmorton, 1975). It is in the derived repleta radiation, along with the repleta, mesophragmatica, dreyfusi, and canalinea species groups (Figure 1.2). It has never been examined in a phylogenetic study. dreyfusi species group. The dreyfusi species group consists of nine known species (Table 1.2; Baechli, 2005). It is considered to be closely related to the repleta, mesophragmatica, canalinea, and coffeata species groups (Figures 1.2, 1.4b, 1.5; Throckmorton, 1975; Tatarenkov and Ayala, 2001; Remsen and O’Grady, 2002). Phylogenetic relationships within this species group have never been examined. melanica species group. Stalker (1966) examined the phylogenetic relationships of some members of the melanica group using polytene chromosome banding patterns,

20

How to look at flies

D. paramelanica D. melanica D. euronotus D. melanura D. nigromelanica D. micromelanica Figure 1.13. Phylogeny of the melanica species group based on polytene banding patterns (Stalker, 1966).

•

•

interspecific hybridization studies, and geographic distribution (Figure 1.13). He considered D. micromelanica and D. nigromelanica to be the basal-most members of this group. Drosophila melanura, D. euronotus, and the D. melanica–D. paramelanica sibling pair are successively more derived. There are currently twelve species in this group (Table 1.2; Baechli, 2005), none of which have been studied with molecular characters. Members of this group have been reported from sap and slime fluxes on a number of species, including red oak, black locust, elm, and willow (Throckmorton, 1975). According to Throckmorton (1975), all species in this group can be readily cultured in the laboratory. This group is related to other members of the virilis-repleta radiation (Figures 1.2, 1.4b, 1.5; Throckmorton, 1975; Tatarenkov and Ayala, 2001; Remsen and O’Grady, 2002). mesophragmatica species group. The mesophragmatica species group is a small group containing nine described species, all of which are Neotropical in distribution (Table 1.2; Baechli, 2005). Phylogenetic relationships (Figure 1.14) in this group have been inferred using polytene chromosome banding patterns (Brncic et al., 1971) and isozymes (Nair et al., 1971). These studies suggest that D. pavani and D. gaucha are sister species and placed basal to D. mesophragmatica and its relatives (Figure 1.14). Most species of the mesophragmatica group are easily culturable on standard cornmeal media. nannoptera species group. The four species in the nannoptera group are found in the desert regions of North and South America (Table 1.2; Baechli, 2005). The phylogeny of the nannoptera group has been inferred using polytene chromosome banding patterns (Figure 1.15, Ward and Heed, 1970). Their results show that D. wassermani (a species referred to as “species W” and described by Pitnick and Heed, 1994) and D. nannoptera are sister taxa. With the exception of a single heterozygous inversion in D. nannoptera, these two species are homosequential. Drosophila acanthoptera is the sister taxon of the D.wassermani–D. nannoptera clade, and D. pachea is basal in this group. A fifth species in this group has recently been discovered in southern Mexico (Castrezana, unpublished). Most of the

Phylogenetic relationships of Drosophilidae

21

D. gasici D. brncici D. mesophragmatica D. viracochi D. pavani D. gaucha Figure 1.14. Phylogeny of the mesophragmatica species group based on polytene chromosomes and isozymes (Brncic et al., 1971; Nair et al., 1971).

D. wassermani D. nannoptera D. acanthoptera D. pachea Figure 1.15. Phylogeny of the nannoptera species group based on polytene chromosomes (Ward and Heed, 1970).

•

•

nannoptera species can be cultured in the laboratory, although some require special diets (see Chapters 7–11). Additional systematic work on the nannoptera group, especially using molecular characters, should be performed to test relationships within this clade. peruviana species group. The Neotropical peruviana species group consists of a single, flower breeding species (Table 1.2; Baechli, 2005). Throckmorton (1975) considered D. peruviana to be most closely allied with the nannoptera and bromeliae groups (Figure 1.2). polychaeta species group. The polychaeta species group is characterized by having three dorsocentral setae (most members of the genus Drosophila have only two – see Chapter 2). This group currently contains a total of eight species (Table 1.2; Baechli, 2005), distributed mainly in the Asian tropics. Throckmorton (1975) and Tatarenkov and Ayala (2001) placed the polychaeta group close to the base of the virilis-repleta radiation (Figures 1.2, 1.4b). Remsen and O’Grady (2002) suggested that this group came out in a clade separate from the remainder of the genus Drosophila, as the sister taxon of the clade formed by Dettopsomyia, Liodrosophila, and Sphaerogastrella

22

How to look at flies

•

•

(Figure 1.5). The latter result is consistent with the geographic distributions of these species, but additional work will be required to resolve the placement of the polychaeta species group within the genus Drosophila. Relationships within this group are not well understood, and have yet to be tested with phylogenetic methods. repleta species group. The repleta species group is one of the largest and most extensively studied groups in the subgenus Drosophila, with nearly 100 species (Table 1.2; Baechli, 2005). This group, largely through the efforts of Patterson, Stone and others (see Patterson and Stone, 1952; Barker and Starmer, 1982; Barker et al., 1990 for a review), has served as a model system for evolutionary and ecological studies. Wasserman (1982, 1992) has examined the phylogeny of the repleta group using polytene chromosome banding patterns (Figure 1.16) and Vilela (1983) has revised this group, largely based on the morphology of the male genitalia. When recoded and analyzed in a parsimony framework, the polytene chromosome phylogeny, while not highly resolved, displays almost no homoplasy (O’Grady et al., 2001). Only a few groups are apparent, including parts of the mojavensis, mulleri, stalkeri, martensis, and buzzatii clusters (Figure 1.16). Durando et al. (2000) have recently used several molecular loci to investigate the phylogeny of the repleta species group (Figure 1.17). Their results suggest that the repleta group is not monophyletic with respect to several closely related taxa placed in the repleta radiation by Throckmorton (1975, see above), including the dreyfusi, canalinea, and mesophragmatica species groups. They state that this is due to poor resolution at the base of the tree, and that additional characters may resolve this issue. Their study was quite informative for some of the more derived lineages in this group. For example, the mulleri and buzzatii complexes were sister taxa and monophyletic. Other complexes and subgroups, such as the eremophila, hydei, mercatorum, and meridiana are also monophyletic, although their exact relationships are unclear. The latter results are in agreement with those of Wasserman’s polytene chromosome studies (Figure 1.16). Spicer and Pitnick (1996) present a phylogeny of the hydei subgroup that gives a more detailed view of evolution in this group than the Durando et al. (2000) tree alone (Figures 1.17, 1.18). robusta species group. The robusta group contains seventeen described species (Table 1.2; Baechli, 2005) found mainly in the temperate regions of North America, Europe, and Asia. There is at least one Afrotropical taxon. Throckmorton (1975) reports that the robusta species group has been reared from sap fluxes. The majority of these species can be reared in the laboratory. Narayanan (1973) has examined the phylogenetic relationships in six species of the robusta group using polytene chromosome banding patterns (Figure 1.19). Drosophila colorata was arbitrarily designated as standard so it appears at the base of this unrooted phylogeny. Based on their chromosomes, D. morkwakii and D. colorata are closely related. Drosophila lacertosa and D. robusta are sister taxa, with D. sordidula and D. pseudosordidula being intermediate species in the phylogeny (Figure 1.19). The robusta group is thought to be most closely related to the melanica species group (Levitan, 1982), with which it shares a common distributional pattern and ecological affinities. Stalker (1966) considered the robusta species group to be the

Phylogenetic relationships of Drosophilidae

23

90

66

93

89

100

aldrichi wheeleri mulleri nigrodumosa arizonae mojavensis & mojavensis mulleri clusters huaylasi navojoa mayaguana parisiena straubae desertorum borborema serido buzzatii cluster buzzatii koepferae martensis uniseta martensis cluster venezolana starmeri stalkeri stalkeri cluster richardsoni hamatofila hexastigma longicornis propachuca pachuca mainlandi ritae spenceri anceps camargoi canalinea ellisoni eremophila mettleri micromettleri leonis gaucha pavani hydei nigrohydei fulvimacula mercatorum paranaensis meridiana meridionalis pegasa neorepleta nigrospiracula virilis melanogaster cyrtoloma longipedis

Figure 1.16. Chromosome phylogeny of the repleta species group (Wasserman, 1992; O’Grady et al., 2001).

24

How to look at flies

D. aldrichi D. wheeleri D. mulleri D. nigrodumosa D. arizonae D. mojavensis D. huaylasi D. navojoa D. mayaguana D. straubae mulleri complex D. parisiena D. desertorum D. longicornis D. propachuca D. pachuca D. mainlandi D. ritae D. hexastigma D. spenceri D. borborema D. serido D. buzzatii D. koepferae buzzatii complex D. starmeri D. venezolana D. martensis D. uniseta D. richardsoni D. stalkeri mulleri complex D. hamatofila fasciola subgroup D. ellisoni D. canalinea *** D. camargoi *** D. eremophila eremophila complex D. mettleri D. micromettleri D. mercatorum mercatorum subgroup D. paranensis D. neorepleta repleta subgroup D. fulvimacula D. gaucha *** D. pavani *** D. hydei hydei subgroup D. nigrohydei D. anceps D. nigrospiracula anceps complex D. leonis D. meridiana D. meridionalis meridiana complex D. pegsa Figure 1.17. Molecular phylogeny of the repleta species group, after Durando et al. (2000).

Phylogenetic relationships of Drosophilidae

25

D. hydei D. eohydei D. nigrohydei D. bifurca Figure 1.18. Molecular phylogeny of the hydei subgroup (repleta species group), after Spicer and Pitnick (1996).

D. lacertosa D. robusta D. pseudosordidula D. sordidula D. colorata D. moriwakii Figure 1.19. Chromosome phylogeny of the robusta species group (Narayanan, 1973).

•

•

sister of the Hawaiian Drosophilidae on the basis of polytene chromosome banding patterns. This is in contrast to Throckmorton (1975), who suggested that the Hawaiian species were more closely allied with the immigrans-Hirtodrosophila radiation and the robusta group was placed in the virilis-repleta radiation (Figure 1.2). Grimaldi (1990) also suggested an alternative sister group for the Hawaiian Drosophilidae (see discussion below). It is clear from a number of molecular studies that the robusta group is more closely related to the Hawaiian taxa than the immigrans-Hirtodrosophila radiation, supporting the hypothesis of Stalker (1966). However, it may not be the exact sister group as other species in virilis-repleta taxa are also closely related to the Hawaiian lineage (Figures 1.4a, 1.4b, 1.5; DeSalle, 1992; Remsen and DeSalle, 1998; Remsen and O’Grady, 2002). tumiditarsus species group. This species group contains a single species, D. repletoides, which is found in China and Japan (Table 1.2; Baechli, 2005). Throckmorton (1975) places this species in the virilis-repleta radiation (Figure 1.2). Tatarenkov and Ayala (2001) suggest that this species may be outside of the virilis-repleta radiation (Figure 1.11). Additional work should be done to clarify the affinities of this taxon. virilis species group. The virilis group has served as a model system for comparative studies and, as such, the phylogenetic relationships within this group have been extensively examined. Spicer has used a variety of techniques, including two-D gel electrophoresis (Spicer, 1988), nucleotide sequences (Spicer, 1992; Spicer and Bell,

26

How to look at flies

2002), and quantitative characters (Spicer, 1995) to infer phylogeny in this group. The virilis species group consists of about thirteen described species that are typically found in a boreal distribution (Table 1.2; Baechli, 2005). Currently, the status of two, D. americana and D. texana, is somewhat questionable. The latter was initially described as a subspecies (Patterson et al., 1940), but has since been considered a full species (Throckmorton, 1982) on the basis of chromosome and molecular (Spicer and Bell, 2002) differences (Figure 1.20a). McAllister (2002) points out that this chromosomal difference is a widespread polymorphism and subdivision of D. americana is not warranted (Figure 1.20b). Spicer and Bell (2002) also report that the eastern and western forms of another species, D. borealis, do not form a monophyletic assemblage, and may represent two separate species (Figure 1.20a). Nurminsky et al. (1996) also examined the evolution of the Adh locus in the virilis group, and proposed that either this gene had duplicated prior to the diversification of the virilis group and duplicate copies had been lost independently several times,

D. novamexicana D. americana D. texana

(b)

D. lummei D. virilis D. kanekoi D. ezoana D. littoralis D. canadiana D. flavomontana D. montana D. borealis (eastern) D. lacicola (a)

D. borealis (western)

Figure 1.20. (a) Phylogeny of the virilis group based on the work of Spicer (1992) and Spicer and Bell (2002); (b) relationships between D. americana and D. novamexicana after McAllister (2002).

Phylogenetic relationships of Drosophilidae

27

or the duplication even had taken place independently in both the virilis and montana subgroups (Figure 1.21). Genus Drosophila: immigrans-tripunctata radiation Throckmorton’s (1975) immigrans-Hirtodrosophila radiation, much like his virilisrepleta radiation, is a large heterogeneous group containing many species groups and subgenera within the genus Drosophila, as well as a number of distinct genera (Figure 1.2). Based on recent morphological (Grimaldi, 1990; Figure 1.3) and molecular (e.g., Kwiatowski and Ayala, 1999; Remsen and O’Grady, 2002; Figure 1.4) studies, it is clear that the members of the immigrans and tripunctata radiations are closely related. It is likewise clear that the genus Hirtodrosophila forms a genus distinct from the various species groups in the tripunctata and immigrans radiations (Grimaldi, 1988; Figure 1.22). Therefore, rather than discussing Hirtodrosophila here, we will follow

D. americana D. texana D. lummei D. virilis-1 D. virilis-2 D. flavomontana D. borealis D. montana-2 D. lacicola-2 D. lacicola-1 D. montana-1 Figure 1.21. Gene phylogeny of the alcohol dehydrogenase locus in the virilis species group (Nurminski et al., 1999).

Hirtodrosophila Zygothrica Mycodrosophila Drosophila Figure 1.22. Placement of the genus Hirtodrosophila relative to the genus Drosophila, after Grimaldi (1987).

28

How to look at flies

Remsen and O’Grady (2002) and consider the genus Hirtodrosophila separately from the remainder of the immigrans and tripunctata radiation species. With the exception of a few species groups and genera, relationships within the large immigrans-tripunctata radiation have not been extensively studied. Throckmorton (1975) proposed that a basal radiation (the immigrans radiation) led to two further radiations, one in the New World (the tripunctata radiation) and another in the Old World (the Old World Hirtodrosophila radiation; see Hirtodrosophila discussion below). The basal immigrans radiation contains some members of the genus Drosophila, such as the subgenus Siphlodora and the immigrans, testacea, and quinaria species groups, and the genera Zaprionus and Samoaia. The tripunctata radiation is made up of the large tripunctata group and several closely related species groups, including calloptera, cardini, rubifrons, and macroptera (Figure 1.2). Remsen and O’Grady (2002) recovered an immigrans-tripunctata clade in their analysis (Figure 1.5). This consisted of a number of species groups placed in the immigrans and tripunctata radiations (sensu Throckmorton, 1975). However, a monophyletic tripunctata radiation was not recovered in this study (Remsen and O’Grady, 2002). Furthermore, several taxa placed in this group by Throckmorton (1975; Figure 1.2), such as the subgenus Dorsilopha, the Hawaiian Drosophila, and the genera Zaprionus and Scaptomyza, were found to have strong affinities with other groups. The subgenus Dorsilopha, for example, was found to be the sister group of Zaprionus and placed basal to the subgenus Sophophora. The Hawaiian Drosophila and the genus Scaptomyza, referred to as the Hawaiian Drosophilidae (see below), were well supported as sister taxa and collectively placed as the sister group of the virilis-repleta radiation (Figures 1.4, 1.5). These findings are in agreement with some other molecular and morphological studies. Additional taxon and character sampling within this large, complex group will be needed to further resolve phylogenetic relationships. Perlman et al. (2003) have also recently reconstructed a phylogeny of many species in the immigrans and tripunctata radiations in their study of nematode parasitism in mycophagous Drosophila species (Figure 1.23). Their results, while not topologically identical to those of Remsen and O’Grady (Figure 1.5), are similar in that they do not recover monophyletic immigrans and tripunctata radiations. Their phylogeny stresses relationships within the quinaria and testacea species groups, and it will be discussed at more length in those sections (below).

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•

calloptera species group. The calloptera species group is endemic to the Neotropics (Burla and Pavan, 1953) and currently contains eight described species (Table 1.3; Baechli, 2005). This group is characterized by having spectacular wing patterns (Wheeler, 1968) which are reminiscent of the adiastola species group of picture winged Hawaiian Drosophila. Throckmorton (1975) considered this group to be most closely related to the species groups of the tripunctata radiation (Figure 1.2), and Remsen and O’Grady suggest that, within this large radiation, the guarani species group is the sister taxon of the calloptera group (Figure 1.5). cardini species group. The cardini species group contains sixteen described species divided into two subgroups, cardini and dunni (Table 1.3; Heed and Krishnamurthy, 1959; Baechli, 2005). This species group is entirely Neotropical in distribution, and many members of the dunni subgroup are endemic to single Caribbean islands.

Phylogenetic relationships of Drosophilidae

29

D. acutilabella

cardini group

D. cardini D. tripunctata tripunctata group D. brachynephros D. phalerata D. unispina D. curvispina D. falleni D. innubila D. palustris D. subpalustris D. deflecta

quniaria group

D. limbata D. munda D. quinaria D. occidentalis D. suboccidentalis D. recens D. subquinaria D. transversa D. nigromaculata D. guttifera D. kuntzei D. histrio

histrio group

D. macroptera

macroptera group

D. testacea D. orientacea D. neotestacea

testacea group

D. putrida D. immigrans D. busckii D. histriodes D. affinis D. pseudoobscura

Sophophora

D. melanogaster D. yakuba Figure 1.23. Phylogeny of mycophagous Drosophila in the tripunctata and immigrans radiations based on Perlman et al. (2003).

30 Table 1.3.

How to look at flies Species placed in the immigrans-tripunctata radiation

Species group

Species and authority

calloptera

atrata Burla and Pavan, 1953; calloptera Schiner, 1868; kallima Wheeler, 1957; lindae Wheeler, 1968; maracaya Wheeler, 1957; ornatipennis* Williston, 1896; quadrum Wiedemann, 1830; schildi Malloch, 1924

cardini

acutilabella* Stalker, 1953; antillea* Heed, 1962; arawakana* Heed, 1962; bedichecki Heed and Russel, 1971; belladunni* Heed and Krishnamurthy, 1959; cardini* Sturtevant, 1916; cardinoides* Dobzhansky and Pavan, 1943; caribiana* Heed, 1962; dunni* Townsend and Wheeler, 1955; neocardini* Streisinger, 1946; neomorpha Heed and Wheeler, 1957; nigrodunni* Heed and Wheeler, 1957; parthenogenetica* Stalker, 1953; polymorpha* Dobzhansky and Pavan, 1943; procardinoides* Frydenberg, 1956; similis* Williston, 1896

guarani

alexanderi Cordiero, 1951; araucana Brncic, 1957; griseolineata* Duda, 1927; guaraja King, 1947; guarani* Dobzhansky and Pavan, 1943; guaru Dobzhansky and Pavan, 1943; huilliche Brncic, 1957; limbinervis Duda, 1925; maculifrons Duda, 1927; ornatifrons Duda, 1927; peruensis Wheeler, 1959; subabdia* Patterson and Mainland, in Patterson, 1943; tucumana Vilela and Pereira, 1985; urubamba Vilela and Pereira, 1993

immigrans

afoliolata Zhang and Toda, in Zhang, Toda and Watabe, 1995; albomicans* Duda, 1923; annulipes Duda, 1924; aplophallata Zhang and Toda, in Zhang, Toda and Watabe, 1995; audientis Lin and Ting, 1971; balneorum Sturtevant, 1927; bimaculata Zhang and Toda, in Zhang, Toda and Watabe, 1995; brevitabula Zhang and Toda, 1992; burmae Toda, 1986; calceolata Duda, 1926; cheongi Takada and Momma, 1975; circumdata Duda, 1926; clarinervis Toda, 1986; crispipennis Okada and Carson, 1983; curviceps Okada and Kurokawa, 1957; eminentiula Zhang and Shi, in Zhang, Toda and Watabe, 1995; eprocessata Zhang and Toda, in Zhang, Toda and Watabe, 1995; flavimedifemur Zhang and Toda, 1988; flavitibiae Toda, 1986; formosana* Duda, 1926; fuscicostata Okada, 1966; fustiformis Zhang and Liang, 1993; hexastriata Tan, Hsu and Sheng, 1949; hypocausta* Osten Sacken, 1882; ichinosei Zhang and Toda, in Zhang, Toda and Watabe, 1995; immigrans* Sturtevant, 1921; kepulauana* Wheeler, in Wilson et al., 1969; kitagawai Toda, 1986; kohkoa* Wheeler, in Wilson et al., 1969; lineata van der Wulp, 1881; lineolata de Meijere, 1914; longisetae Zhang, Lin and Gan, 1990; maryensis Gupta and Dwivedi, 1980; metasetigerata Gupta and Kumar, 1986; minangkabau Zhang and Toda, in Zhang, Toda and Watabe, 1995; monochaeta Sturtevant, 1927; nakanoi Zhang and Toda, in Zhang, Toda and Watabe, 1995; nasuta* Lamb, 1914; nasutoides Okada, 1964; neohypocausta* Lin and Wheeler, in Lin and Tseng, 1973; neoimmigrans Gai and Krishnamurthy, 1982; neosignata Kumar and Gupta, 1988; neonasuta Sajjan and Krishnamurthy, 1972; nigridentata Toda and Peng, in Zhang, Toda and Watabe, 1995; nigrilineata Angus, 1967; nigrodigita Lin and Ting, 1971; niveifrons Okada and Carson, 1982; nixifrons Tan, Hsu and Sheng, 1949; notostriata Okada, 1966; nullilineata Zhang and Toda, 1988; obscurinervis Toda, 1986; oritisa Chen, 1990; padangensis Zhang and Toda, in Zhang, Toda and Watabe, 1995; pallidifrons Wheeler, in Wilson et al., 1969; papilla Zhang and Shi, in Zhang and Toda, 1992; paraimmigrans Gai and Krishnamurthy, 1986; pararubida* Mather, 1961; parasignata Takada, Momma and Shima, 1973; parustulata Zhang and Toda, 1988; parviprocessata Toda, 1986; pentafuscata Gupta and Kumar, 1986; pentastriata Okada, 1966; pseudotetrachaeta Angus, 1967; pulaua* Wheeler, in Wilson et al., 1969; purpurea Gupta and Sundaran, 1990; quadrilineata de Meijere, 1911; quadriserrata Okada and Carson, 1982; ruberrima de Meijere, 1911; ruberrimoides Zhang and Gan, 1986; rubida* Mather, 1960; rubra Sturtevant, 1927; senticosa Zhang and Shi, in Zhang, Toda and Watabe, 1995; serrulata Zhang and Toda, in Zhang, Toda and Watabe, 1995; shwezayana Toda, 1986; siamana Ikeda et al., 1983; signata* Duda, 1923; spuricurviceps Zhang and Gan, 1986; subfasciata de Meijere, 1914; sulfurigaster* Duda, 1923; synpanishi Okada, 1964; taekjuni Kim and Joo, 2002; taiensis Kumar and Gupta, 1988; taipinsanensis Lin and Tseng, 1973; tetrachaeta Angus, 1964; tetravitta Takada and Momma, 1975; torquata Zhang and Toda, in Zhang, Toda and Watabe,

Phylogenetic relationships of Drosophilidae Table 1.3.

31

(Continued )

Species group

Species and authority

immigrans (continued) pallidipennis quinaria

1995; trichaeta Angus, 1967; trilimbata Bezzi, 1928; ustulata de Meijere, 1908; wangi Toda and Zhang, in Zhang, Toda and Watabe, 1995; xanthogaster Duda, 1924 pallidipennis* Dobzhansky and Pavan, 1943 analspina Singh and Negi, 1995; angularis Okada, 1956; bondarenkoi Sidorenko, 1993; brachynephros Okada, 1956; curvispina Watabe and Toda, 1984; deflecta Malloch, in Malloch and McAtee, 1924; falleni* Wheeler, 1960; guttifera Walker, 1849; innubila Spencer, in Patterson, 1943; kuntzei Duda, 1924; limbata von Roser, 1840; magnaquinaria Wheeler, 1954; mediobandes Dwevedi and Gupta, 1980; munda Spencer, 1942; natasha Gornostayev, 1992; neokuntzei Singh and Gupta, 1981; nigriculter Okada, 1988; nigromaculata Kikkawa and Peng, 1938; occidentalis Spencer, 1942; palustris* Spencer, 1942; parakuntzei Okada, 1973; phalerata* Meigen, 1830; quinaria Loew, 1866; recens Wheeler, 1960; rellima Wheeler, 1960; schachti Baechli, Vilela and Haring, 2002; suboccidentalis Spencer, 1942; subpalustris* Spencer, 1942; suffusca Spencer, in Patterson, 1943; takadai Lee, 1964; tenebrosa Spencer, in Patterson, 1943; triantlia Okada, 1988; transversa Fallen, 1823; unispina Okada, 1956 neotestacea Grimaldi et al., 1992; orientacea Grimaldi et al., 1992; putrida* Sturtevant, 1916; testacea von Roser, 1840 addisoni Pavan, 1950; albescens Frota-Pessoa, 1954; albicans Frota-Pessoa, 1954; albirostris Sturtevant, 1921; angustibucca Duda,1925; arcosae Vela and Rafael, 2001; argenteifrons Wheeler, 1954; bandeirantorum Dobzhansky and Pavan, 1943; bifilum Frota-Pessoa, 1954; bipunctata Patterson and Mainland, 1943; blumelae Pipkin and Heed, 1964; bodemannae Pipkin and Heed, 1964; carlosvilelai Vela and Rafael, 2001; converga Heed and Wheeler, 1957; cuaso Vilela and Ratcov, 2000; cundinamarca Vilela and Bachli, 2000; curvapex Frota-Pessoa, 1954; divisa Duda, 1927; facialba Heed and Wheeler, 1957; fairchildi Pipkin and Heed, 1964; fontdevilelai Vela and Rafael, 2001; fragilis Wheeler, 1949; frotapessoai Vilela and Bachli, 1990; greerae Pipkin and Heed, 1964; hansoni Pipkin,1964; hansonioides Pipkin, 1966; johnstonae Pipkin and Heed, 1964; leticiae Pipkin, 1967; machachensis Vela and Rafael, 2001; mcclintockae Pipkin, 1964; mediodelta Heed and Wheeler, 1957; mediodiffusa* Heed and Wheeler, 1957; medioimpressa Frota-Pessoa, 1954; medioparva Heed and Wheeler, 1957; mediopicta Frota-Pessoa, 1954; mediopictoides* Heed and Wheeler, 1957; mediopunctata Dobzhansky and Pavan, 1943; mediostriata* Duda, 1925; mediovittata Frota-Pessoa, 1954; medioobscurata Duda, 1925; mediocris Frota-Pessoa, 1954; metzii Sturtevant, 1921; morena Frota-Pessoa, 1954; mesostigma Frota-Pessoa, 1954; neoguaramunu Frydenburg, 1956; nigricinta Frota-Pessoa, 1954; paraguayensis Duda, 1927; paramediostriata Townsend and Wheeler, 1955; pasochoensis Vela and Rafael, 2001; pellewae Pipkin and Heed, 1964; pilaresae Vela and Rafael, 2001; platitarsus Frota-Pessoa, 1954; prosimilis Duda, 1927; pruinifacies Frota-Pessoa, 1954; roehrae Pipkin and Heed, 1964; rostrata Duda, 1925; semialba Duda, 1925; setula Heed and Wheeler, 1957; spinatermina Heed and Wheeler, 1957; tomasi Vela and Rafael, 2001; trapeza Heed and Wheeler, 1957; triangula Wheeler, 1949; triangulina Duda, 1927; trifiloides Wheeler, 1957; trifilum Frota-Pessoa, 1954; tripunctata* Loew, 1862; tristriata Heed and Wheeler, 1957; unipunctata* Patterson and Mainland, 1943; valenciai Vela and Rafael, 2001; whartonae Pipkin and Heed, 1964

testacea tripunctata

* Species in culture at the Tucson Stock Center.

Heed and Russell (1971) have generated a polytene chromosome phylogeny of this group (Figure 1.24a) that supports the notion that each subgroup is monophyletic. Within the cardini subgroup, several sibling species pairs are found (D. bedichecki and D. cardinoides, D. parthenogenetica and D. procardinoides, D. neomorpha and

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How to look at flies

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D. polymorpha). Drosophila neocardini and D. cardini are basal in the cardini subgroup. Within the dunni subgroup, chromosomes are unable to resolve much structure, other than the fact the D. acutilabella is basal in this clade. Hollocher (1998) has recently generated a phylogeny of the dunni subgroup using mitochondrial DNA sequences. This phylogeny adds structure lacking in the chromosome study (Figure 1.24b). The dunni-belladunni-similis complex recovered in the chromosome analysis is basal and paraphyletic with respect to the remaining dunni subgroup species. Drosophila nigrodunni and D. antillea form a sibling pair, in agreement with their abdominal pigmentation patterns. The cardini group has been studied extensively, primarily because of the fascinating cline in abdominal patterns found in the Caribbean members of the dunni subgroup (Heed and Krishnamurthy, 1959; Hollocher et al., 2000a, 2000b). Hollocher et al. (2000b) discuss some models to account for this color pattern. guarani and guaramunu species groups. Traditionally, the guarani species group contained two subgroups, guarani and guaramunu. Kastritsis (1969), using polytene chromosome banding patterns, divided this group into two separate species groups based on the fact that the guaramunu species group is more closely related to the tripunctata group than either is to the guarani group. However, there is still some confusion over this placement, and Baechli (2005) lists these species in different subgroups of the same species group. Currently, the guarani species group contains six species that are found in Central and South America (Table 1.3). According to Kastritsis (1969), who reviewed all the available data, D. guarani and D. subabdia are sister taxa. The

D. bedichecki D. cardinoides D. parthenogenetica D. procardinoides D. neomorpha D. polymorpha D. cardini D. neocardini D. belladunni D. dunni D. similis D. nigrodunni D. antillea D. arawakana D. caribiana D. acutilabella

dunni subgroup (a)

cardini subgroup

(b)

Figure 1.24. (a) Polytene chromosome phylogeny of the cardini species group, after Heed and Russell (1971); (b) phylogeny of the dunni subgroup (cardini species group) based on mitochondrial DNA sequences, after Hollocher (1998).

Phylogenetic relationships of Drosophilidae

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33

guaramunu species group contains three described species (Table 1.3) that are also South American in distribution. Chromosome data suggest that, of these species, D. griseolineata and D. guaramunu are sister taxa (Kastritsis, 1969). Four additional species are closely allied to these taxa, but an exact position cannot be determined at this time (Table 1.3). Phylogenetic work on these species is sorely needed. immigrans species group. The immigrans group is a large Old World clade of about a hundred described species (Table 1.3 lists only those species present in the Tucson Stock Center). This group has, interestingly, diversified in parallel with the tripunctata species group in the New World. Throckmorton considered this group a member of the immigrans radiation, a lineage basal to the tripunctata radiation (Figure 1.2). Remsen and O’Grady (2002), using molecular characters, placed this taxon basal to all species in their immigrans-tripunctata clade (Figure 1.5). In spite of recent taxonomic reorganization (Okada and Carson, 1983) removing the lineosa subgroup and placing it within the genus Zaprionus (see below), no inclusive phylogenetic hypothesis exists for the immigrans species group. One study (Yu et al., 1999) has recently examined the phylogeny of the nasuta subgroup (Figure 1.25).

D. s. albostrigata KM, ML D. albomicans CHM D. nasuta D. s. neonasuta D. pallidifrons Taxon I/J D. albomicans HK, KM, GZ D. kepulauana D. s. bilimbata nasuta subgroup

Taxon F D. kohkoa D. pulaua D. s. sulfurigaster D. niveifrons D. immigrans

Figure 1.25. Phylogenetic relationships within the nasuta subgroup (immigrans species group) based on mitochondrial DNA sequences (Yu et al., 1999). Taxa I/J and F are putative new species. Abbreviations after species names refer to collection localities: CHM  Chiangmai, Thailand; GZ  Guangzhou, China; HK  Hekou, China; KM  Kunming, China; ML  Miri, Sarawak, Malaysia.

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Within this subgroup, D. niveifrons is basal and sister to a large clade of relatively closely related taxa. Their phylogeny suggests that D. s. sulfurigaster, which is traditionally composed of several subspecies, may not be monophyletic. Instead, each of the three subspecies examined (neonasuta, albostrigata, bilimbata) may be species distinct from the typical D. s. sulfurigaster. These data also conclude that D. albomicans is not monophyletic at the species level, with populations from Chiangmai, China, being quite different from the other three populations sampled. Additional work, taking into consideration the remaining taxa in the immigrans species group, is needed. pallidipennis species group. The pallidipennis species group contains one species, D. pallidipennis, which is found in the Neotropics (Table 1.3). Two subspecies are currently recognized, D. pallidipennis centralis and D. pallidipennis pallidipennis. Pasteur and Kastritsis (1971) have examined the polytene chromosomes of this species. quinaria species group. The quinaria species group is a Holarctic clade containing 32 described species (Table 1.3; Baechli, 2005). This group has served as a model system for understanding the ecology of mycophagous Drosophilidae and, by extension, insect-host (Jaenike, 1978a, 1978b, 1995; Grimaldi and Jaenike, 1984; Jaenike and James, 1991) and insect-parasite associations (Perlman et al., 2003). Spicer and Jaenike (1996) have proposed a phylogeny of the quinaria species group to better understand the evolution of -amanitin (a RNA polymerase II inhibitor which is toxic to species that are not adapted) tolerance and host use in this group (Figure 1.26). They find that D. phalerata and D. falleni are sister taxa, and basal to the remaining quinaria group species. Drosophila palustris and D. subpalustris are sister species within a larger clade also containing D. recens and D. quinaria. Drosophila guttifera occupies an intermediate position on the phylogeny (Figure 1.26).

D. phalerata D. falleni D. subpalustris D. palustris D. recens D. quinaria D. guttifera Figure 1.26. Phylogeny of the quinaria species group (Spicer and Jaenike, 1996).

Phylogenetic relationships of Drosophilidae

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35

These results are very similar to those of Perlman et al. (2003), which include a larger sample of the total number of species in this group. They recover two major clades in the quinaria group (Figure 1.23). The first contains D. falleni and D. phalerata. While not sister taxa, these species do fall out into a large clade containing D. brachynephros, D. unispina, D. curvispina, and D. innubila. The other major clade includes the sibling species D. palustris and D. subpalustris, along with another species, D. deflecta. Drosophila recens and D. quinaria are in a large clade containing D. limbata, D. munda, D. occidentalis, D. suboccidentalis, D. subquinaria, D. transversa, D. nigromaculata, D. guttifera, and D. kuntzei (Figure 1.23). The quinaria group, like several related immigrans-tripunctata species groups, also possesses a diversity of abdominal pigmentation patterns. testacea species group. The testacea species group is Holarctic in distribution. Currently, this group contains four described species (Table 1.3), all of which are mycophagous. Grimaldi et al. (1992) examined the systematic relationships within this clade. They suggested that there were two major lineages, the testacea and putrida subgroups. The testacea subgroup contains three species: D. testacea, D. orientalis, and D. neotestacea. Drosophila putrida is the only species placed in the putrida subgroup. Grimaldi et al. (1992) suggested this placement because of the highly different male genitalia of D. putrida. Perlman et al. (2002) find a similar topology, although their phylogeny is more resolved than the previous work in that it supports the sister group relationship between D. orientacea and D. testacea (Figure 1.23). tripunctata species group. The tripunctata species group was erected by Sturtevant (1942), based on material from two species. Since that time, this group has been divided into four subgroups (Frota-Pessoa, 1954) containing over 70 described species (Baechli, 2005). Table 1.3 lists only those species present in the Tucson Stock Center. The tripunctata species group is mostly Neotropical in distribution, with its center of species diversity in Brazil. Only the nominal species, D. tripunctata, is found in the Nearctic region, where it is widespread in the eastern United States. Kastritsis (1966) examined the polytene chromosomes of some species in this group, but those data are not amenable to coding for a phylogenetic study. In fact, the tripunctata species group is a very large, poorly studied group that has not been extensively studied using explicit phylogenetic methodology. This is unfortunate because, much like the cardini species group, this clade displays a high diversity of abdominal pigmentation patterns.

Genus Drosophila: Hawaiian Drosophilidae The endemic Hawaiian Drosophilidae is among the largest groups in this family, with about 600 described and over 300 undescribed species (O’Grady et al., 2003). This group is slightly unconventional from a taxonomic standpoint, as the names of many species groups are derived from the secondary sexual characters found in males rather than from species names, as is the case with the continental fauna (i.e. the repleta group). This historical artifact is somewhat confusing, but will be adopted here. Also confusing is the fact that the genus Scaptomyza is the sister taxon of the Hawaiian Drosophila lineage, rendering the genus Drosophila paraphyletic (O’Grady, 2002;

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How to look at flies

Nudidrosophila* Ateledrosophila* modified picture wing mouthpart Idiomyia* crassifemur*

antopocerus* modified tarsus

Titanochaeta?

ciliated tarsus

Scaptomyza Grimshawomyia? Celidosoma?

haleakalae drosophiloid scaptoid

Figure 1.27. Phylogeny of species groups placed in the Hawaiian Drosophilidae based on internal anatomy (Throckmorton, 1966).

Remsen and O’Grady, 2002). We include Scaptomyza here because of its phylogenetic affinities, and point out that revisionary work on this genus Drosophila is sorely needed to resolve this issue. With the exception of the picture wing clade, the majority of species groups in this large adaptive radiation are poorly understood with respect to their phylogenetic relationships (see O’Grady 2002 for a recent review). Throckmorton (1966) proposed a phylogeny of the endemic species of Drosophilidae found in Hawaii (Figure 1.27). He recognized two major sister groups; the Hawaiian Drosophila, and the genus Scaptomyza (Table 1.1). The Hawaiian Drosophila are characterized by a high degree of sexual dimorphism, with males having elaborate modifications to their wings, mouthparts, and forelegs. The genus Scaptomyza is a fascinating group, which contains both Hawaiian and continental species. In contrast to the Hawaiian Drosophila, this genus is characterized by males with few secondary sexual characteristics. Instead, Scaptomyza has a high degree of diversity in the external male genitalic apparatus. Throckmorton’s (1966) phylogeny is essentially identical to that proposed by a variety of molecular analyses (Figure 1.28a; Rowan and Hunt, 1991; Thomas and Hunt, 1991; DeSalle, 1992; Kambysellis et al., 1995; Russo et al., 1995; Remsen and DeSalle, 1998; Remsen and O’Grady, 2002). The placement and monophyly of the Hawaiian Drosophilidae has been called into question in some studies (Grimaldi, 1990; reviewed in DeSalle and Grimaldi, 1991, 1992). In Grimaldi’s morphological phylogeny, the Hawaiian Drosophila were not the sister taxon of the genus Scaptomyza (Figure 1.28b). Instead, the Hawaiian Drosophila lineage was included in a clade of mycophagous genera including Hirtodrosophila

Phylogenetic relationships of Drosophilidae

37

Samoaia Zaprionus Sophophora Drosophila Scaptomyza Hawaiian Drosophila Zygothrica Chymomyza Scaptodrosophila (a)

(b)

Figure 1.28. (a) Phylogeny of the Hawaiian Drosophilidae and related groups based on a variety molecular data (DeSalle, 1992; Remsen and DeSalle, 1998); (b) phylogeny of the Hawaiian Drosophilidae and related groups based on Grimaldi’s (1990) morphological analysis.

and its relatives (Figure 1.28b). This was similar to Throckmorton’s Old World Hirtodrosophila radiation (Figure 1.2, discussed above). Grimaldi further suggested that the Hawaiian Drosophila were not closely related to the genus Drosophila at all and represented their own genus, Idiomyia. Scaptomyza was included in a separate lineage, more closely related to the genus Drosophila than the Hawaiian Drosophila. It should be noted that Grimaldi’s (1990) analytical methods have been criticized and his data reanalyzed (O’Grady, 2002; Remsen and O’Grady, 2002; see also above). This reanalysis, while not as resolved as the original morphological hypothesis or the molecular studies, also does not conflict with either. It is therefore clear from all data analyzed that the Hawaiian Drosophila and Scaptomyza form a clade. Within the Hawaiian Drosophila lineage, Throckmorton (1966) proposed that the haleakalae and ciliated tarsus species groups were basal (Figure 1.27). The leaf breeder clade (sensu Heed, 1968), consisting of the modified tarsus and antopocerus groups, was the sister group of a clade containing the modified mouthparts and picture wing groups. Two genera, Ateledrosophila and Nudidrosophila, were placed associated with the picture wing and modified mouthparts groups. These two genera have since been synonymized with the genus Drosophila on the basis of male genitalic morphology (Kaneshiro, 1976). Relationships within the genus Scaptomyza were much less clear, owing to the large number of undescribed species in this group. Recently, several studies have examined the phylogenetic relationships of the Hawaiian Drosophilidae using molecular sequence data (Kambysellis et al., 1995; Baker and DeSalle, 1997; Bonacum et al., in press). These results, with some notable exceptions (discussed below), all agree with those of Throckmorton (Figure 1.29). Kambysellis et al. (1995) focused on the picture wing species, although they did

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picture wing modified mouthpart modified tarsus antopocerus ciliated tarsus fungus breeder Scaptomyza crassifemur Figure 1.29. Summary of the molecular hypotheses of intergroup relationships in the endemic Hawaiian Drosophilidae (Kambysellis et al., 1995; Baker and DeSalle, 1999).

include some modified tarsus, antopocerus, and modified mouthpart species. Baker and DeSalle took a broader taxonomic approach, although with much reduced taxon sampling. Their results were congruent with Throckmorton’s (1966) in that they proposed a basal haleakalae group (Figure 1.29). The leaf breeder clade was sister to the modified mouthpart-picture wing clade. The only drawback of this study was that only three or four taxa from each group were sampled, and several groups (e.g., ciliated tarsus, nudidrosophila) were not sampled at all. Bonacum and colleagues (Bonacum, 2001; Bonacum et al., in press) have greatly improved on both the taxon and character sampling of the Hawaiian Drosophilidae (Figure 1.30). They included multiple representatives of all the major groups of Hawaiian Drosophila, as well as many continental and Hawaiian Scaptomyza species. For the most part, their results were in agreement with the previously proposed notions of phylogeny in this group (Figures 1.27, 1.29). Among the most interesting results of this study is that the nudidrosophila species group seems to make the picture wing clade paraphyletic. This is similar to the placement of this species in Throckmorton’s (1966) phylogeny (Figure 1.27). Additional study of these species will be required in order fully to resolve this placement. Bonacum et al. (in press) also suggested that the ciliated tarsus species group, once considered basal within the Hawaiian Drosophila lineage (Throckmorton, 1966), was actually a member of the modified tarsus group (Figure 1.30). Finally, this study also was the first to use cladistic methodology to address phylogenetic relationships in the genus Scaptomyza. Bonacum’s results not only supported Throckmorton’s assertion that Titanochaeta and Grimshawomyia were closely related to Scaptomyza, but also argued that these taxa be synonymized with Scaptomyza (Bonacum 2001; O’Grady et al., 2003). The placement and phylogeny of Scaptomyza will be discussed below.

•

picture wing clade. The picture wing clade (sensu Kaneshiro et al., 1995) is the largest and most well-studied group in the Hawaiian Drosophila lineage. This group is characterized by males and females with spectacularly pigmented wings, the

Phylogenetic relationships of Drosophilidae

39

D. adunca antopocerus D. longiseta D. diamphidiopoda D. apodasta D. latigena D. septuosa modified tarsus D. petalopeza D. medialis D. cracens modified mouthpart D. adventitia D. primaeva primaeva D. hirtitibia nudidrosophila D. konaensis D. cyrtoloma D. melanocephala D. hanaulae D. ingens D. neoperkinsi D. oahuensis D. neopicta D. nigribasis planitibia D. substenoptera D. differens D. planitibia Picture D. heteroneura Wing D. silvestris D. hemipeza D. picticornis D. setosifrons D. glabriapex D. lineosetae D. ochracea grimshawi D. villosipedis D. inedita D. prolaticilia D. grimshawi D. adiastola adiastola D. aethostoma D. infuscata D. involuta D. kambysellisi modified mouthpart D. soonae D. biseriata D. commatifemora D. hystricosa D. dolichotarsis D. haleakalae D. inciliata D. longiperda haleakalae D. iki D. nigra D. polita D. nigella D. multicilata S. anomala Scaptomyza S. elmoi D. melanogaster Figure 1.30. Molecular phylogeny of the Hawaiian Drosophilidae, after Bonacum et al. (in press).

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How to look at flies

•

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patterns of which may be dimorphic. In spite of the attention this group has received, taxonomic ranks in the picture wings are somewhat confusing. Some authors refer to the entire group as the picture wing species group, while other authors consider the planitibia, grimshawi, and adiastola to be at the species group rank. Kaneshiro et al. (1995) have designated clades within the larger, more inclusive picture wing clade, easing the conflicting situation of a species group within a species group. We follow their convention, as shown in Table 1.1, for all species present in the Tucson Stock Center. Based on polytene chromosome banding patterns and male genitalia, Kaneshiro et al. (1995) have proposed a phylogeny (Figure 1.31) for the picture wing clade. This tree shows that the planitibia clade is sister to the large grimshawi clade (which contains the grimshawi and glabriapex species groups). The adiastola clade is basal to the planitibia-grimshawi clade. Two taxa on the primaeva species group, D. primaeva and D. attigua, are basal to the picture wing clade. planitibia species group. The planitibia species group is a well-studied group of sixteen species in the picture wing clade of the Hawaiian Drosophila (Table 1.1). Initially, this species group was placed in its own genus, Idiomyia, because it possessed an additional cross vein in the wing (Grimshaw, 1901). Carson and Stalker (1968) used polytene chromosomes to demonstrate that this group was actually placed in the picture wing clade. Bonacum et al. (2005) have recently examined the phylogeny of the planitibia species group (Figure 1.32). Their results suggest there are four major lineages within this group, the cyrtoloma, neopicta, picticornis, and planitibia subgroups. The neopicta and cyrtoloma subgroups are sister taxa. The planitibia subgroup, which contains the hypercephalic species D. heteroneura, is sister to the cyrtolomaneopicta clade. The picticornis subgroup is basal in the planitibia species group, a placement that is congruent with morphology in that this subgroup lacks the supernumerary cross vein that characterizes the remainder of this group (Figure 1.32). haleakalae species group. The haleakalae species group is the basal group in the Hawaiian Drosophila lineage. This group is characterized by less dramatic secondary sexual phenotypes in males which, in many ways, are more similar to the genus Scaptomyza, particularly in their mating behaviors. The haleakalae species group has recently been revised (Hardy et al., 2001) and a combined molecular and morphological phylogeny is available (Figure 1.33; O’Grady and Zilversmit, 2004). This phylogeny differs in many ways from the taxonomic framework outlined by Hardy et al. (2001). This is largely due to the dearth of synapomorphic morphological characters that define each species subgroup within the haleakalae group. One notable result in this phylogeny is the close relationship of D. nigra, D. fungiperda, and D. nigella (Figure 1.33). These taxa, placed in two separate taxonomic groups, all have lost the rimmed labellum that is characteristic of the remainder of the haleakalae species group. This phylogeny also suggests a close relationship of two species placed in the venusta cluster, D. venusta and D. longipedis. Finally, two partial subgroups are recovered in the molecular analysis, the polita and haleakalae subgroups (Figure 1.33). This suggests that the taxonomy-based subgroups may actually reflect some historical relationships, but not exactly as proposed by Hardy et al. (2001).

Phylogenetic relationships of Drosophilidae

41 D. hawaiiensis, D. heedi, D. silvarentis D. musaphila, D. gymnobasis, D. recticilia, D. turbata D. gradata D. hirtipalpis, D. psilotarsalis D. flexa, D. formella, D. villitibia D. lasiopoda D. engyochracea, D. balioptera, D. sobrina, D. orthofascia D. murphyi D. ciliaticrus, D. reynoldsiae D. ochracea, D. sejuncta, D. limitata, D. claytonae D. grimshawi, D. affinisdisjuncta, D. disjuncta, D. bostrycha, D. pullipes, D. obatai, D. mulli, D. villosipedis, D. atrimentum, D. orphnopeza, D. crucigera, D. sodomae, D. sproati D. punalua, D. prostopalpis, D. prolaticilia, D. basisetae, D. uniseriata, D. ocellata, D. paucipuncta, D. paucicilia D. virgulata, D. digressa, D. hexachaetae D. spaniothrix, D. macrothrix, D. odontophallus, D. psilophallus, D. tarphytrichia D. gymnophallus, D. liophallus D. discreta, D. lineosetae, D. faciculisetae D. glabriapex, D. pilimana, D. vesciseta, D. aglaia, D. conspicua, D. alsophila, D. micromyia D. assita, D. montgomeryi D. distingueda, D. divaricata, D. inedita

grimshawi

D. differns, D. planitibia, D. heteroneura, D. silvestris D. melanocephala, D. ingens D. oahuensis, D. hanaulae, D. neoperkinsi, D. cyrtoloma, D. neopicta, D. hemipeza, D. nigribasis D.obscuripes, D. substenoptera D. picticornis, D. setosifrons D. setosimentum, D. ochrobasis D. clavisetae, D. neoclavisetae, D. neogrimshawi D. adiastola, D. touchardia, D. spectabilis, D. cilifera, D. toxochaeta, D. peniculipedis D. hamifera, D. paenehamifera, D. varipennis, D. truncipenna D. ornata

Figure 1.31. Chromosome phylogeny of the picture wing clade (Kaneshiro et al., 1995).

planitibia

adiastola

42

How to look at flies

D. hanaulae D. ingens D. cyrtoloma D. melanocephala

cyrtoloma subgroup

D. obscuripes D. neoperkinsi D. oahuensis D. nigribasis D. neopicta

neopicta subgroup

D. substenoptera D. hemipeza D. silvestris D. heteroneura

planitibia subgroup

D. differens D. planitibia D. picticornis

picticornis subgroup

D. setosifrons D. primaeva D. adunca Figure 1.32. Phylogeny of the planitibia species group (picture wing clade) after Bonacum et al. (2005).

Genus Scaptomyza Scaptomyza is a very complex and poorly studied taxon. While several studies strongly support the placement of this genus within the Hawaiian Drosophilidae lineage as the sister taxon of the Hawaiian Drosophila (Figures 1.2, 1.4b, 1.5; Throckmorton, 1966; Okada 1973; Remsen and DeSalle 1998; Remsen and O’Grady, 2002), the monophyly of Scaptomyza is far from certain (Fig. 1.3; Hackman, 1959, 1982; Grimaldi, 1990). Over 150 described species are endemic to the Hawaiian Archipelago. The remaining hundred or so described species of Scaptomyza are placed in nine groups and are found elsewhere. Table 1.1 lists the species present in the Tucson Stock Center. Relationships among and within most of the major lineages of Scaptomyza are not well understood. Okada (1973) proposed a phylogeny of the Scaptomyza based on a phenetic algorithm (Figure 1.34). This tree indicates that the Hawaiian Scaptomyza are not monophyletic, implying more than one colonization of (or migration from) the Hawaiian Islands. Recently, two genera (Titanochaeta and Grimshawomyia) and one

Phylogenetic relationships of Drosophilidae

43

D. melanosoma D. fungiperda

fungiperda complex

D. nigella D. nigra D. fulgida D. dolichotarsis

several complexes, haleakalae subgroup

D. longiperda D. ochropleura D. haleakalae D. canipolita

polita subgroup

D. bipolita D. insignita D. polita D. paraanthrax

iki complex

D. iki D. multiciliata D. scitula D. melanoloma

anthrax subgroup

modified tarsus group modified mouthpart group picture wing group Figure 1.33. Molecular phylogeny of the haleakalae species group (O’Grady and Zilversmit, 2004).

subgenus of Drosophila (Engiscaptomyza) were transferred to Scaptomyza based on molecular and morphological studies (O’Grady et al., 2003). It is clear the further molecular and morphological studies need to be done to resolve additional issues of relationships in this interesting genus.

Genus Drosophila: subgenus Sophophora The subgenus Sophophora was erected by Sturtevant (1939, 1942) when he subdivided the genus Drosophila into subgenera and species groups. This subgenus currently

44

How to look at flies

Bunostomaψ Rosenwaldia* Elmomyza* Alloscaptomyza* Tantalia* Parascaptomyzaψ Macroscaptomyza Metascaptomyza Mesoscaptomyza Dentiscaptomyza Exalloscaptomyza* Hemiscaptomyza Scaptomyza Figure 1.34. Phylogeny of the major lineages within the genus Scaptomyza based on Ohada’s phenetic algorithm. * denotes entire lineages endemic to Hawaii,  indicates lineages where some species are endemic to Hawaii.

includes seven species groups: melanogaster, obscura, saltans, willistoni, dispar, fima, and dentissima (Sturtevant 1939, 1942; Burla, 1954b; Mather, 1954; Tsacas, 1979, 1980), and approximately 300 species (Wheeler, 1982, 1986; Lemeunier et al., 1986). Based on biogeographical data, a common ancestral “protomelanogaster” lineage gave rise to both the melanogaster and obscura species groups in southeast Asia during the mid-Oligocene period (Throckmorton, 1975). The saltans and willistoni species groups comprise a Neotropical clade of species, closely related to one another, but distinct from all other Sophophoran forms (Throckmorton, 1975). Throckmorton (1975) considered the saltans and willistoni species groups to be derivative within the subgenus, originating after the divergence of the melanogaster and obscura species groups from the “protomelanogaster” ancestor. Throckmorton (1975) considered species placed in the subgenus Sophophora to be part of a large radiation of flies that also contained the genera Chymomyza and Neotanygastrella. In his scheme, the “Sophophoran radiation” was basal and gave rise to the remainder of the genus Drosophila (Throckmorton, 1975). Recent molecular studies support the notion that Sophophora is basal within the Drosophilinae and quite distantly related to the subgenus Drosophila (Remsen and DeSalle, 1998; Kwiatowski and Ayala, 1999; Remsen and O’Grady, 2002). Morphology, DNA–DNA hybridization, and phylogenetic analysis of a variety of nucleotide sequences indicate that the subgenus Sophophora is monophyletic (Throckmorton, 1975; Lemeunier et al., 1986; Lachaise et al., 1988; DeSalle, 1992; Thomas and Hunt, 1991; Russo et al., 1995), as are its four largest species groups, melanogaster, obscura, saltans, and willistoni (reviewed in Powell, 1997). Furthermore, these data also support the monophyly of the Old World melanogaster-obscura

Phylogenetic relationships of Drosophilidae

Neotropical Clade

45

saltans group willistoni group (in part) willistoni group (in part) fima group

Old World Clade

obscura group melanogaster group (in part) melanogaster group (in part) Drosophila

Figure 1.35. Summary of Pelendakis et al.’s (1991) phylogeny of Sophophora.

and Neotropical saltans-willistoni clades (Figure 1.35). In all of these studies, the subgenus Sophophora was monophyletic with respect to both the subgenus Drosophila and several genera within the family Drosophilidae. However, two studies, both by Pelendakis and colleagues, have suggested that Sophophora is not monophyletic (Pelendakis et al., 1991; Pelendakis and Solignac 1993). Their results suggested that the melanogaster species group, as traditionally defined, is not a monophyletic lineage. The ananassae subgroup (melanogaster species group) was shown to be the sister taxon of the obscura species group. However, it should be noted that bootstrap support for this relationship is not high. The fima species group is the sister group to the obscura-ananassae clade. The saltans species group was monophyletic in this study, as was the saltans-willistoni clade, but the willistoni species group is paraphyletic with respect to the saltans species group (Figure 1.35). The result that was most incongruent with the previous morphological and molecular studies, however, was the finding that Sophophora was paraphyletic with respect to the subgenus Drosophila. Katoh et al. (2000) also suggested the Sophophora was not monophyletic. Their analysis found that the genus Lordiphosa (this group was once considered a subgenus in Drosophila, but Grimaldi (1990) elevated it to generic rank) was closely related to the Neotropical saltans and willistoni species groups. They further suggested that the Old World obscura and melanogaster lineage was more closely related to the subgenus Drosophila (Figure 1.36), although support for this assertion was quite low. Recently, O’Grady and Kidwell (2002) have used three molecular data sets to examine the phylogeny of the subgenus Sophophora. Their results are not in agreement with those of Pelendakis and Solignac (1991) in that all four of the major species group are monophyletic (Figure 1.37). They did find, however, that the willistoni species group was not monophyletic in some analyses, suggesting that support for this clade is not high (Figure 1.37). However, the study of Remsen and O’Grady (2002), which

46

How to look at flies

Scaptomyza Hawaiian Drosophila robusta group virilis group repleta group polychaeta group Lordiphosa (tenuicauda group) Mycodrosophila Hirtodrosophila immigrans group funebris group quinaria group Liodrosophila obscura group Old World melanogaster group Sophophora Zaprionus Lordiphosa (denticeps, fenestratum, miki groups) willistoni group Neotropical saltans group Sophophora Scaptodrosophila Amiota Leucophenga Chymomyza Figure 1.36. Summary of Katoh et al. (2000), with specific emphasis on the placement of Lordiphosa relative to Sophophora.

sampled extensively within the genus Drosophila, strongly supports the monophyly of Sophophora (Figure 1.5) as a lineage distinct from the genus Drosophila. Instead suggesting Sophophora has affinities with Chymomyza and Scaptodrosophila, two basal drosophilines.

•

melanogaster species group. Drosophila melanogaster is one of the single most important model organisms in biology. The melanogaster species group has, in a similar fashion, served as a model system for studies of speciation, population genetics, molecular evolution, and the evolution of development (Krietman, 1983; Turelli and Orr, 2000; Turelli et al., 2001; Kopp and True, 2002a, 2002b). The first major taxonomic revision of this group, aside from its establishment by Sturtevant (1939), was done by Bock and Wheeler (1972). Since that time, a large number of species have been described for both the African and Asian-Pacific regions (refer to Table 1.4 for those species present in the Tucson Stock Center).

Phylogenetic relationships of Drosophilidae

47

D. mauritiana D. simulans D. melanogaster D. yakuba D. eugracilis D. lutescens

melanogaster

D. tsacasi D. triauraria D. quadraria D. biauraria D. affinis D. azteca D. subobscura D. madeiriensis D. guanche D. bifasciata

obscura

D. miranda D. pseudoobscura D. persimilis D. sturtevanti D. milleri D. saltans D. prosaltans D. lusaltans D. austrosaltans

saltans

D. subsaltans D. emarginata D. neocordata D. nebulosa D. willistoni D. tropicalis D. insularis D. pavlovskiana D. paulistorum D. equinoxialis D. capricorni D. sucinea D. fumipennis Figure 1.37. Phylogeny of Sophophora, based on O’Grady and Kidwell (2002).

willistoni

48 Table 1.4.

How to look at flies Species placed in the subgenus Sophophora

Species group

Species and authority

melanogaster


ananassae* Doleschall, 1858; auraria* Peng, 1937; baimai* Bock and Wheeler, 1972; barbarae* Bock and Wheeler, 1972; biarmipes*; biauraria* Bock and Wheeler, 1972; bicornuta* Bock and Wheeler, 1972; bipectinata* Duda, 1923; birchii* Dobzhansky and Mather, 1961; diplacantha* Tsacas and David, 1978; elegans* Duda, 1927; ercepeae* Tsacas and David, 1975; erecta* Tsacas and Lachaise, 1974; eugracilis* Bock and Wheeler, 1972; ficusphila* Kikkawa and Peng, 1938; greeni* Bock and Wheeler, 1972; jambulina* Parshad and Paika, 1964; kanapiae* Bock and Wheeler, 1972; kikkawai* Burla, 1954; lacteicornis* Okada, 1965; lini* Bock and Wheeler, 1972; lucipennis* Lin, in Bock and Wheeler, 1972; lutescens* Okada, 1975; malerkotliana* Parshad and Paika, 1964; mauritiana* Tsacas and David, 1975; mayri* Mather and Dobzhansky, 1962; melanogaster* Meigen, 1830; mimetica* Bock and Wheeler, 1972; nikananu* Burla, 1954; orena* Tsacas and David, 1978; orosa* Bock and Wheeler, 1972; pallidosa* Bock and Wheeler, 1972; parabipectinata* Bock, 1971; paralutea* Bock and Wheeler, 1972; parvula* Bock and Wheeler, 1972; pennae* Bock and Wheeler, 1972; phaeopleura* Bock and Wheeler, 1972; prostipennis* Lin, in Bock and Wheeler, 1972; pseudoananassae* Bock, 1971; pseudotakahashii* Mather, 1957; pulcherella* Tan, Hsu and Sheng, 1949; punjabiensis* Parshad and Paika, 1964; quadraria* Bock and Wheeler, 1972; rufa* Kikkawa and Peng, 1938; sechellia* Tsacas and Baechli, 1981; seguyi* Smart, 1945; serrata* Malloch, 1927; simulans* Sturtevant, 1919; takahashii* Sturtevant, 1927; teissieri* Tsacas, 1971; triauraria* Bock and Wheeler, 1972; tsacasi* Bock and Wheeler, 1972; varians* Bock and Wheeler, 1972; vulcana* Graber, 1957; yakuba* Burla, 1954

obscura

affinis* Sturtevant, 1916; algonquin Sturtevant and Dobzhansky, 1936; alpina Burla, 1948; ambigua* Pomini, 1940; athabasca Sturtevant and Dobzhansky, 1936; azteca* Sturtevant and Dobzhansky, 1936; bifasciata* Duda, 1923; cariouae Tsacas, in Tsacas et al., 1985; cuauhtemoci Felix and Dobzhansky, in Felix et al., 1976; dobzhanskii Patterson, 1943; eniwae Takada, Beppu and Toda, 1979; epiobscura Parshad and Duggal, 1966; eskoi Lakovaara and Lankinen, 1974; frolovae Wheeler, 1949; guanche* Monclus, 1976; helvetica Burla, 1948; hubeiensis Sperlich and Watabe, in Watabe and Sperlich, 1997; imaii Moriwaki and Okada, in Moriwaki et al., 1967; inexspectata Tsacas, 1988; kitumensis Tsacas, in Tsacas et al., 1985; krimbasi Tsacas, in Tsacas et al., 1985; lowei Heed, Crumpacker and Ehrman, 1968; madeirensis Monclus, 1984; maya Heed and O’Grady, 2000; microlabis Seguy, 1938; miranda* Dobzhansky, 1935; narragansett* Sturtevant and Dobzhansky, 1936; novitskii Sulerud and Miller, 1966; obscura Fallen, 1823; persimilis* Dobzhansky and Epling, 1944; pseudoobscura* Frolova, in Frolova and Asterauv, 1929; seminole Sturtevant and Dobzhansky, 1936; sinobscura Watabe, in Watabe et al., 1996; subobscura* Collin, in Gordon, 1936; subsilvestris Hardy and Kaneshiro, 1968; tolteca* Patterson and Mainland, 1944; tristis Fallen, 1823; tsukubaensis Takamori and Okada, 1983

saltans

austrosaltans* Spassky, 1957; cordata Sturtevant, 1942; dacunhai* Mourao and Bicuda, 1967; elliptica Sturtevant, 1942; emarginata* Sturtevant, 1942; lusaltans* Magalhaes, 1962; magalhaesi Mourao and Bicudo, 1967; milleri* Magalhaes, 1962; neocordata* Magalhaes, 1956; neoelliptica Pavan and Magalhaes, in Pavan, 1950; neosaltans Pavan and Magalhaes, in Pavan, 1950; nigrosaltans Magalhaes, 1962; parasaltans Magalhaes, 1956; prosaltans* Duda, 1927; pseudosaltans Magalhaes, 1956; pulchella Sturtevant, 1916; rectangularis Sturtevant, 1942; saltans* Sturtevant, 1916; septentriosaltans Magalhaes and Buck, in Magalhaes, 1962; sturtevanti* Duda, 1927; subsaltans* Magalhaes, 1956

Phylogenetic relationships of Drosophilidae Table 1.4.

49

(Continued )

Species group

Species and authority

willistoni

abregolineata Duda, 1925; bocainensis Pavan and Cunha, 1947; bocainoides Carson, 1954; capnoptera Patterson and Mainland, 1944; capricorni* Dobzhansky and Pavan, 1950; changuinolae Wheeler and Magalhaes, 1962; equinoxialis* Dobzhansky, 1946; fumipennis* Duda, 1925; insularis Dobzhansky, 1957; megalagitans Wheeler and Magalhaes, 1962; mangabeirai Malogolowkin, 1951; nebulosa* Sturtevant, 1916; neoalagitans Wheeler and Magalhaes, 1962; parabocainensis Carson, 1954; paulistorum* Dobzhansky and Pavan, in Burla, 1949; pavlovskiana Kastritsis and Dobzhansky, 1967; pittieri Baechli and Vilela, 2002; pseudobocainensis Wheeler and Magalhaes, 1962; subinfumata Duda, 1925; sucinea* Patterson and Mainland, 1944; tropicalis* Burla and Cunha, in Burla et al., 1949; willistoni* Sturtevant, 1916

* Species in culture at the Tucson Stock Center;
All taxa in these groups were not listed.

•

Recently, Schawaroch (2002) has generated a molecular phylogeny of this entire group (Figure 1.38). Kopp and True (2002a) have done a more targeted study of the Oriental subgroups (Figure 1.39), a large clade within the melanogaster species group (Ashburner, 1989, 2004). The results of both studies are quite congruent. For example, both suggest that the suzukii subgroup is not monophyletic. Kopp and True’s (2002a) phylogeny indicates that members of the suzukii subgroup form three lineages, two of which are related to the takahashii subgroup and another that is close to the elegans subgroup (Figure 1.39). Schawaroch’s (2002) results are similar. One species, D. lucipennis, is closely related to the elegans subgroup (Figure 1.38). Another suzukii lineage is related to the takahashii subgroup, a result also seen in Kopp and True (2002a). A third lineage, represented by D. biarmipes, shows an affiliation with the eugracilis-melanogaster lineage (Figure 1.38). A point of contention between the two studies is their placement of Drosophila eugracilis. Schawaroch (2002) places this species close to the melanogaster subgroup. Kopp and True (2002a), on the other hand, suggest that D. eugracilis is basal to all the other “Oriental subgroups” and that the melanogaster subgroup is sister to the suzukii-takahashii lineage (Figures 1.38, 1.39). Kopp and True also included the rhopaloa subgroup as the sister taxon of the elegans-suzukii lineage. This species was not surveyed by Schawaroch (2002); instead, she sampled extensively within the ananassae and montium subgroups to give a more holistic picture of evolution within the entire melanogaster species group. The ananassae and montium subgroups are monophyletic sister taxa (Figure 1.38). These two large clades are the sister groups of the so-called “Oriental subgroups”. obscura species group. The phylogenetic relationships of species in the Drosophila obscura species group have been reviewed by O’Grady (1999). The obscura group was initially divided into two lineages by Sturtevant (1942). The affinis subgroup consisted exclusively of New World species, with the obscura subgroup containing species found in both Old and New Worlds. In an early morphological study, Buzzati-Traverso and Scossiroli (1955) concluded that, within the “traditional” obscura subgroup (sensu Sturtevant 1942), there were two distinct lineages of

50

How to look at flies

D. paralutea D. prostipennis takahashii subgroup D. lutescens D. takahashii D. mimetica suzukii subgroup (in part) D. biarmipes D. eugracilis eugracilis subgroup D. melanogaster melanogaster subgroup D. teissieri D. yakuba D. lucipennis suzukii subgroup (in part) D. elegans elegans subgroup D. ficusphila ficusphila subgroup D. ananassae D. pallidosa D. phaeopleura ananassae subgroup D. malerkotliana D. varians D. ercepeae D. greeni D. seguyi D. nikananu D. vuncana D. diplacantha D. tsacasi D. bicornuta D. birchii D. barbarae D. mayri montium subgroup D. serrata D. watanabei D. punjabensis D. orosa D. kikkawai D. lini D. rufa D. quadraria D. triauraria D. auraria D. biauraria D. parvula D. kanapiae D. baimaii Figure 1.38. Phylogeny of the melanogaster species group, after Schawaroch (2002).

Phylogenetic relationships of Drosophilidae

51

D. eugracilis

eugracilis

D. lucipennis

suzukii*

D. elegans

elegans

D. gunungcola D. fuyamai

rhopaloa

D. orena melanogaster

D. teissieri D. melanogaster D. suzukii

suzukii*

D. biarmipes D. lutescens D. trilutea takahashii

D. takahashii D. prostipennis D. pseudotakahashii D. mimetica suzukii* D. ficusphila D. bipectinata

ficusphila ananassae

D. ananassae Figure 1.39. Phylogeny of the melanogaster species group, after Kopp and True (2002a).

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Nearctic species and several lineages of Palearctic species. The obscura group is now divided into five subgroups (Figure 1.40), two in the New World (affinis and pseudoobscura) and three in the Old World (obscura, subobscura, and microlabis). The affinis and pseudoobscura species groups are sister taxa. The affinities of the Old World lineages remain somewhat unclear, although they are considered basal to the New World forms (Figure 1.40). Table 1.4 lists the species of the obscura group. saltans species group. The saltans species group is exclusively Neotropical in distribution (Table 1.4). Magalhaes (1962) and Throckmorton and Magalhaes (1962) proposed five subgroups. The cordata and elliptica subgroups were basal within the saltans group. The subsaltans subgroup was considered the sister group of the clade formed by the more derived sturtevanti and saltans species groups. O’Grady et al. (1998) used four molecular loci to examine the phylogeny of this group. Their results largely agreed with those of Magalhaes (1962). The saltans subgroup, represented by D. saltans, D. lusaltans, D. austrosaltans, and D. prosaltans, was most

52

How to look at flies

D. affinis D. athabasca affinis subgroup

D. algonquin D. azteca D. tolteca D. naragansett D. persimilis D. pseudoobscura D. miranda

pseudoobscura subgroup

D. lowei D. ambigua D. obscura D. tristis

obscura subgroup

D. subsilvestris D. imaii D. bifasciata D. subobscura D. maderiensis

subobscura subgroup

D. guanche D. microlabis

microlabis subgroup

D. kitumensis Figure 1.40. Summary phylogeny of the obscura species group (O’Grady 1999).

derived and sister to the subsaltans subgroup (Figure 1.41). The sturtevanti subgroup formed a clade sister to these derived taxa. The elliptica, represented by D. emarginata, and cordata, represented by D. neocordata, subgroups were most basal in the saltans group (Figure 1.41).

Phylogenetic relationships of Drosophilidae

53

D. lusaltans D. prosaltans saltans subgroup D. saltans D. austrosaltans D. subsaltans

parasaltans subgroup

D. sturtevanti sturtevanti subgroup

D. milleri

elliptica subgroup

D. emarginata D. neocordata

cordata subgroup

Figure 1.41. Phylogeny of the saltans species group (O’Grady et al., 1998).

D. prosaltans D. saltans D. neocordata D. emarginata D. sturtevanti D. subsaltans Figure 1.42. Phylogeny of the saltans species group (Rodriguez-Trelles et al., 1999).

The saltans group phylogeny produced by Rodriguez-Trelles et al. (1999) was quite different. They suggested that the parasaltans subgroup was basal in the saltans group (Figure 1.42). The saltans subgroup was sister taxon to the cordata subgroup and occupied a derived position in the tree. The elliptica and sturtevanti subgroups were successive sister groups to this more derived clade. This topology is not in agreement with the morphological (Magalhaes, 1962), biochemical (Magalhaes and Throckmorton, 1962), or cytological hypotheses (Bicudo, 1973a, 1973b). Such strong and widespread conflict between two phylogenetic hypotheses is unusual in drosophilid systematics (see, however, DeSalle and Grimaldi, 1991, 1992). Rodriguez-Trelles and Ayala (1999) vehemently argued for their topology over the alternative, based largely on the fact that they had employed a maximum likelihood algorithm and used a large, contiguous sequence fragment, the Xdh gene. It should be noted, however, that O’Grady et al. (1998) also used maximum likelihood, and the sum of their four sequence fragments was nearly the same size

54

How to look at flies

D. equinoxialis D. paulistorum/ D. pavlovskiana D. willistoni D. tropicalis D. insularis D. nebulosa

(a)

(b)

Figure 1.43. (a) Phylogeny of the willistoni species group (Gleason et al., 1998); (b) alternative phylogeny of the willistoni species group based on allozyme analyses (Ayala et al., 1974).

•

(in terms of total character and phylogenetically informative characters) as the Xdh gene. One issue that Rodrigues-Trelles and Ayala (1999) fail to mention is that they examined far fewer species (six versus nine) than the previous study. Furthermore, O’Grady et al. (1998) also examined multiple populations from three widespread species – D. sturtevanti, D. prosaltans, and D. emarginata – such that their ingroup contained fourteen terminal taxa (as opposed to six). Reduced taxon sampling in the latter study (Rodriguez-Trelles and Ayala, 1999) could also be the cause for the difference in tree topology. Reconciling these differences by adding additional characters and taxa will help elucidate relationships within the saltans species group. willistoni species group. Like the saltans group, the willistoni species are entirely Neotropical in distribution (Table 1.4). Gleason et al. (1998) have provided an excellent phylogeny of the willistoni species group based on the nucleotide sequences of several genes. Their phylogeny suggests that D. paulistorum is paraphyletic with respect to D. pavlovskiana. They suggest that there is a conflict in species concepts, biological versus phylogenetic, in this case. The D. pavlovskiana-D. paulistorum lineage is the sister taxon of D. equinoxialis (Figure 1.43a). Drosophila tropicalis is sister to this group, in contrast to the results of Ayala et al. (1974), where D. willistoni and D. tropicalis were each other’s closest relatives and sister taxa to the D. paulistorum-D. equinoxialis clade (Figure 1.43b). Drosophila insularis and D. nebulosa occupy the more basal position on the phylogeny (Figure 1.43a).

Genus Hirtodrosophila The genus Hirtodrosophila was initially described as a subgenus in the genus Drosophila (Sturtevant, 1942). In his revision of the family, Grimaldi (1990) formally elevated this to generic rank. Throckmorton (1975) considered this genus a member of the Old World Hirtodrosophila radiation, a group that includes, in addition to the genus Hirtodrosophila and its relatives in the genus Mycodrosophila, the Hawaiian Drosophila and the genus Scaptomyza. Morphological (Grimaldi, 1990) and molecular

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(DeSalle, 1992) analyses suggest that this group is actually quite distinct from the Drosophila and the Hawaiian Drosophilidae, and we treat it as distinct in our discussion (see also Table 1.1). Hirtodrosophila is a cosmopolitan genus, but the highest diversity is concentrated in the tropics. A single species, H. duncani, is in culture at this time (Table 1.1). The majority of species in this genus are mycophagous (Grimaldi, 1987). The Neotropics, for example, most likely contain over one hundred undescribed species. Phylogenetic relationships in this group are not well understood, but Grimaldi (1987) places these close to the genera Zygothrica and Mycodrosophila (Figure 1.22), a relationship that is also supported by some molecular data (Figure 1.5).

Genus Liodrosophila Liodrosophila is a small Old World genus of 65 known species (Baechli, 2005). This group is distinguished by having a shiny, metallic sheen to the ground color of the body. Okada (1974) has revised Liodrosophila and placed this genus in a phylogenetic context (Figure 1.44). He considered this genus to be the sister taxon to another Old World tropical group, Sphaerogastrella. This is in agreement with Grimaldi (1990) and his placement of these two taxa in the Styloptera Genus Group (Figure 1.3). Remsen and O’Grady (2002) also found a close association between Liodrosophila and Sphaerogastrella (along with a third genus, Dettopsomyia). They referred to this grouping as the formosa clade (Figure 1.5) to avoid confusion with the monophyletic genus Styloptera. Although only one species, L. aerea, has traditionally been available in culture at the Tucson Drosophila Stock Center, several Liodrosophila species are culturable (Table 1.1). This taxon represents a unique lineage in the Drosophilidae that is not well studied with respect to its phylogeny and ecology.

Genus Samoaia Samoaia is a small genus of seven described species endemic to the islands of Samoa. These species are unique for their beautifully patterned wings and thoraces. Wheeler and Kambysellis (1966) reviewed this genus, describing three new species. A cytological study of this group (Ellison, 1968) has been done. There are two major lineages: the “black wing group”; and a sister clade containing two described species, S. leonensis and S. attenuata, and a third undescribed species, “afiamalu”. Species relationships among four taxa are shown in Figure 1.45. It is clear from a number of studies that Samoaia is closely related to the genus Drosophila. Grimaldi (1990) placed Samoaia in a clade with the genus Zaprionus (see below), sister to a monophyletic genus Drosophila (Figure 1.3). However, several molecular studies place it either within or sister to the subgenus Drosophila, to the exclusion of the subgenus Sophophora (Figure 1.4). Remsen and O’Grady (2002) suggested that this species was more basal within the Drosophilidae, close to the mycophagous genus Hirtodrosophila (Figure 1.5). It is clear that the placement of this genus needs to be examined in more detail, particularly with respect to the subgenera Drosophila and Sophophora. Only a single species, S. leonensis, is currently in culture in the Tucson Stock Center (Table 1.1), but several others can be raised in the lab (Ellison, 1968; Throckmorton, 1975).

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L. bimaculata L. quadrimaculata L. ornata L. coeruleifrons L. rugulosa L. nitida L. spinata L. madagascarensis L. sinuata L. aerea L. rufa L. divergens L. castanea L. bicolor L. ciliatipes L. fuscipennis L. fuscata L. bifurcata L. submarginalis L. marginifrons L. ceylonica L. globosa L. pusilla L. dictenia L. dimitada L. onchopyga L. varians L. fasciata Figure 1.44. Phylogeny of the genus Liodrosophila (Okada, 1974).

sp. nov. "afiamalu" S. leonensis S. attenuata blackwing group (includes at least one new species and several known taxa) Figure 1.45. Phylogeny of the genus Samoaia based on polytene banding patterns (Ellison, 1968).

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Genus Scaptodrosophila The genus Scaptodrosophila was initially described as a subgenus of Drosophila, and named Pholadoris (Sturtevant, 1939). Grimaldi (1990) formally elevated this group to generic rank and demonstrated that this group actually occupies a basal position within the Drosophilinae (Figure 1.3). This result is in agreement with Throckmorton’s (1975) placement of this group in the basal Scaptodrosophila radiation. Many Scaptodrosophila species possess prescutellar acrostichal setulae (Chapter 2), a character also seen in the basal subfamily Steganinae. The various molecular studies done to date (Figures 1.4, 1.5) support the placement of this genus as basal (see DeSalle and Grimaldi, 1991, 1992; DeSalle, 1992 for further discussion). Scaptodrosophila is a fascinating group that has undergone a large radiation in Australia (Bock and Parsons, 1975), with over half of the roughly 200 species in this genus endemic to this continent. Little phylogenetic work has been done on the genus Scaptodrosophila so the monophyly of this large radiation, or the species groups established within it (Bock and Parsons, 1978), is uncertain at this point. Scaptodrosophila species in the Tucson Stock Center are listed in Table 1.1.

Genus Zaprionus Zaprionus currently contains about 30 described species (Table 1.1). The highest diversity of this genus is found in sub-Saharan Africa, where many species are endemic. Zaprionus is distinctive in that all species possess a number of bright white or silvery stripes extending, in many cases, longitudinally from the fronto-orbital plates down the mesonotum to the scutellum. Zaprionus is a monophyletic group in all analyses to date (Figures 1.2–1.5), although the placement of this taxon is somewhat unclear. Throckmorton (1975) considered Zaprionus a member of the immigrans-Hirtodrosophila radiation (Figure 1.2). Grimaldi (1990) placed Zaprionus as the sister taxon of the genus Samoaia (Figure 1.3) on the basis of his morphological cladistic analyses. Some molecular studies (Figure 1.4) indicate that Zaprionus is close to the subgenus Dorsilopha (above) within a large clade containing many subgenus Drosophila species groups. The recent movement of the lineosa subgroup (immigrans species group) to the genus Zaprionus suggests a close relationship between these taxa (see above). Other studies (Figure 1.5) place Zaprionus as the sister taxon of Dorsilopha and close to the subgenus Sophophora. The placement of Zaprionus (and Samoaia for that matter) is still uncertain. It is clear, however, that both are imbedded within the Drosophila, rendering this large genus polyphyletic.

Conclusions Our understanding of phylogenetic relationships within the family Drosophilidae has increased greatly in the 30 years since Throckmorton’s (1975) review. A number of phylogenetic studies have used both morphology (Okada, 1989; Grimaldi, 1990) and molecules (e.g. DeSalle, 1992; Remsen and DeSalle, 1998; Kwiatowski and Ayala, 1999;

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Tatarenkov and Ayala, 2001; Remsen and O’Grady, 2002) to infer relationships among the many genera in this family. Although no firm consensus exists – and several key relationships are openly contentious – a picture of the evolutionary history of this group is emerging. The largest impediment is that the molecular studies are still woefully inadequate with respect to their taxon sampling. However, some more extensively sampled studies have been published (Remsen and O’Grady, 2002), and additional work is currently underway. Much phylogenetic work in the Drosophilidae today is comparative. Phylogenetic trees of drosophilid species are being employed as necessary tools to understand the evolution of transposable elements (Silva and Kidwell, 2004), parasitism (Perlman et al., 2003), development (Hollocher et al., 2000a, 2000b; Kopp and True, 2002a, 2000b; Gompel and Carroll, 2003), and a number of other phenomena. Within the genus Drosophila, the phylogenetic relationships of over ten species groups (virilis, repleta, quinaria, testacea, saltans, obscura, melanogaster, willistoni, picture wing, cardini, haleakalae), have been published since 1995. In many cases these phylogenies are in agreement with previous work and are serving to refine our notions of evolution at the species level. Further basic systematic work is needed not only to fill in the gaps in our knowledge, but also to build upon the comparative framework that is already in place in this family. The challenge for the next decade will be to increase the holdings of species in the stock center, expanding its taxonomic breadth and focusing on groups that are currently not culturable.

Acknowledgements The following scientists have graciously provided comments on various drafts of this chapter: James Bonacum, Rob DeSalle, Steve Perlman.

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Rodriguez-Trelles, F., Tarrio, R. and Ayala, R. J. (1999). Molecular evolution and phylogeny of the Drosophila saltans species group inferred from the XDH gene. Mol. Phylog. Evol. 13(1), 110–121. Rowan, R. G. and Hunt, J. A. (1991). Rates of DNA change and phylogeny from the DNA sequences of the alcohol dehydrogenase gene for five closely related species of Hawaiian Drosophila. Mol. Biol. Evol. 8(1), 49–70. Russo, C. A. M., Takezaki, N. and Nei, M. (1995). Molecular phylogeny and divergence times of drosophilid species. Mol. Biol. Evol. 12(3), 391–404. Schawaroch, V. (2002). Phylogeny of a paradigm lineage: the Drosophila melanogaster species group (Diptera: Drosophilidae). Biol. J. Linn. Soc. 76, 21–37. Silva, J. C. and Kidwell, M. G. (2004). Evolution of P elements in natural populations of Drosophila willistoni and D. Sturtevanti. Genetics 168(3), 1323–1335. Spicer, G. S. (1988). Molecular evolution among some Drosophila species groups as indicated by two-dimensional electrophoresis. J. Mol. Evol. 27, 250–260. Spicer, G. S. (1992). Reevaluation of the phylogeny of the Drosophila virilis species group (Diptera: Drosophilidae). Ann. Ent. Soc. Am. 85(1), 11–25. Spicer, G. S. (1995). Phylogenetic utility of the mitochondrial cytochrome oxidase gene: molecular evolution of the Drosophila buzzatii species complex. J. Mol. Evol. 41(6), 749–759. Spicer, G. S. and Bell, C. D. (2002). Molecular phylogeny of the Drosophila virilis species group (Diptera: drosophilidae) inferred from mitochondrial 12S and 16S ribosomal RNA genes. Ann. Ent. Soc. Am. 95(2), 156–161. Spicer, G. S. and Jaenike, J. (1996). Phylogenetic analysis of breeding site use and alpha-amanitin tolerance within the Drosophila quinaria species group. Evolution 50(6), 2328–2337. Spicer, G. S. and Pitnick, S. (1996). Molecular systematics of the Drosophila hydei subgroup as inferred from mitochondrial DNA sequences. J. Mol. Evol. 43(3), 281–286. Stalker, H. D. (1966). The phylogenetic relationships of the species in the Drosophila melanica group. Genetics 53, 327–342. Sturtevant, A. H. (1939). On the subdivision of the genus Drosophila. Proc. Natl. Acad. Sci, USA 25, 137–141. Sturtevant, A. H. (1942). The classification of the genus Drosophila with the description of nine new species. Univ. Texas Publ. 4213, 5–51. Tatarenkov, A. and Ayala, F. J. (2001). Phylogenetic relationships among species groups of the virilis-repleta radiation of Drosophila. Mol. Phylogenet. Evol. 21, 327–331. Tatarenkov, A., Zurovcova, M. and Ayala, F. J. (2001). Ddc and amd sequences resolve phylogenetic relationships of Drosophila. Mol. Phylogenet. Evol. 2, 321–325. Thomas, R. H. and Hunt, J. A. (1991). The molecular evolution of the alcohol dehydrogenase locus and the phylogeny of Hawaiian Drosophila. Mol. Biol. Evol. 8, 687–702. Throckmorton, L. H. (1962). The problem of phylogeny in the genus Drosophila. Univ. Texas Publ. 6205, 207–344. Throckmorton, L. H. (1966). The relationships of the endemic Hawaiian Drosophilidae. Univ. Texas Publ. 6615, 335–396. Throckmorton, L. H. (1975). The phylogeny, ecology, and geography of Drosophila. In: Handbook of Genetics (R. C. King, ed.), Vol. 3, pp. 421–469. Plenum, New York. Throckmorton, L. H. (1982). The virilis species group. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thmopson and H. L. Carson, eds), Vol. 3b, pp. 421–469. Academic Press, New York. Throckmorton, L. H. and Magalhaes, L. E. (1962). Changes with evolution of pteridine accumulations in species of the saltans group of the genus Drosophila. Univ. Texas Publ. 6205, 489–505. Toda, M. J. (1986). Drosophilidae (Diptera) in Burma. Kontyu 54(4), 634–653.

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How to look at flies Tosi, D. and Pereira, M. A. Q. R. (1993). Karyotypes and phylogenetic relationships in the Drosophila species of the annulimana group (Diptera: Drosophilidae). Rev. Bras. Genet. 16, 321–333. Tsacas, L. (1979). Contribution des donnees africainesa la comprehension e la biogeographie et de l’evolution du sous-genre Drosophila (Sophophora) Sturtevant (Diptera: Drosophilidae). Compte rendu des séances de la Societé de biiogeographie 48, 29–51. Tsacas, L. (1980). Les groupes d’especes du sous-genre Sophophora Sturtevant (Diptera, Drosophilidae, Drosophila) et le role du fonctionnement des genitalias males dans la definition des taxons supraspecifiques. Bull. Soc. Zool. Fr. 105(4), 529–543. Turelli, M. and Orr, H. A. (2000). Dominance, epistasis and the genetics of postzygotic isolation. Genetics 154, 1663–1679. Turelli, M., Barton, N. H. and Coyne, J. A. (2001). Theory and speciation. Trends Ecol. Evol. 16, 330–343. Val, F. C. D. and Marques, M. D. (1984). Drosophilidae (Diptera) from the Pantanal of Mato Grosso (Brazil) with the description of a new species belonging to the bromeliae group of the genus Drosophila. Pap. Avul. Zool. 39(11), 223–230. Vilela, C. R. (1983). A revision of the Drosophila repleta species group (Diptera: Drosophilidae). Rev. Bras. Entomol. 27, 1–114. Vilela, C. R. (1984). Notes of the type-series of Drosophila coffeata Williston, with description of a new neotropical species of the subgenus Drosophila (Diptera, Drosophilidae). Revta. Bras. Eng. 28(1), 59–64. Ward, B. L. and Heed, W. B. (1970). Chromosome phylogeny of Drosophila pachea and related species. J. Hered. 61, 248–258. Wasserman, M. (1982). The repleta species group. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3b, pp. 61–140. Academic Press, New York. Wasserman, M. (1992). Cytological evolution of the Drosophila repleta species group. In: Drosophila Inversion Polymorphism (C. B. Krimbas and J. R. Powell, eds), pp. 455–552. CRC Press, Boca Raton, FL. Wheeler, M. R. (1968). Some remarkable new species of Drosophila. Univ. Texas Publ. 6818, 431–442. Wheeler, M. R. (1982). Drosophilidae: a taxonomic overview. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3b, pp. 1–105. Academic Press, New York. Wheeler, M. R. (1986). Additions to the catalog of the world’s Drosophilidae. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3e, pp. 395–409. Academic Press, New York. Wheeler, M. R. and Kambysellis, M. P. (1966). Notes on Drosophilidae (Diptera) of Samoa. Univ. Texas Publ. 6615, 533–563. Yu, H., Wang, W., Fang, S. et al. (1999). Phylogeny and evolution of the Drosophila nasuta subgroup based on mitochondrial ND4 and ND4L gene sequences. Mol. Phylog. Evol. 13(3), 556–565.

CHAPTER 2

Morphological characters Contents

• • • • •

Head Thorax Abdomen Glossary of morphological terms References

The adult fly can be divided into three major regions: head, thorax, and abdomen (Figure 2.1). Within each of these regions a variety of structures have been described and studied (McAlpine, 1981), and are useful characters for identification of Drosophilidae. Reviews of drosophilid morphology have been done by Sturtevant (1916, 1921, 1939, 1942), Duda (1935), Ferris (1950), Wheeler (1952, 1982), and Grimaldi (1987, 1990). The flybase website is an excellent resource (http://flybase.bio.indiana.edu/.bin/fbimage), and has a number of additional images.

Head The drosophilid head is a complex structure which possesses a number of taxonomically important characters. The shape of the head itself is often important. A number of taxa in this family are hypercephalic, and have conspicuously broadened heads (Grimaldi and Fenster, 1989; Figure 2.2). Males of these species typically engage in territorial guarding behavior, and will often joust or head-butt other males attempting to enter their space. This is the case in the well-known Hawaiian Drosophila species, D. heteroneura (Figure 2.3). The antennae consist of a number of segments, the apical-most of which is narrowed into an arista (Figure 2.4). The arista typically has a number of long dorsal and ventral branches, or rays. Arista with multiple long branches are often referred to as plumose. This is typical of many species in the genus Drosophila. Those arista without branches are pubescent. Several genera in the subfamily Steganinae, such as some species of Gitona and Amiota, have pubescent arista. The antopocerus species group of the Hawaiian Drosophila is unique in that it has an elongate arista with numerous fine hairs (Figure 2.4). The chaetotaxy of the third antennal segment is also important in differentiating between some genera. Hirtodrosophila, for example, possesses a third antennal segment with elongate setae.

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Figure 2.1. Dorsal view of a typical drosophilid showing head, thorax, and abdomen (from Grimaldi, 1987). The following morphological characters are further defined in the glossary: costal vein (C), crossveins (h, CuA2, dm-cu, r-m), dorsocentral setae (dc), fronto-orbital plate (fop), humeral setae (hs), subcostal vein (Sc), tergites (tII-VI), wing veins (R1, R2
3, R4
5, M1, CuA1, A1, A2). Courtesy of American Museum of Natural History.

The fronto-orbital plates, which are situated above the compound eye, bear three orbital setae; one proclinate and two reclinate (Figure 2.5). The ocellar triangle, which is located in the dorsomedial region of the head, also has a pair of anteriorally directed setae. Behind the ocellar triangle, on the posterodorsal margin of the head, are the postocellar setae. The inner and outer vertical setae (Figure 2.6) are on the posterolateral margins of the head. The relative placement and length of the orbital, ocellar, vertical, and postocellar setae are all important characters used to diagnose drosophilid genera. The genus Drosophila is characterized by three elongate orbitals, with the proclinate being inserted anterior-most (Figure 2.6). In contrast, the anterior reclinate is inserted anterior to the proclinate in the genus Chymomyza (Figure 2.5). The genus Liodrosophila is different still, with the anterior reclinate setae being minute or absent. The face of most drosophilid species is not prominent, and tends to be either flat or only slightly carinate. Some taxa, such as the genus Zygothrica, have very prominent, nose-like carina which differentiate them from all other species in this family. The shape of the carina can be flat, rounded, or sulcate. The carina is usually wider basally and the ratio of the widths at top and bottom is sometimes used to identify species. The mouthparts of most drosophilids are not ornate (Figure 2.6). Some Hawaiian taxa, however, possess elaborate modifications to their mouthparts, particularly the labellum

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Figure 2.2. Some hypercephalic species of Drosophilidae (from Grimaldi and Fenster, 1989). Courtesy of American Museum of Natural History.

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Figure 2.3. Hypercephaly and male–male sparring behavior in Drosophila heteroneura (photograph courtesy of K. Y. Kaneshiro).

Figure 2.4. Antennae of Drosophila adunca, a species in the Hawaiian antopocerus group. The image is taken at 110 and shows the first, second and third antennal segments, with the arista modified arista inserted into the third segment.

Figure 2.5. Head of Chymomyza amoena, showing the placement of the proclinate (po) and reclinate (aro, pro) orbital setae. The position of the ocellar triangle (ot) and the inner and outer vertical (iov) setae are also shown.

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Figure 2.6. Head of D. immigrans, showing the placement of the proclinate (po) and reclinate (aro, pro) orbital setae and the ocellar triangle (ot).

(Figure 2.7). The color and chaetotaxy of the palps is often diagnostic of different species, and is most often used to diagnose very closely related species (O’Grady et al., 2003a). A number of oral setae are located on either side of the face. The first oral setae is typically the largest and is known as the vibrissae. The second oral (or subvibrissal) setae is also important in species identification. The number and relative size of the oral setae are diagnostic for many species. The compound eye is usually red in the Drosophilidae, and the pilosity is sometimes used to differentiate between closely related forms. The gena, or cheeks, are located on the ventral margin of the eye, and their width relative to the eye is sometimes used to distinguish between closely related species, such as D. melanogaster and D. simulans.

Thorax The thorax is comprised of three fused segments; the prothorax, mesothorax, and metathorax (Figure 2.8). The prothorax and metathorax are very reduced relative to the mesothorax, which has been greatly enlarged to accommodate the musculature that powers the wings. The humeral callus, which bears a pair of taxonomically important setae, is located on the prothoracic segment (Figure 2.8). The mesonotum covers the majority of the dorsal surface of the mesothoracic segment (Figures 2.8, 2.9). There are several important setae on the mesonotum, including several rows of acrostichal setulae and a number of paired dorsocentral setae (Figure 2.9). The number and disposition of the acrostichals is very important to species identification. Most Drosophila possess either six or eight regular rows. The majority of species in the genus Scaptomyza, however, only have two to four rows. Many larger species of Drosophila, as well as other genera, are characterized by having ten or more irregular rows. Several taxa, such as the genus Scaptodrosophila and some species in the annulimana group, also possess two or four enlarged acrostichal setae in the posterior-most row. These are referred to as prescutellar setae, and they have evolved independently in several places in the Drosophilidae.

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Figure 2.7. Mouthpart diversity in the endemic modified mouthpart species group of Hawaiian Drosophilidae (from O’Grady et al., 2003b).

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dc

Mn

Sc

W

Hu

Ha

H

Ms

C1

Abd St C2

C3

Figure 2.8. Thorax of a typical drosophilid (lateral view), showing the major division of this body structure (from Sturtevant, 1916). Abbreviations are as follows: abdomen (Abd), coxae (C1–3), dorsocentral setae (dc), haltere (Ha), head (H), humeral callus (Hu), mesonotum (Mn), mesopleuron (Ms), scutellum (Sc), sternopleuron (St), wing (W).

hs

acr sta

dc

p acr s

7

96

1 ma ide

b sctl s a sctl s Figure 2.9. Dorsal view of P. guttatus (from Wheeler, 1982), showing the following important chaetotaxic characters of the thorax: acrostichal setulae (acr sta), apical scutellar setae (ap sctl s), basal scutellar setae (b sctl s), dorsocentral setae (dc), humeral setae (hs), prescutellar acrostichal setae (p acr s). Source: Manual of Nearctic Diptera, Volume 2, Agriculture and Agri-Food Canada, 1987. Reproduced with the permission of the Minister of Public Works and Government Services Canada, 2005.

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fe ti

bt II III IV V Figure 2.10. Legs of Drosophila, showing the femur (fe), tibia (ti), and five tarsal segments (bt, II–V).

More rarely, taxa will have a set of enlarged acrostichals positioned more anteriorly, close to the transverse suture. These are referred to as presutural setae, and can be found in some species of the mycophagous tripunctata and testacea species groups. Directly posterior to the mesonotum is the scutellum. The orientation of the anterior and posterior scutellar setae (convergent, divergent, or parallel) is important in species identification (Figure 2.9). The katepisternum, referred to by some authors as the sternopleuron, bears a number of katepisternal setae, the number and relative length of which are important – particularly in the genus Drosophila – for defining species and species groups (Figure 2.8). The so-called “sterno index” is the ratio of the anterior to the posterior katepisternal setae. Drosophila willistoni, for example, is characterized by having a very low sterno index of about 0.3. A pair of halteres, or balancers, are located on the metathoracic segment (Figure 2.8). The color of the halteres is often important in differentiating species. Each segment bears a single set of legs. The main leg segments are the coxa, trochanter, femur, tibia, and tarsus (Figure 2.10). There are five tarsal segments. The coloration and chaetotaxy of the legs is widely used in drosophilid systematics. The forelegs of males, in particular, often bear characteristic structures which are useful for species level identification. In addition to chaetotaxy and coloration, the relative lengths of the leg segments are often very important in species identification. The melanogaster and obscura groups, for example, are characterized by having sex combs (Figure 2.11). Other species, such as members of the genus Chymomyza and the immigrans species group, have rows of femoral spines (Figure 2.12). Zaprionus tuberculatus, as well as several other taxa in the genus Zaprionus, have enlarged tubercles on their forelegs (Figure 2.13).

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Figure 2.11. Sex combs of Drosophila ananassae of the melanogaster species group.

Figure 2.12. Foreleg of Drosophila immigrans, showing femoral spines.

Perhaps the greatest diversity of foreleg modification comes in the modified tarsus species group of Hawaiian Drosophila. Members of this clade have spoon-like structure, thickened setae, or are missing an entire tarsal segment – an unusual condition in the Diptera. The wings are attached to the mesothoracic segment (Figure 2.14), and are rich in taxonomic characters. Coloration of the wings can be hyaline, variously spotted (e.g.,

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Figure 2.13. Foreleg of Zaprionus sepsoides showing femoral tubercule. hum brk

R1

sc brk R2+3 r-m

bm dm CuA2

dm-cu

R4+5 M1

cup A1 CuA1

Figure 2.14. Wing of Stegena coleoptrata showing typical venation for this genus and the following important wing characters in the Drosophilidae: crossveins (bm-cu, dm-cu, r-m, and CuA2), humeral break (hum brk), subcostal break (sc break), wing cells (bm, dm, cup), wing veins (R1, R2
3, R4
5, M1, CuA1, A1).

Figure 2.1), or fuscous. Many groups, such as the Hawaiian picture wing and calloptera species groups, are characterized by magnificently patterned wings (Figures 2.15–2.17). The venation pattern is mostly constant, but some taxa, such as the planitibia species group of Hawaiian Drosophila, have additional cross veins (Figure 2.16).

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Figure 2.15. Wing of Drosophila adiastola.

Figure 2.16. Wing of Drosophila heteroneura.

Figure 2.17. Wing of Drosophila calloptera.

Abdomen The entire dorsal and lateral surface of the abdomen, or dorsolateral plate, is a heavily chitinized region. The dorsolateral region of each abdominal segment is referred to as a tergite. The tergites telescope out, growing smaller as they extend posteriorly, and overlap slightly such that the posterior portion of each segment overlaps the anterior portion of the next most posterior segment (Figure 2.18). Tergites 2–9 and 2–8 are apparent in males and females, respectively (Grimaldi, 1987). The pigmentation patterns of the tergites are taxonomically important, especially between closely related species in the repleta (Patterson 1943, Patterson and Mainland 1944), cardini (Heed and Krishnamurthy 1959), tripunctata (Frota-Pessoa, 1954) and other subgenus Drosophila species groups. The ventral region of each abdominal segment is called a sternite, or ventral plate. There are six hairy, chitinized, quadrilaterally-shaped sternites visible in the female, and four visible in the male. In females the five anterior sternites are similar in shape but the sixth is narrower, with a deep notch in one side. The three anterior sternites in males are like those of the female, but the fourth is larger and broader than the others.

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Figure 2.18. Tergites of Drosophila immigrans.

The internal and external male terminalia provides many characters useful for the identification of species. Tergite 9 in males is referred to as the epandrium (Figure 2.19). It may possess a number of characteristic ventral epandrial lobes, sometimes referred to as paralobes. The chaetotaxy of the ventral epandrial lobes is quite variable in Drosophilidae, some taxa may possess thin setulae, larger setae, or even distinct spines on this structure. Posterior to the epandrium are the paired cerci, or anal plates. Posteroventral to the epandrium are the paired surstyli, each of which may bear a number of distinctive prensisetae (Figure 2.19). Grimaldi (1987) has suggested that the surstyli may actually be tergite 10 or 11. A number of drosophilid groups, such as the melanogaster species group and the genus Scaptomyza, have elaborate modifications to their surstyli, epandrial lobes, cerci, or combinations or these. In the case of Scaptomyza, these modifications are diagnostic for the genus and the component subgenera (O’Grady et al., 2003b). The genitalia are the structures associated with sternite 9, or the hypandrium. The posterolateral regions of the hypandrium are typically lobate, and are referred to as gonopods (Figure 2.20). The paraphyses are posterolateral to the gonopods, and often have a number of setae or setulae. The aedeagus, or penis, is positioned centrally with respect to the remainder of the hypandrium. It is divided into two parts, the basiphallus (or shaft) and the distiphallus. The distiphallus may have a number of spines or setulae, and is often sclerotized to a greater or lesser degree. The shape and size of the distiphallus is highly variable among species.

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Figure 2.19. Male terminalia of a typical drosophilid (from Grimaldi, 1987) showing the cerci (ce), epandrium (ep), prensisetae (prs), and surstyli (sry). Courtesy of American Museum of Natural History.

Figure 2.20. Internal male genitalia of a typical drosophilid showing the relative position of the hypandrium (hyp) and distiphallus (ds). Courtesy of American Museum of Natural History.

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Figure 2.21. Ovipositor of Drosophila aenicta.

The female terminalia has not traditionally been used in species identification (however, see Hardy et al., 2001), even though it is highly variable between closely related taxa (Throckmorton, 1966; Kambysellis, 1993). The major external structure is the ovipositor (Figure 2.21). This is a paired structure which bears a number of ovisensilla, the placement and size of which are quite variable and are useful in the identification of species. The shape and coloration of the ovipositor plates themselves are also good species-level characters.

Glossary of morphological terms acrostichal setulae (acr sta; Figure 2.9). Small setulae found on the mesonotum (Mn; Figure 2.8). A series of rows runs from the anterior of the mesonotum to the scutellum (Sc; Figures 2.8, 2.9). The number of rows is usually counted between the anterior dorsocentral setae (dc), as they can be irregular more posteriorly. The number of acrostichal setae typically ranges from zero to twelve rows in Drosophilidae. anal plate. See cercus. antennae. This jointed appendage is attached to the anterodorsal margin of the head. The three main parts are the basal segment (or first antennal segment), pedecil (or second antennal segment), and first flagellomere (third antennal segment). The distal region of the first flagellomere is referred to as the arista. anterior dorsocentral setae. See dorsocentral setae. anterior reclinate orbital setae (aro; Figures 2.5, 2.6). Head setae inserted into the fronto-orbital plate (fop; Figure 2.1) and directed posteriorally. See orbital setae. anterior (or basal) scutellar setae (b sctl s; Figure 2.9). Large setae located at the margins of the anterior end of the scutellum (Sc; Figures 2.8, 2.9), near the base of the triangle. These can be convergent, divergent, parallel or cruciate in orientation. See scutellum.

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arista. Distal portion of the third antennal segment. This structure is typically plumose, with several long dorsal and ventral branches (rays), in the genus Drosophila. It may, however, also be bare or micropubescent in some clades. Ends in a terminal fork, which is usually not counted when the dorsal and ventral branches are added up for species identification. basitarsus. First tarsal segment, where tarsus is joined to the tibia. See leg. carina. Nose-like structure on the face of the fly. This can be prominent and sharply pointed, sulcate medially, or rounded. The ratio between the width of the carina at the base and at the top is often used to described species. cercal clasper. Enlarged setae on cercus. Often called the secondary clasper. cercus (ce; Figure 2.19). Plate associated with the anal region of the fly. Bears the secondary claspers in some taxa. clasper. Both primary (surstylus) and secondary (cercal clasper) are referred to by various authors. claws. Modified setae at the tips of the pretarsus. convergent. Refers to scutellar setae which angle towards one another but do not cross. See anterior and posterior scutellar setae. costal index. Length of the second costal section divided by the length of the third costal section. See wing. coxa (or coxae, C1–3; Figure 2.8). Attachment point between the leg and the thorax. See leg. cruciate. Refers to scutellar setae which cross. See anterior and posterior scutellar setae. divergent. Refers to scutellar setae which angle away from one another. See anterior and posterior scutellar setae. dorsocentral setae (dc; Figures 2.1, 2.8, 2.9). Large setae located on the posterolateral region of the mesonotum (Mn; Figure 2.8), anterior to the scutellum (Sc; Figure 2.8). Typically, there are two sets of dorsocentrals, anterior and posterior, although this number ranges between one and four sets in Drosophilidae. epandrium (ep; Figure 2.19). Part of the male terminalia corresponding to tergite 9. May have a number of ventral lobes, or extensions, as well as a number of setulae, setae or spines. face. Area on the anterior of the fly, between the antennae and oral cavity. See also carina. femur. Third major division of the leg, located between the trochanter and tibia. See leg. first antennal segment. Segment joining the head to the remainder of the antennae. Also referred to as the basal antennal segment. See antennae. frons. Region on the anterodorsal portion of the head, located between the frontoorbital plates. A number of setulae may be inserted into this region. Coloration and pollinosity are important characters. fronto-orbital plate (fop; Figure 2.1). Sclerite located on the dorsolateral margin of the head, directly above the compound eyes. Bears the orbital setae (po, aro, pro; Figures 2.5, 2.6). See head.

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gena. The cheek, located along the ventral margin of the eye. gyres. Twists of the testes. halteres (Hg; Figure 2.8). Also referred to as balancers by some authors. These are knob-like projections attached to the metanotum. Possibly homologous to the hind wing of other insects. Haltere color is often used to distinguish between species. head. Most anterior body segment of the fly. Comprised of a complex series of sclerites including the ocellar triangle, fronto-orbital plate, frons, compound eye, gena, face, clypeus, and mouthparts. humeral setae (hs; Figures 2.1, 2.9). Between one and three setae are typically located on the humerus (Hu; Figure 2.8). Also referred to as postpronotal setae. See humerus. humerus (or humeral callus; Hu; Figure 2.8). Structure located on anterolateral region of the thorax. The humeri are the only prothoracic structures visible dorsally; each appears as a small hump on the anterior portion of the thorax. There are typically between one and three humeral setae (hs; Figures 2.1, 2.9), in addition to several other small hairs on this sclerite. hyaline. Clear, without pigmentation. hypandrium. Part of the male terminalia corresponding to sternite 9. This structure is the male genitalia. katepisternal setae. Setae located on the katepisternum. These vary in number in Drosophilidae from none to five, although most taxa have three. The sterno index is calculated using the relative lengths of these setae. katepisternum. Pleural sclerite located directly above the coxa of the second leg. See mesonotum, katepisternal setae, and sterno index. legs. The legs of Drosophilidae consist of a number of separate structures, including the coxae, trochanter, femur, tibia, tarsus, and pretarsus. The coxa serves as the point of attachment between the leg and thoracic segments. The trochanter is a small second segment between the coxa and femur. The femur and tibia are long segments bearing a number of taxonomically diagnostic setae or, in some groups, specialized structures. The tarsus of most drosophilid species includes five segments. The first tarsal segment is often referred to as the basitarsus. The tarsal segments are typically shorter than the femora and tibia, but also bear important chaetotaxic characters. The pretarsus is the distal-most segment and is composed of a pair of small curved claws and a pulvillus. longitudinal. Oriented along the long axis of the tarsus. Refers to the sex combs present in the melanogaster and obscura species groups. mesonotum (Mn; Figure 2.8). The dorsal region of the mesothorax, extending along the dorsum of the fly from the joint with the head (H; Figure 2.8) to the scutellum (Sc; Figure 2.8). mesothorax. Largest thoracic segment. Mesothoracic parts include the mesonotum and mesopleuron, these two parts being divided by the notopleural suture. The pleural suture divides the mesopleuron into the episternum (anterior) and epimeron (posterior). The episternum is split into the anepisternum (dorsal) and katepisternum (ventral) by the anapleural suture. The katepisternum is referred to as the sternopleuron by some authors. The sterno index is an important measure in the identification of Drosophilidae. The transepimeral suture divides the epimeron into the anepimeron

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(dorsal) and katepimeron (ventral). The anepisternum and anepimeron are always bare in Drosophilidae. The wing is attached directly above the anepimeron. micropubescent. Refers to an arista with either a few short branches or no branches at all. mouthparts. Feeding apparatus of the fly. Also used in mating in some taxa. Includes the labium, labrum, and labellum. ocellar setae. Pair of anteriorally directed setae typically located in the ocellar triangle. Some taxa have ocellar setae which have been displaced and are inserted anterior to the ocellar triangle. ocellar triangle (ot; Figures 2.5, 2.6). This region is located posteromedially on the head. It bears three ocelli and a pair of anteriorally directed ocellar setae. oral setae. Setae directly flanking either side of the face. The first oral, or vibrissae, is usually the largest. orbital setae. Three orbital setae, one proclinate (proclinate orbital setae) and two reclinate (anterior and posterior reclinate orbital setae), are inserted into the fronto-orbital plate. The placement and length of these setae relative to one another is important in delineating closely related taxa. These setae are sometimes numbered 1–3, starting with the posterior reclinate. parallel. Refers to scutellar setae which are parallel to one another. See anterior and posterior scutellar setae. paralobes. Ventral lobes on the epandrium. pilose (pilosity). Refers to the density of short hairs (pile) on the eye. Many densely pilose species appear to have coarse, fuzzy eyes, while other taxa appear smooth. pleurae. A complicated number of fused plates along the “sides” of the thorax, directly ventral to the mesonotum. plumose. Having many long-rayed branches. See arista. pollinose. Appearing to be covered with pollen. Usually used to refer to the frons or mesonotum. Degree of pollinosity is an important character used to differentiate between some species. posterior reclinate orbital setae (pro; Figures 2.5, 2.6). Head setae inserted into the fronto-orbital plate (fop; Figure 2.1) and directed posteriorly. See orbital setae. posterior (or apical) scutellar setae (ap sctl s; Figure 2.9). Large setae located at the margins of the posterior end of the scutellum (Sc; Figures 2.8, 2.9), near the point of the triangle. These can be convergent, divergent, parallel or cruciate in orientation. See scutellum. postocellar setae. Set of setae inserted on the posterodorsal margin of the head, directly posterior to the ocellar triangle. prensisetae (prs; Figure 2.19). Setae, or teeth, on the surstyli (sry; Figure 2.19). Typically arranged in a number of regular or irregular rows. Some authors refer to these as clasper teeth or the clasper comb. prescutellar acrostichal setae (p acr s; Figure 2.9). Some drosophila taxa have one or two pairs of prescutellars located between the posterior dorsocentral setae (dc; Figures 2.1, 2.8, 2.9). These are inserted into the mesonotum (Mn; Figure 2.8).

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pretarsus. Sixth and final division of the leg. Located at the apex of the leg, attached to fifth tarsomere. See leg. primary clasper. See surstylus. proclinate orbital setae (po; Figures 2.5, 2.6). Head setae inserted into the fronto orbital plate (fop; Figure 2.1) and directed anteriorally. See orbital setae. prothorax. Anterior-most thoracic segment, located where the head is joined to the thorax. Prothoracic parts include the humerus and proepisternum. pulvillus. Fine, hair-like structure at the apex of the pretarsus. See leg. sclerite. Chitinous body part or segment. scutellar setae. See anterior and posterior scutellar setae. scutellum (Sc; Figures 2.8, 2.9). Triangular structure on the dorsum of the fly, directly posterior to the mesonotum (Mn; Figure 2.8). This structure is typically bare of small hairs, but always bares two sets of distinct marginal setae, the anterior (b sctl s; Figure 2.9) and posterior (ap sctl s; Figure 2.9) scutellar setae. second antennal segment. Middle segment of the antennae. Typically bears several stout setae, in addition to numerous short setulae. Also referred to as the pedicel. See antennae. sex combs. Modified setae located on the tarsal segments of some species in the subgenus Sophophora. The number of combs and the number of setae (or teeth) per comb varies between species. sternite. Ventral region of each abdominal segment. sterno index. Length of the anterior katepisternal setae divided by the length of the posterior katepisternal setae. This measure is often used to separate clades within the genus Drosophila. sternopleural setae. See katepisternal setae. sternopleuron (St; Figure 2.8). See katepisternum. sulcate. Depressed or caved in. surstylus (sry; Figure 2.19). Paired structure located posteroventrally to the epandrium (ep; Figure 2.19). Often bears a number of presisetae (prs; Figure 2.19). This structure is used in mating to grasp and position the female. Often referred to as the primary clasper. tarsomere. A segment of the tarsus. See also leg. tarsus. Fifth major division of the leg, located between tibia and pretarsus. Typically contains five distinct segments, or tarsomeres. See leg. tergites (tII-IV; Figure 2.1). Dorsolateral region of each abdominal segment. Tergites one and two are fused to form syntergite 1
2. The tergites telescope out, growing smaller as they extend posteriorly, and overlap slightly such that the posterior portion of each segment overlaps the anterior portion of the next most posterior segment. The pigmentation patterns of the tergites are taxonomically important, especially between closely related species in the repleta, cardini, tripunctata, and other subgenus Drosophila species groups.

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terminal fork. Apical-most bifurcation of the arista. testes. Coiled, sperm-producing organs found in the male. Many species have testes which are visible through the tissue of the abdomen. The color and number of coils (gyres) are diagnostic at the species level. third antennal segment. Apical-most segment of the antennae. Also referred to as the first flagellomere. Distal region of this segment is called the arista. See antennae. thorax. Large, complex body part composed of three fused segments – the prothorax, mesothorax, and metathorax – each of which bears a single set of legs. Wings attach to the mesothoracic segment, and the halteres are located on the metathoracic segment. tibia. Fourth major division of the leg, located between the femur and tarsus. See leg. transverse. Oriented either perpendicular or diagonal to the long axis of the tarsus. Refers to the sex combs present in the melanogaster and obscura species groups. trochanter. Second leg segment, between coxa and femur. See leg. unicolorous. All one color, without distinct stripes, spots, or other patterns. vibrissae. First setae in the oral row. wing. The wing is a membranous structure connected to the mesothoracic segment. A number of veins subdivide the wing into cells. The marginal vein, or costa (C; Figures 2.1 and 2.14) has two breaks: the proximal one, close to the humeral cross vein (h; Figure 2.1) is referred to as the humeral break (hum brk; Figure 2.14); the distal one, before the apex of the first vein, is called the subcostal break (sc brk; Figure 2.14). The remaining long veins are: R1, R2
3, R4
5, M1, CuA1, A1). Crossveins include r-m, dm-cu, and bm-dm, among others. See Figures 2.1 and 2.14 for more detail. Several wing vein indices have been proposed and are widely used to differentiate among species. Refer to Figures 2.1 and 2.14 for locations of the wing veins. The costal index is the length of the second costal section divided by the third costal section. The fourth vein index is the length of the distal section (fourth) of M1 divided by the third section of M1. The 4c index is the length of the third section of C divided by the third section of M1. The 5x index is the length of the distal (third) section of CuA1 divided by the length of cross vein dm-cu.

References Duda, O. (1935). Drosophilidae. In: Der Fliegen der palaearktishen Region (E. Lindner, ed.), Vol. VI, pp. 1–118. E. Schweizerbart’sche Verlagsbuchhandlung, Stuttgart. Ferris, G. F. (1950). External morphology of the adult. In: Drosophila Guide: Introduction to the Genetics and Cytology of Drosophila melanogaster (M. Demerec and B. P. Kaufmann, eds), 5th edn, pp. 368–419. Carnegie Institution of Washington, Washington, DC. Frota-Pessoa, O. (1954). Revision of the tripunctata group of Drosophila with description of fifteen new species (Drosophilidae, Diptera). Archos Mus. Parana. 10, 253–304. Grimaldi, D. A. (1987). Phylogenetics and taxonomy of Zygothrica (Diptera: Drosophilidae). Bull. Am. Mus. Nat. Hist. 186, 103–268.

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How to look at flies Grimaldi, D. A. (1990). A phylogenetic, revised classification of genera in the Drosophilidae (Diptera). Bull. Am. Mus. Nat. Hist. 197, 1–139. Grimaldi, D. A. and Fenster, G. (1989). Evolution of extreme sexual dimorphisms: structural and behavioral convergence among broad-headed male Drosophilidae (Diptera). Am. Mus. Nov. 2939, 1–25. Hardy, D. E., Kaneshiro, K. Y., Val, F. C. and O’Grady, P. M. (2001). Review of the haleakalae species group of Hawaiian Drosophila (Diptera: Drosophilidae). Bishop Museum Bull. Entomol. 9, 1–88. Heed, W. B. and Krishnamurthy, N. B. (1959). Genetic studies on the cardini group of Drosophila in the West Indies. Univ. Texas Publ. 5914, 155–179. Kambysellis, M. P. (1993). Ultrastructural diversity in the egg chorion of Hawaiian Drosophila and Scoptomyza: ecological and phylogenetic considerations. Intl J. Insect Morphol. Embryol. 22, 417–446. McAlpine, J. F. (1981). Morphology and terminology – adults. In: Manual of Nearctic Diptera (J. F. McAlpine, B. V. Peterson, G. E. Shewell et al., eds), Vol. 1, pp. 9–63. Agriculture Canada, Ottawa. O’Grady, P. M., Kam, M. W. Y., Val, F. C. and Pereirra, W. (2003a). Revision of the Drosophila mimica subgroup, with descriptions of ten new species. Ann. Entomol. Soc. Am. 96(1), 12–38. O’Grady, P. M., Bonacum, J., DeSalle, R. and Val, F. C. (2003b). The placement of the Engiscaptomyza, Grimshawomyia, and Titanochaeta, three clades of endemic Hawaiian Drosophilidae. Zootaxa 159, 1–16. Patterson, J. T. (1943). Drosophilidae of the Southwest. Univ. Texas Publ. 4313, 7–216. Patterson, J. T. and Mainland, G. B. (1944). Drosophiliae of Mexico. Univ. Texas Publ. 4445, 9–101. Sturtevant, A. H. (1916). Notes on North American Drosophilidae with descriptions of twentythree new species. Ann. Ent. Soc. Am. 9, 323–343. Sturtevant, A. H. (1921). The North American species of Drosophila. Publ. Carnegie Instn. 301, 1–150. Sturtevant, A. H. (1939). On the subdivision of the genus Drosophila. Proc. Natl. Acad. Sci. USA 25, 137–141. Sturtevant, A. H. (1942). The classification of the genus Drosophila with the description of nine new species. Univ. Texas Publ. 4213, 5–51. Throckmorton, L. H. (1966). The relationships of the endemic Hawaiian Drosophilidae. Univ. Texas Publ. 6615, 335–396. Wheeler, M. R. (1952). The Drosophilidae of the Nearctic region exclusive of the genus Drosophila. Univ. Texas Publ. 5204, 162–218. Wheeler, M. R. (1982). Drosophilidae: a taxonomic overview. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3b, pp. 1–105. Academic Press, London.

CHAPTER 3

Key to species

The following key identifies a number of species maintained by the Drosophila Species Stock Center, as well as a number of taxa that are not currently in culture. Several caveats should be stressed here. First, it is possible that, when using this key for wild collected Drosophilidae, species not included here will either not key properly or will key to another species. Identifications should always be checked against the original description of the species in question to verify your determination. Wheeler (1981, 1986) provides a somewhat outdated printed catalog of species descriptions prior to 1986. Taxodros (Baechli, 2005) is a more up-to-date online catalog that should prove extremely useful. Any characters that do not seem to fit original descriptions should be examined in further detail. Sending questionable species to experts for identification and/or examining type material may be necessary. Second, while we have made every effort to minimize the need for dissections in this key, in many cases dissection may be required to positively identify species. Grimaldi (1987) provides some discussion of appropriate dissection techniques for Drosophilidae. The following publications were used as references to assemble the couplets: Patterson, 1943; Patterson and Mainland, 1944; Burla et al., 1949; Magalhaes, 1962; Hardy, 1965; Sulerud and Miller, 1966; Strickberger, 1968; Bock, 1971, 1976; Bock and Wheeler, 1972; Wheeler, 1981, 1986; Ashburner, 1989; O’Grady, 1999; Heed and O’Grady, 2000; O’Grady and Kidwell, 2002. Refer to catalogs (Wheeler, 1981, 1986; Baechli, 2005) for original descriptions. We would also like to thank the following individuals for their assistance with these keys: Wyatt Anderson (obscura group), Sergio Castrezana (photographs, cactophilic species, general advice), Tamar Erez (nannoptera group), W. B. Heed (repleta group and lengthy discussion of Drosophila biology, morphology, and taxonomy), and Bryant McAllister (virilis group). Participants and instructors in the annual Drosophila Workshops held in Tucson from 2001 to 2003 also provided excellent feedback on various portions of these keys.

Key to species in the Drosophila Species Stock Center 1.

– Acrostichal setulae in two to four rows (Figure 3.1) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 – Acrostichal setulae in six or more rows (Figure 3.2) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7

2.

– Slender flies; habitus unicolorous or with stripes, never with elaborate patterns of pigmentation; wings typically hyaline, although sometimes with apical spot and/or infuscations at crossveins; arista with only one to two ventral rays in addition to terminal fork . . . . . . . . . . . . . . . . . . . genus Scaptomyza Hardy, 3 – Habitus with distinct pattern of spots and stripes, similar to that of wings; acrostichals in two rows; three sets of dorsocentrals present (Figure 3.3) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . genus Samoaia, 6

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Figure 3.1. Mesonotum of Scaptomyza adusta showing four rows of acrostichal setulae between anterior dorsocentral setulae.

Figure 3.2. Mesonotum of Drosophila immigrans with eight rows of acrostichal setulae between anterior dorsocentrals.

Scaptomyza 3.

– Paralobes absent (Figure 3.4); dark species, mesonotum and abdomen polished unicolorous dark brown to black; arista with two ventral rays . . . . . . . . . . . . . . . . . . . . . . . . . . . . subgenus Bunostoma, 4

Key to species

87

Figure 3.3. Patterned mesonotum of Samoaia leonensis.

Figure 3.4. Paralobes absent, posteroventral projections on ninth tergite in Scaptomyza anomala. From Hardy (1965), University of Hawaii Press.

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Figure 3.5. Paralobes present in Parascaptomyza. From Hardy (1965), University of Hawaii Press.

– Paralobes present (Figure 3.5); lighter species, brown stripes present on mesonotum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . subgenus Parascaptomyza, 5

subgenus Bunostoma 4.

– Acrostichal setulae in two rows; legs entirely yellow; pleura entirely dark brown to black; single humeral seta present; ventral margin of ninth tergite with prominent posteroventral projections (Figure 3.4) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . S. anomala Hardy – Acrostichal setulae in four rows; legs yellow, tinged with brown on apices of tarsi; pleura tinged faintly with yellow; ventral margins of ninth tergite nearly vertical (Figure 3.6) . . . . . . . . . . . S. palmae Hardy

subgenus Parascaptomyza 5.

– Secondary clasper on anal plate with four short, thick setae at tip; about nine thinner setae below; surstylus “C-shaped” with three distinct regions of prensisetae; one row of about six thin prensisetae on upper portion, thirteen prensisetae in middle portion, and six prensisetae on lower portion . . . . . . S. elmoi Takada – Secondary clasper on anal plate with two heavy, elongate setae at tip; surstylus not “C-shaped” but flatter, with only two regions of prensisetae; about eight prensisetae above and seven prensisetae setae below; a number of thinner, medially directed setae present on lower margin of surstylus . . . . . . . S. adusta Loew

Key to species

89

Figure 3.6. Ventral margins of ninth tergite in Scaptomyza palmae. From Hardy (1965), University of Hawaii Press.

Figure 3.7. Head setae of Chymomyza amoena depicting placement of proclinate setae (po) relative to the anterior reclinate orbital (aro).

Samoaia 6.

– Wings and thorax with distinct pattern of tan, gray and black; legs banded; spiracles six and seven always found at ventral or posteroventral margin of tergite six . . . . . Samoaia leonensis Wheeler & Kambysellis

7.

– Postocellar setae small and inconspicuous, less than one-third the length of the ocellars; proclinate orbital seta arises posterior to anterior reclinate; proclinate roughly equal in length to anterior reclinate (Figure 3.7);

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Figure 3.8. Foreleg of Chymomyza amoena.

Figure 3.9. Head setae of Drosophila immigrans depicting placement of the proclinate setae (po) relative to the anterior reclinate orbital (aro).

inner margin of forefemur always armed with a row of stout setae (Figure 3.8) . . . . . . . genus Chymomyza, 8 – Postocellar setae well developed; proclinate typically arises anterior to or even with the anterior reclinate orbital (in D. mimica the proclinate is posterior to the anterior reclinate but the other characters do not fit) (Figure 3.9) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9

Chymomyza 8.

– Legs entirely yellow; wings with a distinct pattern of three brown infuscations, one below apex of second vein, another across crossvein dm-cu, and a third at the apex of the wing . . . . . . . C. amoena Loew – Femora, tibia, and basitarsi dark; costal cell darkened; apex of wing whitish . . . . . . C. procnemis Williston

Hirtodrosophila 9.

– Third antennal segment covered with elongate setulae, roughly three times the length of ground setulae (Figure 3.10) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hirtodrosophila pictiventris Duda – Setulae on third antennal segment not elongate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10

Key to species

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Figure 3.10. Third antennal segment of Hirtodrosophila duncani showing elongate setulae.

Figure 3.11. Head and thoracic vittae in Zaprionus ghesquierei.

Liodrosophila 10.

– Anterior reclinate setae minute or absent; ground coloration of thorax and abdomen has a distinct metallic tinge . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Liodrosophila aerea Okada – Anterior reclinate present, not minute; ground color not metallic . . . . . . . . . . . . . . . . . . . . . . . . . . 11

11.

– Multicolored vittae present on the head and notum (Figures 3.11–3.13); head with three vittae, one medial and two along the fronto-orbital plate, which are bordered by dark brown or orange; notum light orange with one or more orange vittae present . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Zaprionus, 12 – Vittae on head and notum either unicolorous or not present . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16

92

Figure 3.12. Head and thoracic vittae in Zaprionus sepsoides.

Figure 3.13. Head and thoracic vittae in Zaprionus tuberculatus.

How to look at flies

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Figure 3.14. Prominent tubercule on forefemur of Zaprionus sepsoides.

Figure 3.15. Unadorned forefemur of Zaprionus ghesquierei.

Zaprionus 12.

– Forefemur with short stout knob, or tubercule, located near middle of the posteroventral surface (Figure 3.14) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 – Forefemur unadorned (Figure 3.15) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14

13.

– Ventral surface of knob with one straight and one hook shaped seta . . . . . . Z. tuberculatus Malloch

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Figure 3.16. Prescutellar acrostichal setulae (p acr s) in Scaptodrosophila latifasciaformis.

– Ventral surface of knob with two strong setae; dorsal surface with a single seta (Figure 3.14) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Z. sepsoides Duda 14.

– Wings strongly shining, brownish . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Z. badyi Burla – Wings hyaline, without shining brownish color . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15

15.

– Mesonotum and scutellum between the two middle stripes black; scutellum with a white apical spot (Figure 3.11); frons lacking white medial stripe . . . . . . . . . . . . . . . . . . . . . . . . . Z. ghesquièrei Collart – Scutellum lacking apical white spot, mesonotum and scutellum otherwise not as above; frons with a fine, white medial stripe anterior to ocelli . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Z. inermis Collart

16.

– Three subequal katepisternal setae present; prescutellar acrostichal setulae usually present (Figures 3.16, 3.17) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . genus Scaptodrosophila, 17 – Middle katepisternal seta either absent or much smaller than anterior and posterior katepisternals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Drosophila, 21

Scaptodrosophila 17.

– Lower portion of genital arch densely setose; anal plate with dense setae, especially at the apex; prensisetae on surstylus arranged in a concave row (Figure 3.18) . . . . . . . . . . . . . . . . . . victoria group, 18 – Lower portion of genital arch with few setae; anal plate not setose; prensisetae on surstylus arranged in a straight row . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . S. latifasciaformis Duda

18.

– Eight or more setae on hypandrium (Figure 3.18) . . . . . . . . . . . . . . . . . . . . S. lebanonensis Wheeler – No more than seven setae on hypandrium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19

Key to species

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Figure 3.17. Prescutellar acrostichal setulae (p acr s) in Scaptodrosophila lebanonensis.

Figure 3.18. Genital arch of Scaptodrosophila lebanonensis.

19.

– Palps dark brown to black . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . S. brooksae Pipkin – Palps light tan, sometimes darker at apex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20

20.

– Dark species; face and front dark brown; gena dark brown; ocellar triangle and orbits black . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . S. stonei Pipkin

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– Light species; face and front yellowish brown; cheeks yellow; ocellar triangle and orbits brown . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . S. pattersoni Pipkin

Drosophila subgenus Dorsilopha 21.

– Yellowish species with a longitudinally striped pleurae (Figure 3.19); preapical setae absent on second and third tibiae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. (Dorsilopha) busckii – Characters not as above, most notably the pleurae without lateral stripes . . . . . . . . . . . . . . . . . . . . 22

22.

– Second to fifth abdominal tergites with posterior dark bands not broken in mid-dorsal line (Figure 3.20) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . subgenus Sophophora, 23 – Abdominal tergites broken or interrupted at the mid-dorsal line, this characteristic is often difficult to see in dark species (Figure 3.21) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . subgenus Drosophila, 100

subgenus Sophophora 23.

– Dark species . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24 – Yellowish species . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25

24.

– Sex combs present on foretarsus of males (Figure 3.22); lacking opaque areas on tergites of female; preapical setae on first tibae usually long; second oral bristle small (less than half the length of first oral); middle orbital bristle large . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . obscura group, 26 – Sex combs absent on foretarsus of males, opaque areas present on tergites of female; other characters not as above . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . saltans group, 36

25.

– Sex combs present on the foretarsus of most, but not all, males (Figures 3.23, 3.24); sometimes sex comb is composed of only one or two setae; sterno index generally above 0.3 . . . . . melanogaster group, 44

Figure 3.19. Lateral view of Drosophila (Dorsilopha) busckii.

Key to species

97

Figure 3.20. Dorsal view of abdomen in Drosophila similans showing unbroken mid-dorsal line characteristic of the subgenus Sophophora.

Figure 3.21. Dorsal view of abdomen in Drosophila immigrans showing interruption of mid-dorsal line found in most subgenus Drosophila species.

98

How to look at flies

Figure 3.22. Sex combs of obscura group, Drosophila algonquin. From Sulerud and Miller (1966).

Figure 3.23. Sex combs of melanogaster group, Drosophila nikananu.

– Sex combs absent on foretarsus of males; sterno index about 0.3 (except in D. sucinea) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . willistoni group, 94 26.

27.

The obscura species group – Acrostichal setulae in six rows; sex comb on basitarsus only . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27 – Acrostichal setulae in eight rows; sex combs on first and second tarsal segments . . . . . . . . . . . . . 30 – Sex comb large, covering two-thirds of basitarsus, with about seven to ten teeth (Figure 3.25); mesonotum without markings; testes short and only slightly coiled (more than nine times as long as

Key to species

99

Figure 3.24. Sex combs of melanogaster group, Drosophila phaeopleura.

Figure 3.25. Sex comb of Drosophila tolteca. From Sulerud and Miller (1966).

Figure 3.26. Sex comb of Drosophila affinis. From Sulerud and Miller (1966).

wide, more than three gyres); sterno index about 0.7; costal index about 2.2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. tolteca Patterson & Mainland – Sex comb small, covering only the tip of the basitarsus, with four to six teeth . . . . . . . . . . . . . . . . 28 28.

– Large teeth on sex comb (Figure 3.26) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. affinis Sturtevant – Small teeth on sex comb . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29

29.

– Frons distinctly pollinose . . . . . . . . . . . . . . . . . . . . . . . . . D. narragansett Sturtevant & Dobzhansky – Frons dull, dark brown; mesonotum pollinose, sometimes marked with four dark brown longitudinal bands (two inside and two outside dorsocentral rows); front not pollinose, shining; six or fewer prensisetae on surstylus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. azteca Sturtevant & Dobzhansky

30.

– First tarsomere equal in length to second tarsal segment; sex combs long . . . . . . . . . . . . . . . . . . . 31 – Basitarsus distinctly longer than second tarsal segment; sex combs shorter . . . . . . . . . . . . . . . . . . 35

31.

– Pleura and basal two tergites yellowish . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. guanche Monclus – Pleura and abdomen entirely dark . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32

32.

– Proximal sex comb with greater than ten teeth; distal sex comb with nine to thirteen teeth . . . . . . 33 – Proximal and distal sex combs with fewer than ten teeth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34

100

How to look at flies

33.

– Wings slightly darkened along costal fringe; ventral margin of cercus with patch of short dense setulae; external process of epandrium rounded and bulging at base and drawn out into a thin projection; surstylus large and cup-shaped, laterally compressed and containing a very short and square looking comb with six to eight prensisetae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. subobscura Collin – Wings hyaline along costal margin; ventral margin of cercus with long setulae . . . D. ambigua Pomini

34.

– Proximal sex comb with six to ten teeth; distal sex comb with five to eight teeth; cheeks wide, gena about one-third width of the eye; dark species with indistinct grayish stripes on the mesonotum; relatively large species . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. miranda Dobzhansky – Proximal sex comb with six to eight teeth; distal sex comb with five to six teeth; surstylus with six or seven prensisetae; gena narrow, about one-fourth width of the eye; species lighter, with more distinct grayish stripes on mesonotum; relatively smaller than above . . . . . . . . . . . . D. pseudoobscura Frolova (NOTE: D. persimilis Dobzhansky and Epling also keys to this couplet.)

35.

– Tip of aedeagus sharply cut; paraphyses slender . . . . . . . . . . . . . . . . . . . . . . . . D. bifasciata Pomini – Tip of aedeagus slender; paraphyses broadened . . . . . . . . . . . . . . . . . . . . . . . . . . . D. obscura Fallen

36.

The saltans species group – Dark species, mesonotum with striped or spotted pattern; some small hairs present below carina . . . 37 – Dark to yellowish species; mesonotum unicolorous, without clear pattern; subcarinal hairs may be present or absent . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41

37.

– Mesonotal pattern includes, among the stripes or spots, one unpaired dark brown to black median streak or spot in front of the anterior dorsocentrals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 38 – Pattern on mesonotum lacks unpaired dark brown to black median spot or streak; sixth tergite of female with one opaque area in each side, near the posterior margin (Figure 3.27) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. sturtevanti Duda

38.

– Spermatheca without invagination (Figure 3.28a); penis not cylindric, without lateral flaps . . . . . 39 – Spermatheca with invagination (Figure 3.28b); penis cylindric, with lateral flaps . . . . . . . . . . . . . 40

39.

– Spermatheca without collar at base; seventh sternite of females pointed at apices, with sharply cut V-shape medially; male genitalia as in Figure 3.29 . . . . . . . . . . . . . . . . . . . . . . . . D. saltans Sturtevant – Spermathecae with collar at base; seventh sternite of females pointed at apices but with a more U-shaped median; male genitalia as in Figure 3.30 . . . . . . . . . . . . . . . . . . . . . D. lusaltans Magalhaes

40.

– Spermathecae with long invagination; sixth sternite of females rounded at apices, median region broad; penis as in Figure 3.31 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. prosaltans Duda

Figure 3.27. Sixth tergite of Drosophila sturtevanti female. From Magalhaes (1962).

Key to species

101

– Spermathecae with thin invagination; sixth sternite of females somewhat pointed at apices, with thin, deeply incised V-shape; penis as in Figure 3.32 . . . . . . . . . . . . . . . . . . . . . . . D. austrosaltans Spassky 41.

– Yellowish species; abdomen dark yellow . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42 – Dark species; abdomen dark brown to black; pleurae lacks distinct stripes; sixth tergite of female without a median opaque area . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. neocordata Magalhaes

Figure 3.28. Spermatheacae of (a) D. lusaltans and (b) D. nigrosaltans, showing presence and absence of invagination. From Magalhaes (1962).

Figure 3.29. Male genitalia of D. saltans.

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Figure 3.30. Aedeagus of D. lusaltans. From Magalhaes (1962).

Figure 3.31. Male genitalia of D. prosaltans.

42.

– Pleurae with distinct brownish stripes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. milleri Magalhaes – Pleurae not as above . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43

43.

– Abdomen yellow with indistinct brown basal bands; aedeagus as in Figure 3.33 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. emarginata Sturtevant

Key to species

103

Figure 3.32. Male genitalia of D. austrosaltans.

Figure 3.33. Male genitalia of D. emarginata.

(NOTE: a darker form of D. emarginata also exists and will key to D. neocordata except in the characters of male genitalia.) – Abdomen with dark brown band on fifth tergite that reaches the lateral margin of tergite . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. subsaltans Magalhaes

104

How to look at flies

Figure 3.34. Sex comb of Drosophila melanogaster. From Bock and Wheeler (1971).

Figure 3.35. Sex comb of Drosophila takahashii. From Bock and Wheeler (1971).

44.

45.

The melanogaster species group – Sex combs absent . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. lucipennis Lin (suzukii subgroup, in part) – Sex combs present on one or more tarsal segments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45 – Sex combs present on basitarsus only (Figure 3.34) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 46 – Sex combs present on more than the basitarsal segment (Figure 3.35) . . . . . . . . . . . . . . . . . . . . . . 57

Key to species

105

Figure 3.36. Sex comb of Drosophila denticulata. From Bock and Wheeler (1971).

Figure 3.37. Sex comb of Drosophila eugracilis. From Bock and Wheeler (1971).

46.

– Sex comb composed of two to three stout, claw-like setae on the apex of the basitarsus (Figure 3.36) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. denticulata Bock & Wheeler (denticulata subgroup) – Sex combs not as above . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47

47.

– Hooked setae present on male mid-legs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48 – Hooked setae absent on mid-leg of male . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49

48.

– Sex comb with only two strong setae (Figure 3.37) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. eugracilis Bock & Wheeler (eugracilis subgroup)

106

How to look at flies

Figure 3.38. Sex comb of Drosophila biarmipes. From Bock and Wheeler (1971).

Figure 3.39. Primary and secondary claspers in Drosophila nikananu. From Bock and Wheeler (1971).

– Sex combs in two rows, each with about three setae (Figure 3.38) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . biarmipes Malloch (suzukii subgroup, in part) 49.

– Primary and secondary claspers present (Figure 3.39); sex comb composed of a single row of eight or nine stout setae inserted on distal section of basitarsus (Figure 3.40) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. nikananu Burla (montium subgroup, in part) – Primary claspers only present (Figure 3.41) . . . . . . . . . . . . . . . . . . . . . . melanogaster subgroup, 50

Key to species

107

Figure 3.40. Sex comb of Drosophila nikananu. From Bock and Wheeler (1971).

Figure 3.41. Primary claspers present in the melanogaster subgroup, Drosophila melanogaster. From Bock and Wheeler (1971).

50.

– Aedeagus with lateral expansions; epandrium with posterior expansion; anal plates lacking ventral processes and teeth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51 – Aedeagus lacks lateral expansion; epandrium without posterior expansion (except in D. teissieri where it is triangular); anal plates with ventral process or strong teeth . . . . . . . . . . . . . . . . . . . . . . . 54

108

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51.

– Epandrial expansion longer than wide . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52 – Epandrial expansion wider than long . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53

52.

– Epandrial expansion narrow, fingerlike . . . . . . . . . . . . . . . . . . . . . . . . D. mauritiana Tsacas & David – Epandrial expansion elongate, sinous, and slightly enlarged at apex and pointed ventrally . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. sechellia Tsacas & Baechli

53.

– Expansion on epandrium trapeziodal in shape . . . . . . . . . . . . . . . . . . . . . . . D. melanogaster Meigen – Expansion on epandrium large, semicircular, clearly viewed laterally . . . . . . D. simulans Sturtevant

54.

– Anal plates lack ventral process, but have strong teeth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55 – Anal plates lack teeth (D. yakuba does possess small spines) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 56

55.

– Epandrium with a small, triangular expansion inserted anterior to clasper; phallus pointed . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. teissieri Tsacas – Epandrium lacking expansion; phallus truncated, boomerang-shaped, with small spines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. orena Tsacas & David

56.

– Anal plates knee-shaped in lateral view; with prolonged, flat, ventral process; phallus slender, curved dorsally . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. yakuba Burla – Anal plates without ventral process; large, straight, gutter-like phallus with saw-shaped fringes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. erecta Tsacas & Lachaise

57.

– Sex combs present on first to third tarsomeres (Figure 3.42) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58 – Sex combs present on first and second tarsomeres (Figure 3.35) . . . . . . . . . . . . . . . . . . . . . . . . . . 62

Figure 3.42. Sex comb of Drosophila elegans. From Bock and Wheeler (1971).

Key to species

109

58.

– Primary claspers only present . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 – Primary and secondary claspers present (Figure 3.43) . . . . . . . . . . ananassae subgroup (in part), 60

59.

– Sex combs with two transverse rows (one or two basal, two or three distal teeth) on basitarsus, two or three rows (zero to one basal; one or two middle, one or two distal teeth) on second tarsal segment, and two teeth on third tarsomere, one above the other; one very large ventromedial tooth present on anal plate (Figure 3.42) . . . . . . . . . . . . . . . . . . . . . . . . . . . D. elegans Bock & Wheeler (elegans subgroup) – Sex comb with three transverse rows (one to three basal, two to four middle, four or five distal teeth) on basitarsus, three or four rows (zero to one, three to four, three to five, three to four) on second tarsal segment, and two rows on third tarsal segment (one basal, one or two distal); anal plate with numerous setae, single large tooth not present (Figure 3.44) . . . . . . . . D. varians (ananassae subgroup, in part)

60.

– Single seta present on third tarsomere . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61 – Sex comb (Figure 3.45) on first and second tarsal segments in three or four rows (each with three to five teeth), sex comb on third tarsal segment with two rows of setae (two teeth each) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. phaeopleura Bock & Wheeler

61.

– First tarsal segment with five rows of setae (two to eight rows of teeth each) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. ananassae Doleschall – Sex comb on first and second tarsal segments (Figure 3.46) with three rows of setae (one to five teeth each) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. pallidosa Bock & Wheeler

62.

– Primary claspers only present . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63 – Primary and secondary claspers present . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 71

Figure 3.43. Primary and secondary claspers in the ananassae subgroup, Drosophila phaeopleura. From Bock and Wheeler (1971).

110

How to look at flies

Figure 3.44. Sex comb of Drosophila varians. From Bock and Wheeler (1971).

Figure 3.45. Sex comb of Drosophila phaeopleura. From Bock and Wheeler (1971).

63.

– Hooked setae on male mid-tibia and basitarsus absent . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. mimetica Bock & Wheeler (suzukii subgroup, in part) – Hooked setae present on mid-legs of males . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 64

64.

– Sex combs longitudinal, running along entire length of basitarsus and second tarsomere (Figure 3.47) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. ficusphila Kikkawa & Peng (ficusphila subgroup) – Sex combs transverse (Figure 3.35) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 65

Key to species

111

Figure 3.46. Sex comb of Drosophila pallidosa. From Bock and Wheeler (1971).

Figure 3.47. Sex comb of Drosophila ficusphila. From Bock and Wheeler (1971).

65.

– Primary surstylus with a ventrolateral comb of long rounded black setae, only a few teeth dorsolaterally (Figure 3.48) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . takahashii subgroup, 66 – Primary surstylus large, with several sets of distinctly different teeth (Figure 3.49) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. pulcherella (suzukii subgroup, in part)

112

How to look at flies

Figure 3.48. Male genitalia in the takahashii subgroup, Drosophila lutea. From Bock and Wheeler (1971).

Figure 3.49. Male genitalia of Drosophila pulcherella. From Bock and Wheeler (1971).

66.

– Wings of male infuscated (Figure 3.50) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67 – Wings hyaline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 68

67.

– Sex comb (Figure 3.51) with single row on basitarsus (four teeth) and two rows on second tarsomere (one tooth basally, three distally); basal branch of posterior paramere pointed apically and not serrate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. trilutea Bock & Wheeler

Key to species

113

Figure 3.50. Wing of Drosophila prostipennis. From Bock and Wheeler (1971).

Figure 3.51. Sex comb of Drosophila trilutea. From Bock and Wheeler (1971).

– Sex comb (Figure 3.52) with two rows (three teeth each) on basitarsus and two or three indistinct rows on second tarsomere (one, one, two); basal branch of posterior paramere long and serrate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. prostipennis, Lin 68.

– Basal branches of posterior paramere very short . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69 – Basal branches of posterior paramere long and serrated . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70

69.

– Sex comb (Figure 3.53) with two rows on basitarsus (two long teeth basally and five long teeth distally) and two rows on second tarsomere (one tooth basally and four teeth distally) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. takahashii Sturtevant and D. pseudotakahashii Mather Drosophila pseudotakahashii was described on the basis of crosses between strains of D. takahashii that only produced hybrids when the female was from Australia. The Australian form became

114

How to look at flies

Figure 3.52. Sex comb of Drosophila prostipennis. From Bock and Wheeler (1971).

Figure 3.53. Sex comb of Drosophila takahashii. From Bock and Wheeler (1971).

D. pseudotakahashii. The male genitalia must be dissected to differentiate these two species. Mather (1957) reports that the posterior paramere is vestigial and the basal process of the novosternum is long and serrated in D. pseudotakahashii. The latter character is short and conical in D. takahashii. 70.

– Sex comb (Figure 3.54) with two rows on basitarsus (three teeth each) and three rows on second tarsomere (one basal, one middle, and two distal) . . . . . . . . . . . . . . . . . . . D. paralutea Bock & Wheeler

Key to species

115

Figure 3.54. Sex comb of Drosophila paralutea. From Bock and Wheeler (1971).

Figure 3.55. Sex comb of Drosophila malerkotliana. From Bock and Wheeler (1971).

71.

– Sex combs transverse (Figures 3.55, 3.56) . . . . . . . . . . . . . . . . . . . ananassae subgroup (in part), 72 – Sex combs longitudinal (Figure 3.57) . . . . . . . . . . . . . . . . . . . . . . . . montium subgroup (in part), 74

72.

– Sex comb (Figure 3.55) with single row on basitarsus (three teeth) and two rows on second tarsomere (one basal and two distal teeth) . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. malerkotliana Parshad & Paika

116

How to look at flies

Figure 3.56. Sex comb of Drosophila bipectinata. From Bock and Wheeler (1971).

Figure 3.57. Male genitalia of Drosophila kanapiae. From Bock and Wheeler (1971).

Another species, D. pseudoananassae Bock, will also run to this couplet. The subspecies of D. malerkotliana and D. pseudoananassae are very closely related. Refer to Bock (1971) to key these species further. – Sex comb with more than one row of setae on basitarsus; other characters not as above . . . . . . . . 73

Key to species

117

Figure 3.58. Male genitalia of Drosophila baimaii. From Bock and Wheeler (1971).

Figure 3.59. Male genitalia of Drosophila parvula. From Bock and Wheeler (1971).

73.

– Abdomen of male entirely pale brown; sex comb on first tarsomere in two rows (six basal, eight distal teeth); single seta present on second tarsomere (Figure 3.56) . . . . . . . . . . . . . . D. bipectinata Duda – Abdomen of male black posteriorly; sex combs as above . . . . . . . . . . . . . . D. parabipectinata Bock

74.

– Secondary clasper not fused to anal plate (Figures 3.57–3.59) . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75 – Secondary clasper either partially or completely fused to anal plate . . . . . . . . . . . . . . . . . . . . . . . . 77

118

How to look at flies

Figure 3.60. Male genitalia of Drosophila lini. From Bock and Wheeler (1971).

75.

– Large black setae present on anal plate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76 – Anal plate lacks thick, elongate black setae; secondary clasper elongate and ovoid, with one large, curved medial seta (Figure 3.57) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. kanapiae Bock & Wheeler

76.

– Single dark seta present on anal plate; secondary clasper nearly circular, with a cluster of seven to fifteen setae (Figure 3.58) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. baimaii Bock & Wheeler – Two prominent setae present on anal plate; secondary clasper oval, with a single large black seta (Figure 3.59) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. parvula Bock & Wheeler

77.

– Secondary clasper partially fused to anal plate, either via a membranous connection or a narrow constriction (Figures 3.60, 3.61) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 78 – Secondary clasper completely fused to anal plate (Figures 3.62–3.64) . . . . . . . . . . . . . . . . . . . . . . 79

78.

– Secondary clasper connected to anal plate via a thin, membranous connection; two large setae present on secondary clasper, roughly equal in size (Figure 3.60) . . . . . . . . . . . . . D. lini Bock & Wheeler – Secondary clasper partially connected, but without membranous connection; two to three large setae present on secondary clasper, these setae are larger dorsally (Figure 3.61) . . . . . . . D. vulcana Graber

79.

– Secondary clasper possesses prominent lateral setae, each roughly equal in size to the typical medial setae on this structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 80 – Secondary clasper lacks large lateral setae, medial setae only present . . . . . . . . . . . . . . . . . . . . . . 84

80.

– Four medial and two lateral setae present on secondary clasper (Figure 3.62) . . . . D. rufa Kikkawa & Peng – Fewer than four medial setae present on secondary clasper; number of lateral setae varies . . . . . . 81

Key to species

119

Figure 3.61. Male genitalia of Drosophila vulcana. From Bock and Wheeler (1971).

Figure 3.62. Male genitalia of Drosophila rufa. From Bock and Wheeler (1971).

81.

– Three medial setae present on secondary clasper . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82 – Two medial setae present on secondary clasper . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83

82.

– Two lateral setae present on secondary clasper (Figure 3.63) . . . . . . . . . . . . . . D. lacteicornis Okada – Three lateral setae present on secondary clasper (Figure 3.64) . . . . D. punjabiensis Parshad & Paika

120

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Figure 3.63. Male genitalia of Drosophila lacteicornis. From Bock and Wheeler (1971).

Figure 3.64. Male genitalia of Drosophila punjabiensis. From Bock and Wheeler (1971).

83.

– Secondary clasper nearly oval in shape; larger medial seta on secondary clasper inserted dorsally (Figure 3.65) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. jambulina Parshad & Paika – Secondary clasper large, with ventral projection, not oval in shape; medial setae roughly equal in size (Figure 3.66) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. orosa Bock & Wheeler

84.

– Five medial setae on secondary clasper (Figure 3.67) . . . . . . . . . . . . . . . D. tsacasi Bock & Wheeler – Four or fewer medial setae on secondary clasper . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85

Key to species

121

Figure 3.65. Male genitalia of Drosophila jambulina. From Bock and Wheeler (1971).

Figure 3.66. Male genitalia of Drosophila orosa. From Bock and Wheeler (1971).

85.

– Only a single medial seta present on secondary clasper (Figure 3.68) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. mayri Mather & Dobzhansky – More than one medial setae on secondary clasper . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86

86.

– Basitarsal sex comb with twenty or fewer teeth; sex comb on second tarsomere with seventeen or fewer teeth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87

122

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Figure 3.67. Male genitalia of Drosophila tsacasi. From Bock and Wheeler (1971).

Figure 3.68. Male genitalia of Drosophila mayri. From Bock and Wheeler (1971).

– Basitarsal sex comb with more than twenty teeth; sex comb on second tarsomere with more than seventeen teeth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89 87.

– Sex comb on basitarsus with twenty teeth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 88 – Sex comb on basitarsus with only eighteen teeth; comb on second tarsomere with thirteen teeth; always only two medial setae on secondary clasper (Figure 3.69) . . . . . D. barbarae Bock & Wheeler

Key to species

123

Figure 3.69. Male genitalia of Drosophila barbarae. From Bock and Wheeler (1971).

Figure 3.70. Male genitalia of Drosophila seguyi. From Bock and Wheeler (1971).

88.

– Sex comb on second tarsal segment with seventeen teeth; either two or three medial setae present on secondary clasper (Figure 3.70) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. seguyi Smart – Sex comb on second tarsomere with only fourteen teeth; three medial setae always present on secondary clasper (Figure 3.71) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. greeni Bock & Wheeler

124

How to look at flies

Figure 3.71. Male genitalia of Drosophila greeni. From Bock and Wheeler (1971).

Figure 3.72. Male genitalia of Drosophila triauraria. From Bock and Wheeler (1971).

89.

– Sex comb on basitarsus with thirty or more teeth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 90 – Sex comb on basitarsus with fewer than thirty teeth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91

90.

– Sex comb on first tarsal segment with thirty-one teeth; sex comb on second tarsal segment with nineteen teeth; two medial setae on secondary clasper (Figure 3.72) . . . . . . . . D. triauraria Bock & Wheeler – Sex comb on basitarsus with thirty teeth; sex comb on second tarsomere with twenty-one teeth; three medial setae on secondary clasper (Figure 3.73) . . . . . . . . . . . . . . . . . . . . D. pennae Bock & Wheeler

Key to species

125

Figure 3.73. Male genitalia of Drosophila pennae. From Bock and Wheeler (1971).

Figure 3.74. Male genitalia of Drosophila serrata. From Bock and Wheeler (1971).

91.

– Two medial setae on secondary clasper (Figure 3.74) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 92 – Three medial setae on secondary clasper (Figure 3.75) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93

92.

– Abdominal tergites brown with dark posterior bands in both sexes; posterior margins of tergites with sparse rows of large setae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. serrata Malloch

126

How to look at flies

Figure 3.75. Male genitalia of Drosophila birchii. From Bock and Wheeler (1971).

– Abdominal tergites yellow to rufous, with dark posterior bands in both sexes; sparse rows of large setae absent from posterior margins of abdominal tergites . . . . . . . . . . . . . . . . . . . . D. kikkawai Burla 93.

94.

– Two medial setae on secondary clasper large, subequal in size, third seta smaller . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. bicornuta Bock & Wheeler – All three medial setae on secondary clasper large, roughly equal in size Several species (D. auraria Peng, D. biauraria Bock & Wheeler, D. birchii Dobzhansky & Mather, D. quadraria Bock & Wheeler) key to this couplet and genitalia dissections will have to be done to separate them fully. The original descriptions should be consulted in the case of these taxa. The willistoni species group – Wings with distinct infuscations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 – Wings hyaline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 96

95.

– Carina broad; second oral 3/4 first; middle orbital 1/2 other two . . . . . . . D. nebulosa Sturtevant Not exactly fitting above . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. fumipennis Duda

96.

– Sterno index 0.7; second oral about two-thirds length of the first . . . . . . . D. sucinea Patterson & Mainland – Sterno index about 0.3; second oral not as above . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97

97.

– Maxillary palps inflated and broadly rounded at the apex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. tropicalis Burla & Cunha, in Burla et al. – Palps more slender and pointed at the tips . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 98

98.

– Ocelli dark . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 – Ocelli light . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. equinoxialis Dobzhansky

99.

– Large hook-shaped process on inner margin of the hypandrium; middle orbital setae inserted intermediate between anterior and posterior orbital setae . . . . . . . . . . . . . . . . . . . . . D. willistoni Sturtevant

Key to species

127

Figure 3.76. Dorsocentral setae in Drosophila polychaeta.

– Inner margin of the hypandrium lacking large hook-shaped process, with only a small tubercule; middle orbital inserted at the same level as the anterior orbital. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. paulistorum Dobzhansky & Pavan, in Burla et al.

subgenus Drosophila 100.

– Anal plate partially fused to genital arch; aedeagus laterally flattened, at least apically, and apparently bilobed at least partially; darker species (usually); tend to be found in sap fluxes, fruits, cactus, etc., rarely reared from fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101 – Lighter species (usually); almost exclusively mycophagous; other characters not as above . . . . 155

The virilis-repleta radiation 101.

102.

– Three pairs of dorsocentral setae; acrostichal setulae in eight irregular rows (Figure 3.76) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . polychaeta group, 102 – Two pairs of dorsocentral setae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 The polychaeta species group – Pale yellow species, surstylus with a row of five or six teeth . . . D. polychaeta Patterson & Wheeler – Blackish brown species surstylus with a row of four teeth . . . . . . . . . . . . . . . D. asper Lin & Tseng

103.

– Each seta or setulae on mesonotum not arising from a single dark spot . . . . . . . . . . . . . . . . . . . 104 – Each seta or setulae on mesonotum arising from a single dark spot; spots sometimes fused into stripes or other patterns (Figure 3.77) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 127

104.

– Prescutellar setulae present (Figure 3.78) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 – Prescutellar setulae absent . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109

105.

The bromeliae species group – Thorax dull yellow-brown in color; single oral setae present . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . bromeliae group, Drosophila bromeliae Sturtevant – Dull black species; anterior scutellar setae convergent; testes orange . . . . . nannoptera group, 106

128

How to look at flies

Figure 3.77. Mesonotum of Drosophila aracataca showing each setae arising from a single black spot. This character is seen in several species groups, including the repleta and annulimana.

Figure 3.78. Thorax of Drosophila bromeliae showing prescutellar setulae.

106.

The nannoptera species group – Dark species, mostly black on dorsum; venter dark orange and black; sternites with heavy black pigment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 107 – Lighter species, light to medium brown dorsally; ventral surface yellow brown or orange brown; sternites lack pigment or are light yellow brown . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 108

Key to species

129

Figure 3.79. Male genitalia of Drosophila nannoptera, ventral view.

Figure 3.80. Male genitalia of Drosophila wassermani, ventral view.

107.

– Sternites four and five square and intact, not split; epandrial lobes symmetrical, short and narrowed to thin points (Figure 3.79) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. nannoptera Wheeler – Sternite five rectangular and split medially; epandrial lobes asymmetrical with left lobe longer than the right . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. pachea Patterson & Wheeler

108.

– Eyes red; epandrial lobes large and elaborate (Figure 3.80) . . . . . . D. wassermani Pitnick & Heed – Eyes dark, nearly black; epandrial lobes short and rounded with six or seven prensisetae (Figure 3.81); setulae on costal margin darker and thicker than typical; enlarged setae also present at regular intervals on the costa (Figure 3.82) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. acanthoptera Wheeler

130

How to look at flies

Figure 3.81. Male genitalia of Drosophila acanthoptera, ventral view.

Figure 3.82. Wing of Drosophila acanthoptera.

109.

– Acrostichal setulae in six rows . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 110 – Acrostichal setulae in eight or more rows . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 122

110.

– Anterior scutellar setae convergent (Figure 3.83); wings hyaline . . . . . . . . . . . melanica group, 111 – Anterior scutellar setae divergent; one or both crossveins infuscated; sterno index 0.8–0.9; third costal section with heavy setae on basal half or more . . . . . . . . . . . . . . . . . . . . . . . . virilis group, 115

111.

– Anterior scutellars divergent . . . . . . . . . . . . . . . . . . . . . . . . D. nigromelanica Patterson & Wheeler – Anterior scutellars convergent . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 112

112.

– Second oral setae about half the length of first . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113 – Second oral setae short, about one-third the length of first . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 114

113.

– Legs yellowish brown, hind tibia darker near base; two heavy setae on first tibia, one on second, preapicals on all three . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. paramelanica Patterson

The melanica species group

Key to species

131

Figure 3.83. Convergent anterior scutellar setae in Drosophila micromelanica.

– Legs brownish black, mid and hind femora and all tarsi paler; apical setae on first and second tibia, preapicals on all three . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. euronotus Patterson & Ward 114.

115.

– Costal index greater than 3.0 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. melanica Sturtevant – Costal index less than 3.0 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. micromelanica Patterson The virilis species group – Puparia gray . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. virilis Sturtevant – Puparia red to brownish . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116

116.

– Basal one-third to two-thirds of third costal section, with heavy seta . . . . . . . . . . . . . . . . . . . . . 117 – Third costal section with heavy setae on basal three-fourths . . . . . . . . . . . . . . . . D. americana, 121

117.

– Mesonotum with two prominent stripes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118 – Dark species, nearly black in ground color; stripes on mesonotum indistinct; abdomen dark brown to black, shining; second setae in oral row about half the length of first . . . . . . D. lacicola Patterson

118.

– Second oral setae not more than one-fourth length of first; heavy seta on third costal section scarcely extending beyond basal three-fifths . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119 – Second oral setae greater than two-thirds length of first; heavy seta on third costal section extending to basal two-thirds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120

119.

– Mesonotum and scutellum largely tan, pollinose; dark brown stripes present inside each dorsocentral row; pleurae significantly darker than mesonotum; legs pale yellowish tan . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. flavomontana Patterson – Thorax dark blackish brown, with a brownish black strip down the entire acrostichal region; legs brownish, distal tarsal joints black . . . . . . . . . . . . . . . . . . . . . . . . . . D. montana Patterson & Wheeler

120.

– Second oral three-fourths length of first; terminal tarsal segments slightly darker than the remainder of the leg; abdomen dark gray to black . . . . . . . . . . . . . . . . . . . . . . . . . . . D. novamexicana Patterson

132

How to look at flies

– Second oral two-thirds length of first; legs pale grayish yellow; abdomen grayish brown, lighter on basal segment in midline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. littoralis Meigen 121.

– Metaphase plate of male and female with three pairs rods, one pair Vs, and one pair dots . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. americana texana Patterson, Stone & Griffen – Metaphase plate of female: one pair rods, two pairs Vs, and one pair dots; metaphase plate of male: four rods, three Vs, and one pair dots . . . . . . . . . . . . . . . . . . . . . . D. americana americana Spencer

122.

– Anterior scutellar setae divergent . . . . . . . . . . . . . . . . . . . . . . . . . . . . mesophragmatica group, 123 – Anterior scutellar setae convergent . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .124

123.

124.

125.

The mesophragmatica species group – 5X index about 1.3 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. pavani Brncic – 5X index 1.0–1.1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. gaucha Jaeger & Salzano – Costal index greater than 3.0 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . robusta group, 125 – Costal index less than 3.0 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 126 The robusta species group – Only one prominent oral, second oral about one-fourth length of first; third costal section with heavy setulae on basal one-third . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. sordidula – Second oral setae two-thirds length of first; third costal section with heavy setulae on basal twofifths or more . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. lacertosa

The dreyfusi species group 126. – Thorax plain brown, lacking a distinct pattern; acrostichal setulae in eight to ten irregular rows . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . dreyfusi group, Drosophila camargoi The canalinea species group – Thorax black, patterned; acrostichal setulae in eight regular rows . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Drosophila canalinea Patterson & Mainland, 83 127.

128.

129.

– Apices of third and fourth longitudinal veins distinctly convergent (Figure 3.84); discal and second basal cells separated by a rudimentary vein; costal lappet present at subcostal break (Figure 3.85); proclinate inserted posterior to anterior reclinate . . . . . . . . . . . . . . . . . . . . . . . . . annulimana group, 128 – Apices or third and fourth longitudinal veins distinctly divergent; discal and second basal cells united; anterior reclinate inserted posterior to proclinate orbital . . . . . . . . . . . . . . repleta group, 130 The annulimana species group – Prescutellar acrostichal setae present . . . . . . . . . . . . . . . . . . . . . . . . . . . D. aracatacas Vilela & Val – Prescutellar acrostichal setae absent . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 129 – Pattern of pigment on mesonotum light, about 50 per cent of the ground color tan (Figure 3.86) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. gibberosa

Key to species

Figure 3.84. Convergent third and fourth longitudinal veins of D. aracatacas.

Figure 3.85. Costal lappet of D. aracatacas.

Figure 3.86. Pigmentation on mesonotum of D. gibberosa.

133

134

How to look at flies

Figure 3.87. Pigmentation on mesonotum of D. talamanca.

– Pattern of pigment on thorax dark, about 25 per cent of the ground color tan (Figure 3.87) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. talamanca

130.

The repleta species group – Large black species with red testes and a dark eye; mesonotum pale brown with dark brown spots that fuse on both sides of mid-dorsal line; ground color of mesonotum distinctly bluish gray; abdominal banding blackish in a complex pattern created by a dark marginal bar that is thickest in the posterior corner of tergites two, three and four, and usually connects with posterior band . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. buzzatii Patterson & Wheeler – Species may be large and dark, but not exactly as above; ground color of mesonotum without bluish hue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131

131.

– Males display “guarding behavior” and ride on backs of females, sometimes for several minutes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 132 – No guarding behavior observed . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 133

132.

– Testes yellow to burnt orange in color; mesonotum tan with darker brown spots; abdomen in males and females not elongated . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. mainlandi Patterson – Testes large, bright lemon yellow in color; mesonotum with dark gray spots; abdomen elongates in males and females . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. pegasa Wasserman

133.

– Frons glossy, with scattered setae that do not form a distinct V-shaped pattern; testes red; front broad; cheek narrow, less than one-fourth width of eye at widest point; eye round in shape . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. mettleri Heed – Frons not as above, typically with a distinct V-shaped pattern of setae; other characters not as above . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 134

Key to species

135

134.

– Coxae of forelegs darker than the remainder of the leg . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 – Coxae of forelegs about the same color as the remainder of the leg . . . . . . . . . . . . . . . . . . . . . . 137

135.

– Posterior crossveins clouded; lateral areas of tergites with light grayish areas near lateral margin, especially on first three tergites; costal index about 3.3, heavy setae on basal half of third costal section . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. nigricruria Patterson & Mainland, in Patterson (1943) – Posterior crossveins not clouded . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 136

136.

– Apical band on tergites not widened to form a triangle in the lateral area . . . . D. repleta Wollaston – Apical band on tergites greatly widened to cover nearly all of the lateral area . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. neorepleta Patterson & Wheeler

137.

– All lateral areas of abdominal segments almost completely covered by expansions of apical bands, solid color without interruptions or light areas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 138 – Some or all of lateral areas of abdominal segments from the more medial portion of the apical band near the angle of the tergite, lateral areas not completely covered with dark markings . . . . . . . . . 148

138.

– Arista with three branches above, excluding terminal fork . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139 – Arista with four branches above, excluding terminal fork . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 144

139.

– Crossveins without clouding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 140 – Crossveins clouded . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 152

140.

– Mesonotum brownish, indistinctly spotted; second oral about the same length as first; sterno index about 0.88; costal index about 2.8 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. ritae Patterson & Wheeler – Mesonotum grayish, distinctly spotted; second oral either absent or about half length of the first . . . . 141

141.

– Eyes dark red or wine in color . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .142 – Eyes bright red . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155

142.

– Only a single prominent oral seta present; long recurved hairs on medial side of male foretarsi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. nigrohydei Patterson & Wheeler – Second oral half length of the first; long recurved hairs present or not . . . . . . . . . . . . . . . . . . . . 143

143.

– Costal index about 3.0; second oral about half length of first . . . . D. nigrospiracula Patterson & Wheeler – Costal index about 3.8; second oral about one-third length of the first . . . . . . . . D. hydei Sturtevant

144.

– Crossveins clouded, apex of wing with some light clouding . . . . . . D. leonis Patterson & Wheeler – Wings entirely hyaline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 145

145.

– Lateral areas of abdominal tergites much darker than medial portions of apical bands; costal index 3.6 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. bifurca Patterson & Wheeler – Lateral areas of abdominal tergites not darker than medial portions of apical bands; costal index 3.0 or less . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 146

136

146.

How to look at flies

– Only one prominent oral seta present . . . . . . . . . . . . . . . . . . . D. fulvimacula Patterson & Mainland – Second oral at least half length of first; mesonotum grayish brown with longitudinal lines . . . . 147

147.

– Long slender species; width of cheek about three-eighths greatest diameter of eye; mesonotum brown with three dark brown longitudinal lines . . . . . . . . . . . . . . D. racemova Patterson & Mainland – Normally proportioned species; width of cheek about one-fourth greatest diameter of eye; mesonotum grayish brown without longitudinal lines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 150

148.

– Arista with four branches above, excluding the terminal fork . . . . . . . . . . . . . . . . . . . . . . . . . . . 149 – Arista with three branches above, excluding the terminal fork . . . . . . . . . . . . . . . . . . . . . . . . . . 151

149.

– Crossveins hyaline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 150 – Crossveins infuscated . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 151

150.

– Mesonotum dark grayish brown; sterno index about 0.73; apical bands on abdomen dark and distinct . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. spenceri Patterson – Mesonotum light brown; sterno index greater than 0.8; abdominal banding light and diffuse. . . . 155

151.

– Mesonotum grayish brown, distinctly spotted; eyes without iridescent highlights; tibia and femur without banding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. anceps Patterson & Mainland – Eyes with distinctly iridescent highlights; tibia and femur banded . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. meridiana Patterson & Wheeler

152.

– Crossveins clouded . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. longicornis Patterson & Wheeler – Crossveins without clouding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 153

153.

– Anterior crossvein black; apex of first costal segment jet black; costal index 3.6 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. hexastigma Patterson & Mainland – Anterior crossvein not darkened, same color as other wing veins; apex of first costal segment either darkened or not, however never jet black; costal index 3.1 or less . . . . . . . . . . . . . . . . . . . . . . . . . 154

154.

– Color pattern in lateral areas of abdominal segments faint, faded, and diffuse; mesonotum light brown . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155 – Color pattern in lateral areas of abdominal segments brown or black, distinct; mesonotum dark grayish-brown . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 157

155.

– Scutellum with very thick, U-shaped dark marking; abdominal tergites with a very faint diffuse apical band . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. mercatorum Patterson & Wheeler – Scutellum with an X-shaped dark marking; abdominal with distinct apical bands . . . . . . . . . . . 156

156.

– Apical bands of abdominal segments three-fourths or greater of the width of the tergite on either side of the mid-dorsal line; apex of first costal section not darkened . . . . . D. hamatofila Patterson & Wheeler – Apical bands of abdominal segments narrow, half or less of the width of the tergite on either side of the mid-dorsal line; apex of first costal section darkened . . . . . . . . . D. mojavensis Patterson & Crow

Key to species

137

157.

– Lateral areas of abdominal tergites not broken into distinct spots, more or less solid, but with distinctly lighter areas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 156 – Lateral areas of abdominal tergites broken into distinct spots . . . . . . . . . . . . . . . . . . . . . . . . . . . 158

158.

– Spots on posterolateral corner of lateral areas irregularly triangular in shape; dark markings are contiguous from irregularly triangle-shaped region in posterolateral corner of abdominal tergites to the margin of tergite . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. arizonae Ruiz, Heed & Wasserman – Spots on posterolateral corner of lateral areas distinctly triangular in shape; dark markings are only narrowly contiguous from triangle shaped region in posterolateral corner of abdominal tergites to margin of tergites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 159

159.

– Abdominal bands distinct; triangular areas in posterolateral corners distinctly darker than apical bands on medial portion of tergites, not united to medial portions of bands; eyes red; testes yellow . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. mulleri Sturtevant – Abdominal bands less distinct; triangular areas in posterolateral corners about the same shade or lighter than apical bands on median portions of tergites, these regions are joined by a narrow band; eyes vermilion; testes orange . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. aldrichi Patterson & Crow

The immigrans-tripunctata radiation 160.

The calloptera species group – Wing and mesonotum with a distinct pattern of dark infuscations (Figure 3.88); frons and face brilliant white (Figure 3.89) . . . . . . . . . . . . . . . . . . . . . . . . . calloptera group, D. ornatipennis Williston

Figure 3.88. Wing pigmentation patterns in D. ornatipennis, calloptera group.

Figure 3.89. Face and frons of D. ornatipennis.

138

How to look at flies

Figure 3.90. Enlarged acrostichal setulae in D. putrida.

– Wings may be infuscated or patterned, but not as above; face and frons not brilliant white in color . . . 161

161.

162.

163.

164.

165.

The testacea species group – Two enlarged acrostichal setulae present in the anterior region of mesonotum, near transverse suture (Figure 3.90) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . testacea group, D. putrida Sturtevant – Acrostichal setule not enlarged at suture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 162 – One or more thick dense spines on anal plate . . . . . . . . . . . . . . . . . . . . . . . . . . funebris group, 163 – Anal plate with cilia or setae, but not with thickened spines . . . . . . . . . . . . . . . . . . . . . . . . . . . . 164 The funebris species group – Anal plate with a single, large, upwardly curved spine (Figure 3.91) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. macrospina Stalker & Spencer, 1939 – Anal plate with a series of thickened spines (Figure 3.92) . . . . . . . . . . . . . . . . D. funebris Fabricius – Inner margin of femur with a row of stout, peg-like setae; this row is sometimes poorly developed (Figure 3.93) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . immigrans group, 165 – Inner margin of femur without a row of stout setae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 169 The immigrans species group – Row of peg-like setae on forefemur poorly developed . . . . . . . . . . . . . . . . . . hypocausta subgroup (Subgroup includes D. hypocausta Osten-Sacken; D. neohypocausta Lin & Wheeler, in Lin & Tseng; D. pararubida Mather; D. rubida Mather.) – Row of peg-like setae well developed . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 166

Key to species

139

Figure 3.91. Anal plate of D. macrospina.

Figure 3.92. Anal plate of D. funebris.

166.

– Ground color of mesonotum pale, with a series of prominent darker longitudinal stripes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . quadrilineata subgroup (sensu Wilson et al., 1969) – Not as above, if stripes are present then they are pale and indistinct . . . . . . . . . . . . . . . . . . . . . . 167

167.

– Foretarsi with some degree of modification . . . . . . . . . . . . . . . . . . . . . . . . . . . immigrans subgroup

140

How to look at flies

Figure 3.93. Inner margin of forefemur of D. immigrans.

(Subgroup includes D. formosana Duda; D. immigrans Sturtevant; D. signata Duda; D. tongpua Lin & Tseng.) – Foretarsi lacking modification, frons partially or entirely silvery-whitish . . . nasuta subgroup, 168 168.

– Entire frons of males silver to white . . . . D. albomicans Duda; D. kepulauana Wheeler, in Wilson et al.; D. kohkoa Wheeler, in Wilson et al.; D. nasuta Lamb (All species are very similar and require dissection of male genitalia to positively identify them – Wilson et al., 1969.) – Only fronto-orbital region silver or whitish in color . . . . . . . . D. pulaua Wheeler, in Wilson et al.; D. sulfurigaster albostrigata Wheeler, in Wilson et al.; D. sulfurigaster bilimbata Bezzi; D. sulfurigaster sulfurigaster Duda (Dissection of male genitalia is required to separate these taxa – Wilson et al., 1969.)

169.

– Costal index greater than 3.9 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 170 – Costal index less than 3.9 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172

170.

The pallidipennis species group – Abdomen with distinct pattern, shining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . cardini group, 171 – Abdomen tannish yellow and with some pattern; species not shining, more diffusely pigmented; costal index about 5.7; acrostichal setulae in eight rows . . . . . . . . pallidipennis group, D. pallidipennis Dobzhansky & Pavan

Key to species

171.

172.

141

The cardini species group – Anterior scutellar setae convergent . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. cardini Sturtevant – Anterior scutellar setae divergent . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. similis Williston – Heavy setulae on basal three-fifths or more of third costal section; sterno index 0.6 or greater . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . quinaria group, 173 – Heavy setulae on basal half or less of third costal section; sterno index 0.5 or less . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . tripunctata group, 184

173.

The quinaria species group – Wings distinctly spotted with thirteen black spots . . . . . . . . . . . . . . . . . . . . . . . D. guttifera Walker – Wings not as above . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 174

174.

– Mesonotum with three longitudinal stripes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 175 – Mesonotum without stripes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 176

175.

– Posterior crossvein slightly curvate; no infuscation at apex of wing . . . . . . . D. palustris Sturtevant – Posterior crossvein strongly curvate; heavy infuscations at the tips of each long vein at the wing apex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. subpalustris Spencer

176.

– Dark pigment on dorsal surface of abdomen forms distinct spots or dots (this character is somewhat variable but typically with about four spots on tergites one to four, and two spots thereafter) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 177 – Dorsal areas of tergites without distinct spots or dots . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 179

177.

– Both crossveins and apex of wing infuscated . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. quinaria Loew – Only crossveins infuscated . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 178

178.

– Wide clouds on crossveins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. recens Wheeler – Crossveins only narrowly clouded . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. falleni Wheeler

179.

– Crossveins not infuscated . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. innubila – One or both crossveins infuscated . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .180

180.

– Both crossveins clouded . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 181 – Only posterior crossveins clouded; dark abdominal banding thins out toward lateral margins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. munda Spencer

181.

– Dark abdominal bands reach lateral margins; narrow band of infuscation on crossvein . . . . . . . 182 – Dark abdominal bands do not reach lateral margins; wide clouds on crossveins . . . . . . . . . . . . 183

182.

– Some abdominal bands show lightened area on mid-dorsal line . . . D. suffusca Spencer, in Patterson – Abdominal banding not lightened at mid-dorsal line . . . . . . . . . D. tenebrosa Spencer, in Patterson

142

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How to look at flies

– Two anterior abdominal segments without dark banding . . . . . . . . . . . . D. magnaquinaria Wheeler – Dark banding appears at seconds abdominal segment . . . . . . . . . . . . . . . . . D. occidentalis Spencer

The tripunctata species group 184. – Acrostichal setulae in six rows; dark brown spot in the mid-dorsal line of tergites four, five, and six . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. tripunctata Loew – Acrostichal setulae in eight rows; dark brown spot present only on tergite six . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. unipunctata Patterson & Mainland, in Patterson

References Ashburner, M. (1989). Drosophila: A Laboratory Handbook and Manual (two volumes). Cold Spring Harbor Laboratory Press, New York. Baechli, G. (2005). Taxodros: The Database on Taxonomy of Drosophilidae, version 2005-1, last accessed 5 January 2005. http://taxodros.unizh.ch/ Bock, I. R. (1971). Taxonomy of the Drosophila bipectinata species complex. Univ. Texas Publ. 7103, 273–280. Bock, I. R. (1976). Drosophilidae of Australia. Aust. J. Zool. 40, 1–105. Bock, I. R. and Wheeler, M. R. (1972). The Drosophila melanogaster species group. Univ. Texas Publ. 7213, 1–102. Burla, H., da Cunha, A. B., Cordeiro, A. R. et al. (1949). The willistoni group of sibling species of Drosophila. Evolution 3, 300–314. Grimaldi, D. A. (1987). Phylogenetics and taxonomy of Zygothrica (Diptera: Drosophilidae). Bull. Am. Mus. Nat. Hist. 186, 103–268. Hardy, D. E. (1965). Diptera: Cyclorrhapha II, Series Schizophora, Section Acalypterae I, Family Drosophilidae. Insects of Hawai’i 12, 1–814. Heed, W. B. and O’Grady, P. M. (2000). Drosophila maya, a new Neotropical member of the Drosophila obscura species group. J. NY Entomol. Soc. 108(1–2), 97–104. Magalhaes, L. E. (1962). Notes on taxonomy, morphology and distribution of the saltans group of Drosophila, with descriptions of four new species. Univ. Texas Publ. 6205, 135–154. Mather, W. B. (1957). Genetic relations of four Drosophila species from Australia (Diptera: Drosophilidae). Univ. Texas Publ. 5721, 221–225. O’Grady, P. M. (1999). Reevaluation of phylogeny in the Drosophila obscura species group. Mol. Phylogen. Evol. 12(2), 124–139. O’Grady, P. M. and Kidwell, M. G. (2002). Phylogeny of the subgenus Sophophora (Diptera: Drosophilidae) based on combined analysis of nuclear and mitochondrial sequences. Mol. Phylogen. Evol. 22, 442–453. Patterson, J. T. (1943). The Drosophila of the Southwest. Univ. Texas Publ. 4313, 7–216. Patterson, J. T. and Mainland, G. B. (1944). The Drosophilidae of Mexico. Univ. Texas Publ. 4445, 9–101. Strickberger, M. W. (1968). Genetics. Macmillan Press, New York. Sulerud, R. L. and Miller, D. D. (1966). A study of key characteristics for distinguishing several Drosophila affinis subgroup species, with a description of a new related species. Am. Mid. Nat. 75, 446–474. Wheeler, M. R. (1981). The Drosophilidae: a taxonomic overview. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3a, pp. 1–97. Academic Press, London. Wheeler, M. R. (1986). Additions to the catalog of the world’s Drosophilidae. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3e, pp. 1–105. Academic Press, London.

CHAPTER 4

Collecting Drosophila in the wild Contents

• • • • • • •

Tools and equipment Preparing and using baits Collecting from natural substrates Collecting and transporting Hawaiian Drosophilidae Permits Transporting live adults or larvae References

Many questions are best addressed using wild caught flies or descendents of recently collected flies, as opposed to ordering cultures from a stock center. Assuming it is possible to travel to the sites where the species of interest are found, a number of practices will enhance the likelihood of successful collecting efforts. In this chapter we discuss tools and techniques for collecting and transporting wild Drosophila.

Tools and equipment Nothing is worse than arriving at a field site to find that some critical piece of equipment for collecting is missing. The chances of this happening are greatly reduced by making a list of items to be used at each step of the trip and packing accordingly. The fly list should be separate from any camping or travel equipment list, and should include the following items: food vials, tape, markers, cotton balls, forceps, cooler, filter paper, yeast, aspirators, instant medium, hand lens, spoon, knife, machete, net, Kleenex (not Kimwipes), anesthetizer, test-tube rack for vials, water, newspaper, fanny pack, and plastic bags. We recommend having available the two types of aspirators shown in Figure 4.1. Both can be constructed with components available in most laboratories. The first, shown in Figure 4.1a, is typical of the aspirator used in the lab to transfer flies without anesthesia. For species the size of D. melanogaster, the glass tubing can be 7 mm in diameter. Many species are larger, however, and require a larger diameter tubing – such as 8 or 9 mm – in order to avoid being damaged during handling. This type of aspirator can hold up to a dozen flies before they must be ejected into a vial. A disadvantage of this type of aspirator is that once flies are inside, a constant suction must be applied to prevent their escape before they eventually are blown gently into a holding

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How to collect wild flies Glass tube

Cheese cloth

Rubber tube

Flexible tube Glass or copper tub Mouth port

Two-hole rubber stopper

Vial

(a)

(b)

Figure 4.1. Two types of aspirators: (a) laboratory aspirator without a holding vial; (b) commercially available aspirator with holding vial.

Figure 4.2. Drosophila net with place to attach a vial in the end.

vial. The advantage of this aspirator is that the operator has control over distributing the collected flies – i.e. they can be ejected individually into vials if desired. The second type of aspirator (Figure 4.1b) deposits the collected flies directly into a holding vial, with or without culture medium, which can hold a larger number of flies.

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This type of aspirator is commercially available from Ward’s Scientific (http://www. wardsci.com) as the Suction Tube Aspirator, or from Bioquip (http://www.bioquip.com) as the 1135A Aspirator. The operator only needs to change the holding vial once it is full of flies. With this type of aspirator, flies will have to be sorted or redistributed later, either with the other type of aspirator or under anesthesia. Either type of aspirator can be used to collect flies from a bait or from a natural substrate. Some natural substrates may be noxious to the investigator, in which case a third model of aspirator, the Blowing Aspirator (Bioquip 1115A), is recommended, as the operator blows out to suck in the flies. A variety of nets have been described and discussed in detail by Carson and Heed (1983). Figure 4.2 shows a net typically used by Drosophila workers, because its design allows a standard 25
95 mm Drosophila vial to be inserted into the end to retrieve flies that are swept into it. At the time of writing this chapter, the only supplier of Drosophila nets is Genesee Scientific (www.geneseesci.com).

Preparing and using baits Carson and Heed (1983) present an excellent overview of different baiting strategies. A wide variety of Drosophila species can be attracted to bait made of fermenting fruit, particularly bananas. To maximize its attractiveness, bait should be prepared several days before it is to be set out. Although fruit bait becomes more attractive after it has fermented for several days, depending upon the substrate and the container used to trap flies, it may eventually become toxic to the flies. Baits that have been around for ten days should be monitored carefully for dead flies, and discarded. The only things required to prepare banana bait are bananas, live baker’s yeast, and water. Ripe bananas should be cut up, with or without their skins, placed in a plastic container, and covered with water. Dry active yeast should be added (one envelope to about 1 kg or 2 lb of bananas) and mixed with the water and bananas. The container should be covered overnight, stirred the next day, and allowed to ferment for an additional 24 hours. While the mixture is fermenting, the container should not be too tightly sealed. The mixture should be stirred again and covered. If any liquid remains, it should be decanted before the material is parceled out into bait containers. Fruits and vegetables in addition to bananas can be prepared for baiting in the same way. Specialist species are often attracted to standard banana baits, but many are not. Mushrooms can be effective baits not only for mycophagous Drosophila but also for many other species. John Jaenike (personal communication) prepares grocery-store mushrooms (Amanita) by soaking them for an hour in water and then placing piles of approximately ten mushrooms on the ground in a shady place near the base of a tree or fallen log. Flies are best collected from these baits in the mornings. Mushroom baits prepared in this way last for several days. In the first few days they attract adults, but later, as they decay, will contain Drosophila larvae that can be reared in the laboratory (see below). Species belonging to the large repleta group are primarily cactophilic. Cacti can be cut up and fermented using baker’s yeast, just as with the bananas. In nature, however, fermenting cacti have their own communities of yeasts (Starmer, 1981; Barker et al., 1983), many of which are available from the Herman J. Phaff Yeast Collection at the

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University of California at Davis (http://www.phaffcollection.org/home.asp) for use in preparing bait. While many of the cactophilic species are attracted to banana baits, larger and more diverse collections are obtained by using cacti in the baits (Markow, unpublished data). Pads of the mission prickly pear, Opuntia ficus-indica, a cactus widely used as an ornamental in the west and southwestern United States, are usually abundant in private yards. In order to obtain tissue of species of columnar cacti we recommend contacting local botanical gardens, as they are often willing, in our experience, to provide material from these species. A number of factors should be considered when making and placing baits. One problem is that other animals may be attracted to the baits, such as ants and wasps, and also large animals that can consume the bait. Baits can be made less accessible to large animals by hanging them on a cord from a tree branch, but depending upon the area this may not always be feasible. Ants are a problem in that they usually attack flies, and once a bait has been discovered by ants, it becomes worthless for collecting Drosophila. Tanglefoot (http://www.tanglefoot.com) makes products that can be applied to the area on the ground surrounding the bait itself or to the tree from which the bait hangs. Flies of some species, such as the mushroom breeders, prefer to be on the ground where mushrooms normally sprout up, so that this is where baits designed for these species should be placed. The exposure of the bait to the sun should also be considered. Flies tend to be most abundant at open baits in the morning and afternoon hours, but avoid direct sun. Furthermore, bait containers can heat up, killing the flies inside. Bait containers can be constructed from a number of different materials. Some investigators use paper cups, others prefer open buckets. Open buckets are typically harvested

Figure 4.3. Trap made from discarded liter plastic beverage bottle.

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using a net. Other devices such as plastic bags and closed containers can be especially useful. Investigators in the Markow laboratory have followed the practice of searching highway pull-outs for one-liter plastic soda bottles which have been improperly discarded, and making them into bait containers (Figure 4.3). Using a pocketknife, an opening is sliced into the side of the top half of the bottle. The bait material is introduced through this and placed in the bottom of the container, and twigs are inserted into the bait as perch sites. These bottles are hung from trees or shrubs by placing a piece of string around their necks. Caps should be screwed on tightly. This type of bait yields large numbers of flies because they tend to remain inside. When harvesting flies from these bottle baits, the side slit is sealed with a piece of tape and a shell vial is slipped over the top immediately after removing the cap. The 25
95 mm shell vial is the same size as the opening of the bottles, and tapping the bait bottle causes the flies to go up into the vial. New vials can be placed over the opening until most of the flies are collected. Remaining individuals can be removed with an aspirator.

Collecting from natural substrates Adults of many species can be collected directly from their feeding or breeding sites. For those species that are not attracted to baits, the only way to collect them is on their natural resources. Drosophila carbonaria, for example, can be seen feeding and ovipositing in the bleeding sap of mesquite trees, but will not come to bait placed nearby. Natural breeding sites are not known for all species, however. For some, but not all, species, feeding and breeding take place at the same site (Spieth, 1952). Adult and larval D. melanogaster, and D. simulans, for example can be collected from a variety of rotting fruits, and a number of species can be netted from compost piles. Mushroom-breeding flies and larvae are both collected from mushrooms that sprout up during the summer in forested areas. Many cactophilic species can be found in large numbers on necrotic cacti. Flower breeders can be found right on the flowers they use (Brncic, 1983). Collecting from natural substrates can be very time consuming, as fly densities are sometimes low and not all substrates located will be suitable for the species of interest – so be prepared to spend time searching. Collecting success will also depend upon the season. In tropical and more temperate areas, species may be active most or all of the year or primarily during the wet season. Species inhabiting colder elevations and latitudes often undergo only one or two generations during the summer. Searching collection records may reveal the month of the year in which certain species have been successfully found. The Tucson Stock Center houses collection records from many investigators, and makes these documents available to visiting scientists. If adult flies cannot be found, it may be possible to rear them from decaying material if larvae are present. In addition, it is often of interest to determine what species are using a particular resource by bringing the material into the laboratory and allowing adults to eclose over a period of time. Rotting material suspected of having Drosophila larvae can be placed in an aquarium or gallon jar, covered with cheesecloth, and observed daily, in the lab, for emergences. Any adult Drosophila should be aspirated out of the

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container daily in order to avoid losing them to predators that also may be in the material, or to the toxic effects of chemical changes in the decaying material. A hole can be made in the cheesecloth in order to introduce the aspirator, and the hole can be covered with a piece of masking tape. Good practices include dating the container and recording the number and sex of each species emerging each day.

Collecting and transporting Hawaiian Drosophilidae Although it was clear that the Hawaiian Islands were home to a diverse array of drosophilid flies (Zimmerman, 1958), early attempts to collect these species using the same collecting methods applied to mainland Drosophila yielded a very small diversity of the known fauna (Carson and Heed, 1983). In addition, standard Drosophila techniques were impractical in Hawaii, where it is extremely difficult to carry large amounts of rotting fruit baits on the steep, narrow trails leading to pristine rainforest. Traditional entomological techniques, such as general sweeping of vegetation, were far more successful than baiting. However, while these methods were good for taxonomic work, they tended to leave specimens in poor condition for establishing laboratory cultures required for genetic work. The initial failure of baiting was likely because of the extremely small population sizes and highly specific host requirements of the Hawaiian species. A variety of techniques have been developed in the past 40 years that have greatly increased the efficacy of collecting these species. When used in combination with sweeping and knowledge of host plants, we can much more completely sample the diversity of Hawaiian Drosophila. Kaneshiro first introduced the idea of using fermenting banana baby food as a substitute for rotting fruit. This is very potent bait; just one or two jars can establish as many bait stations as several pounds of rotting bananas. The baby food is transferred to a small plastic container, inoculated with yeast, and can easily be carried into the field in a backpack. The only requirement is that the container be “burped” periodically to prevent a mess. Once at the collecting site, the bait is spread on tree trunks and on the undersides of branches and fern fronds. Many collectors dye this bait red with food coloring, because Hawaiian Drosophila seem to be attracted to this color. Another fermented bait that has been used with wide success is the mushroom “tea” developed in the late 1970s (Kaneshiro et al., 1977). Commercial mushrooms are fermented with yeast for seven to ten days prior to collecting. Sponges, sometimes dyed red, are soaked in this mushroom tea and hung from the undersides of trees using pushpins. This creates highly pungent bait that attracts many species of Drosophila. This bait was so effective at attracting members of the haleakalae species group that 25 new species in this group alone were discovered over the course of a few years (Hardy et al., 2001). This is impressive considering that at this time (1977–1980) the Hawaiian Drosophila Project had been collecting intensively for more than fifteen years, and had not encountered these species at all. Sweeping of rainforest vegetation is also an effective way to collect many Hawaiian Drosophila species that do not come readily to bait. There are several ways to go about this. Because the Hawaiian rainforest is typically very wet, Carson and Heed (1983)

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suggest dragging heavy canvas “beating bags” through the vegetation. The window of the bag is then put up to the light and the desired flies picked off with an aspirator. This is an arduous method and it tends to damage many smaller flies, but the bag is still effective, no matter how wet it gets. Thinner, non-canvas nets can easily become saturated with water when using this technique, and will damage more flies than they collect. We recommend a standard entomological net (not the smaller Drosophila-specific net or the heavier beating bag), sweeping close to the forest floor and beneath overhanging vegetation, making an effort to avoid wetting the net too much. This method has proven highly effective, although not so much so when raining or when the forest is very wet. Having several changes of net on hand can relieve this problem. Baiting and sweeping are highly effective, but still miss a large number of species. Targeting specific host plants, either by looking for drosophilids on them in the field or by returning to the laboratory with rotting plant material and rearing adults from larvae, will increase the number of species collected. Rearing is particularly important because species are often not attracted to bait, or are not present on the host substrate when the collector looks. Larvae, however, are often present in large numbers. Some species groups, such as the modified mouthpart group, are rarely collected in large numbers without rearing (Kam, 1978; O’Grady et al., 2003). Once flies are collected in the field, getting them to the laboratory becomes another challenge. Collecting in wet forests is difficult because the cotton plugs typically used for Drosophila vials become saturated with water and flies can easily drown. Kaneshiro makes special cloth plugs to keep water from getting into vials (cut-up aloha shirts work best). A paper divider is usually inserted into the vial so flies have an area to perch and the plug does not slip down too far into the vial. Because Hawaiian Drosophila are highly adapted to this high-elevation rainforest habitat, they need to be maintained at low temperature (16°C) and high humidity (nearly 100%). Any deviation from this will lead to reduced fecundity, complete sterility or death. Furthermore, large picture winged Drosophila often get stuck in standard media. A minimal media – containing only a small amount of agar and sugar – has been developed. The insides of these vials are lined with absorbent paper to increase humidity and give the flies ample perching places to prevent them from getting stuck. Once in the lab, Hawaiian Drosophila provide additional culturing challenges (Chapter 8).

Permits Depending upon where the collecting is done and where the flies are going, permits may or may not be required. Obviously permits would not be required to collect flies from your own back yard or the property of someone you know. Usually orchards or other commercial operations such as wineries will give permission to collect on their property. Collecting in United States National Parks and Monuments, however, requires permission from the individual parks, which are listed at http://www.nps.gov. In many countries, collecting insects usually requires permission, which can take months to obtain. Many areas have field stations that can be located through http://www.iobfs.org, the International Organizations of Biological Field Stations. Services at some field stations

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include permits for collecting, in addition to providing a place to sort flies. Another strategy is to contact a local Drosophila biologist or entomologist to identify the appropriate agency from which to seek permission. International collecting trips provide investigators from nations with good research funding to help build the research capabilities of colleagues and students in regions with more limited resources. These interactions can include establishment of formal collaborations, presentations of seminars or technical demonstrations, donations of books, and offers to sponsor students in exchange programs. Many countries require permits to bring live insects or certain plant material across their borders. Each investigator should consult the regulations for their own country. Although Drosophila are not considered to be pests by the United States Department of Agriculture, this agency issues import permits that can be applied for through http://www.aphis.usda.gov/ppq/permits/.

Transporting live adults or larvae Most species can be transported successfully if certain practices are observed. Keeping flies at safe temperatures is one of the most important concerns. If vials of flies are left in a closed automobile, they can overheat and die in a short period of time. Vials are best labeled with collecting data directly on the glass with a marker, and bundled, a half dozen to a group, in several sheets of newspaper taped shut. These bundles should be stored in a cooler. It is not necessary to put ice in the cooler, but if ice is present then the bundles with the vials should not be in direct contact with the ice. Another issue is making certain that flies don’t either become dried out or get stuck in the food or in moisture that has accumulated in the vials. Certain food types or ambient conditions will lead to conditions inside vials that are too dry for certain species. Lining the vials with a piece of filter paper dampened with water will provide flies with a substrate to perch on while preventing vials from drying out. On the other hand, vials should always be checked for condensation inside and “mopped” of excess moisture prior to receiving flies. An excess of flies (more than fifteen) in a vial can lead to the accumulation of moisture from respiration, so it is important to try to keep densities low. Some food types are more prone to these sorts of problems. We recommend cornmeal-based food vials or agar vials for field collecting of flies to be sorted later, especially if the trip is long. Chapter 10 contains recipes for various culture media. On the other hand, if isofemale lines are to be set up in the field (see Chapter 6), banana-based foods (being softer) facilitate oviposition and feeding by first instar larvae. Depending upon the length of the trip, larvae may begin to develop in the vials, causing the food to become sloppy. Changing the vials after four or five days can reduce this problem, but the old vials should be kept to rear the offspring, remembering to identify them as F1 rather than the field collected flies themselves. A number of species that can be collected in nature cannot be cultured in the laboratory. If this is the case, the flies should be used for experiments before they die. For example, if genetic variation is to be examined, the flies should be identified and then either frozen (for allozymes) or placed in ethanol (for DNA). Chapter 6 gives more detail on handling wild-caught specimens.

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References Barker, J. S. F., Toll, G. L., East, P. D., Miranda, M. and Phaff, H. J. (1983). Heterogeneity of the yeast flora in the breeding sites of cactophilic Drosophila. Can. J. Microbiol. 29, 6–14. Brncic, D. (1983). Ecology of flower breeding Drosophila. In: The Genetics and Biology of Drosophila (M. Ashburner, H. L. Carson and J. N. Thompson, eds), Vol. 3d, pp. 333–377. Academic Press, London. Carson, H. L. and Heed, W. B. (1983). Methods of collecting Drosophila. In: The Genetics and Biology of Drosophila (M. Ashburner, H. L. Carson and J. N. Thompson, eds), Vol. 3d, pp. 2–31. Academic Press, London. Hardy, D. E., Kaneshiro, K. Y., Val, F. C. and O’Grady, P. M. (2001). Review of the haleakalae species group of Hawaiian Drosophila (Diptera: Drosophilidae). Bishop Museum Bull. Entomol. 9, viii  88 pp. Kam, M. W. Y. (1978). The biosystematics of the mimica subgroup of the modified mouthparts species group of Hawaiian Drosophila. Thesis, University of Hawaii at Honolulu, vi  63 pp. Kaneshiro, K. Y., Ohta, A. T. and Spieth, H. T. (1977). Mushrooms as bait for Drosophila. Drosophila Information Service 52, 85. O’Grady, P. M., Kam, M. W. Y., Val, F. C. and Perreira, W. D. (2003). Revision of the Drosophila mimica subgroup, with descriptions of ten new species. Ann. Entomol. Soc. Am. 96, 12–38. Spieth, H. T. (1952). Mating behaviour within the genus Drosophila (Diptera). Bull. Am. Mus. Nat. Hist. 99, 395–474. Starmer, W. T. (1981). A comparison of Drosophila habitats according to the physiological attributes of the associated yeast communities. Evolution 35, 38–52. Zimmerman, E. C. (1958). 300 species of Drosophila in Hawaii? A challenge to geneticists and evolutionists. Evolution 12, 557–558.

CHAPTER 5

Distribution Contents

• •

Distribution maps References

Patterson and Stone’s (1952) chapter on Drosophila distributions was probably the first, and arguably the best, synthesis of locality data in this genus to date. They covered the distributions of all species in the genus Drosophila known at the time, over 600 species. Although subsequent workers have done regional treatments of a given biogeographic province (Bachli and Rocha-Pite, 1981; Bock and Parsons 1981; Hardy and Kaneshiro 1981; Okada 1981; Tsacas et al., 1981; Val et al., 1981; Wheeler, 1981) or taxonomic treatments focusing on a single group (Erhman and Powell, 1982; Lakovaara and Saura 1982; Throckmorton, 1982; Wasserman 1982; Lemunier et al., 1986; Lachaise et al., 1988), none have the breadth of the Patterson and Stone volume. Wheeler’s catalog of Drosophilidae (1982, 1986) includes locality data for all species and was invaluable in preparing this chapter, but it lacks detailed maps. Here we attempt to synthesize what is currently known about the distribution of taxa present in culture at the Tucson Stock Center. We also cover the close relatives of these species in order to provide a more complete picture of distributions within the various species groups in the genus Drosophila. Good distributional data are critical to understanding population structure, speciation, and change in community composition over time. This latter point is particularly important now that widespread changes in climate are taking place. Drosophila species can serve as indicators of such events, although only a few studies have attempted this to date (Grimaldi et al., 2000). Several caveats concerning the data presented in this chapter should be stated clearly, however. Distributional records were gleaned from disparate sources such as species descriptions, regional catalogs, and collection records generated over the past 60 or so years. It is certain that additional data on some species exists and was missed in our literature survey. Therefore the maps we show here (Figures 5.1–5.44) are by no means complete or completely accurate, and are not intended to be either. Instead, they should be viewed as approximations of the complex, and constantly changing, populations that make up each species and species group. Furthermore, because some of the records used are very old, there is no guarantee that present-day species compositions are at all similar.

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The other point that should be made is that survey data of this type are, by their very nature, heterogeneous. Unequal attention has been given to some countries (United States, Mexico, Brazil) while other places (Belize, Guatemala, Honduras, Venezuela, and others) remain virtually unknown. For example, Patterson and his colleagues collected extensively in Mexico, and we know more about the fauna of this country than nearly any other. The Guianas, on the other hand, have never been surveyed systematically for Drosophila. Therefore species diversity measures gleaned from these maps may be the result of historical sampling bias rather than actual numbers of species. In spite of the preliminary nature of the data we present here, many useful patterns can be drawn from it. For example, a number of species groups have similar distributions: from the Mexican plateau in the north, through Central America, and down the east side of the Andes into the Amazon Basin of northern Brazil. This is common, and was hypothesized by Throckmorton (1975) to be the result of numerous connections, breaks, and reconnections of the Isthmus of Panama. Modern molecular methods have not been widely applied to such questions, and will undoubtedly be used to investigate the change in Drosophila species compositions over time as well as identify barriers to gene flow that biogeography alone cannot. It is our hope that this chapter will stimulate additional Drosophila collecting in areas that have been traditionally under collected. We also hope that those interested in biogeography begin to examine the rich and complex diversity present in the genus Drosophila.

Distribution maps

Figure 5.1. Distribution of selected taxa in the annulimana species group. The following species are shown: D. gibberosa (vertical lines), D. aracataca and D. talamancana (horizontal lines), D. breuerae (backslashes), D. annulimana (dots), and a number of Brazilian species with (squares).

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Figure 5.2. Distribution of selected taxa in the bromeliae, carbonaria, and carsoni species groups. Ranges of D. bromeliae are shown with crosshatches, D. bromelioides with horizontal lines. The carbonaria (dots) and carsoni (forward slashes) species groups are both monotypic and poorly studied.

Figure 5.3. Distributions of selected taxa in the canalinea species group: D. canalinea (horizontal lines), D. canalinoides (vertical lines), D. annularis (forward slashes), D. paracanalinea (dots), and D. albomarginata (crosshatches). Note significant area of sympatry in the isthmus region.

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Figure 5.4. Distribution of Nearctic and Neotropical species in the coffeata, dreyfusi and melanica groups. The coffeata group is a small, Neotropical clade of four species. The range of one species in this group, D. fuscolineata, is shown with dots. The dreyfusi species group is also Neotropical in distribution. Drosophila boliviana (crosshatches) is from Bolivia, D. fuscipennis and D. lugubripennis (forward slashes) are known from Peru, D. camargoi and D. briegeri (squares) are widely distributed from Costa Rica and Panama, respectively, to Brazil. Three ranges of three species found only in Brazil, D. dreyfusi, D. krugi, and D. wingei are also shown with squares. The distributions of other dreyfusi species fall within the ranges of those shown. The melanica group is Holarctic in distribution. Representative distributions of some Nearctic species are shown here: Drosophila paramelanica (horizontal lines), D. melanica (vertical lines), D. melanissima (ovals), and D. micromelanica (backslashes).

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Figure 5.5. Distribution of selected Asian and European species in the melanica group. Drosophila afer (forward slashes), D. bisetata and D. longiserrata (dots), and D. tsigana (crosshatches) are shown.

Figure 5.6. Distributions of the following species in the mesophragmatica group are shown: D. mesophragmatica (horizontal lines), D. brncici (forward slashes), D. orkui (backslashes), D. altiplanica (dots), and D. pavani (vertical lines). Drosophila gaucha (not shown) is widely distributed in South America.

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Figure 5.7. Distribution of the nannoptera and peruviana species groups. The peruviana group is monotypic and known only from Peru (forward slashes). The nannoptera group contains four described species and is known predominantly from Mexico. Drosophila pachea (horizontal lines) is found in northern Mexico and the southwest United States where it is tightly associated with its host plant Senita. Drosophila acanthoptera (dots) is known from southern Mexico and Venezuela. Drosophila wassermani (crosshatches) and D. nannoptera are sympatric with D. acanthoptera in southern Mexico.

Figure 5.8. Distribution of selected Asian species in the robusta and polychaeta groups: D. cheda (horizontal lines), D. moriwakii, D. neookadai, D. okadai and D. pseudosordidula (forward slashes), D. asper (vertical lines), D. sordidula (dots), and D. lacertosa (squares).

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Figure 5.9. Distribution of some species in the polychaeta group. The polychaeta species group is quite widespread and probably represents an old lineage that is basal to other members of the genus Drosophila. Drosophila fraburu (dots) is known from western Africa. Drosophila hirtipes is endemic to the Seychelles and D. polychaeta is subcosmopolitan.

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Figure 5.10. Distribution of selected taxa in the mercatorum and fasciola subgroups of the repleta species group. Ranges are as follows: D. peninsularis (ovals), D. carcinophila and D. paraguttata (backslashes), D. fulvalineata (dots), D. paranaensis (forward slashes) and D. fasciola (horizontal lines). Drosophila mojuoides is known only from Trinidad. Drosophila coroica (crosshatches) is known only from Bolivia. Drosophila pictura has been collected in both Trinidad and Bolivia. Drosophila pictilis is found only in El Salvador. Most other taxa in this group fall within the ranges shown. Drosophila mercatorum (not shown) is widespread in distribution and should be considered subcosmopolitan.

Figure 5.11. Distribution of selected taxa in the inca and hydei subgroups of the repleta species group: D. bifurca (dots), D. nigrohydei (horizontal lines), D. eohydei (backslashes), D. neohydei (forward slashes), D. guayllambambae and D. inca (crosshatches), and D. novemarista (vertical lines). Drosophlia hydei (not shown) is cosmopolitan.

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Figure 5.12. Distribution of selected species in the repleta subgroup of the repleta group: D. fulvimacula (vertical lines), D. fulvimaculoides (dots), D. linearepleta and D. neorepleta (horizontal lines), D. limensis (forward slashes). Drosophila repleta (not shown) is cosmopolitan.

Figure 5.13. Distribution of species in the anceps and meridiana complexes (repleta group, mulleri subgroup). Ranges of the following taxa are shown: D. nigrospiracula (vertical lines), D. anceps (backslashes), D. leonis (dots), D. meridiana (horizontal lines), D. promeridiana (forward slashes), and D. meridionalis (squares).

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Figure 5.14. Distribution of species in the eremophila and stalkeri complexes (repleta group, mulleri subgroup). The following species are depicted: D. mettleri (forward slashes), D. eremophila (horizontal lines), D. micromettleri (dots), and D. stalkeri (vertical lines). Drosophila richardsoni is known from Puerto Rico to the Lesser Antilles.

Figure 5.15. Distribution of species in the mojavensis and mulleri clusters (repleta group, mulleri subgroup, mulleri complex), showing the ranges of D. wheeleri (crosshatches), D. mojavensis (vertical lines), D. navojoa (dots), D. arizonae (forward slashes), D. aldrichi (squares), D. huaylasi (horizontal lines), and D. nigrodumosa (backslashes). Ranges of D. parisiena and D. straubae (ovals) are also shown. Drosophila mayaguana is widespread in the Caribbean.

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Figure 5.16. Distribution of the ritae cluster (repleta group, mulleri subgroup, mulleri complex) and several miscellaneous species in the repleta group. The following ranges are shown: D. ritae (forward slashes), D. mathisi (backslashes), D. desertorum (dots), D. nigricruria (horizontal lines), and D. hamatofila (vertical lines).

Figure 5.17. Distribution of species in the longicornis cluster (repleta group, mulleri subgroup, mulleri complex). Drosophila longicornis (horizontal lines), D. mainlandi (vertical lines), D. propachuca (forward slashes) and D. pachuca (dots) are shown. The range of an as yet undescribed species, “D. sonorae” is shown with ovals.

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Figure 5.18. Distribution of additional species in the repleta group: D. hexastigma (forward slashes), D. pegasa (dots) and D. spenceri (vertical lines).

Figure 5.19. Distribution of species in the martensis complex (repleta group, mulleri subgroup). Several species are shown: D. martensis (forward slashes), D. starmeri, D. uniseta, and D. venezolana (dots), D. boroborema and D. serido (horizontal lines), and D. koepferae (vertical lines).

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Figure 5.20. Distribution of the calloptera group and two Nearctic species in the robusta group. The following species are shown: D. ornatipennis (ovals), D. kallima (dots), D. calloptera (vertical lines), D. maracaya (squares), D. robusta (forward slashes), and D. colorata (backslashes). Other robusta species are found in Asia (Figure 5.8) and Europe.

Figure 5.21. Distribution of the tumiditarsis group and the Asian species in the virilis group. Drosophila repletoides (forward slashes) is recorded from central China and Japan. Drosophila kanekoi (dots) and D. ezoana (backslashes) are both known from Japan. Other taxa in the virilis group are known from North America (Figure 5.22) and Europe.

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Figure 5.22. Distribution of the Nearctic species of the virilis group is as follows: D. montana (vertical lines), D. flavomontana (horizontal lines), D. novamexicana (dots), D. lacicola (backslashes), D. americana (forward slashes), and D. borealis (ovals). Other species in the virilis group are found elsewhere in the Holarctic (Figure 5.21).

Figure 5.23. Distribution of selected species in the quinaria, testacea, and cardini species groups. Drosophila putrida (testacea group) is found in the central and eastern United States and into southeastern Canada (vertical lines). In the quinaria group the ranges of D. quinaria, D. guttifera and D. recens (vertical lines) overlap with that of D. palustris (dots) in the central and northeastern portions of North America. Drosophila magniquinaria (forward slashes) is known from the Pacific Northwest. The cardini group is mostly Neotropical; D. cardinoides (horizontal lines) is distributed from Mexico to Brazil, D. neocardini (ovals) is found in Colombia and Brazil, and D. procardinoides (backslashes) has been collected in Peru and Bolivia. Two species, D. bedichecki and D. belladunni are endemic to Trinidad and Jamaica (respectively). Drosophila neotestacea (testacea group, not shown) is known from the northern half of the United States, Canada, and Alaska.

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D. cardini

D. dunni D. arawakana D. caribiana D. antillea D. similis

D. nigrodunni

D. parthenogenetica, D. neomorpha

D. polymorpha

Figure 5.24. Distribution of some species in the cardini group. Drosophila cardini (ovals) is widespread in North and South America and the Caribbean. Drosophila parthenogenetica and D. neomorpha are shown with forward slashes, D. polymorpha with vertical lines. A number of species in the dunni subgroup are endemic to single islands in the Antilles.

Figure 5.25. Distribution of species in the guarani group. Ranges of D. guaraja, D. guarani, and D. guaru (horizontal lines), D. griseolineata (dots), D. araucana and D. huilliche (crosshatches), D. peruensis (backslashes), D. alexanderi (squares) and D. limbinervis (ovals) are shown.

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Figure 5.26. Distribution of some species in the immigrans group. D. pulaua is known from Borneo, D. neohypocausta from Taiwan and D. pararubida from the Philippines. Drosophila signata (backslash) is known from Taiwan and Borneo and D. kepulauana (forward slash) is known from Borneo and the Philippines.

Figure 5.27. Distribution of some species in the immigrans group. Drosophila formosana and D. albomicans are distributed from Taiwan to southeast Asia (dots), D. kohokoa is from southeast Asia and Indonesia, D. nasuta (horizontal lines) is thought to be from eastern Africa, Madagascar, and the Seychelles, although this species has become subcosmopolitan in the last 100 years. Two additional species, D. sulfurigaster and D. hypocausta, are widespread from southeast Asia into Micronesia. Drosophila immigrans is cosmopolitan.

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Figure 5.28. Distribution of New World species in the pallidipennis and quinaria groups. The distribution of D. pallidipennis is shown with vertical lines. Ranges of the following quinaria group species are shown: D. innubila, D. suffusca, D. tenebrosa, and D. munda (horizontal lines), D. rellima (backslashes), D. occidentalis and D. suboccidentalis (dots), D. falleni (squares), and D. deflecta (forward slashes).

Figure 5.29. Distributions of selected Asian species in the quinaria and testacea groups: D. parakuntzei (backslashes), D. angularis, D. unispina, D. takadai and D. nigromaculata (forward slashes, D. brachynephros (dots), and D. mediobandes (squares). Drosophila orientacea (testacea group) is known only from the Japanese island of Hokaido.

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Figure 5.30. Distribution of species in the tripunctata group. The following species, all in culture at the Tucson Stock Center, are shown: D. tripunctata (forward slashes), D. unipunctata (vertical lines), D. mediopictoides (dots), D. mediostriata (horizontal lines). Drosophila mediodiffusa is widely distributed in the Caribbean.

Figure 5.31. Distribution of Caribbean and some South American saltans species. Several species are single island endemics, including D. dacunhai from Jamaica, D. lusaltans from Haiti, D. milleri from Puerto Rico, and D. pulchella from St. Vincent Island. Drosophila saltans is known from Cuba and from Mexico through Costa Rica. Drosophila cordata is found only in Guatemala and D. septentriosaltans from Colombia.

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Figure 5.32. Distribution of the South American species of the saltans group. A number of saltans species have only been recorded from Brazil (crosshatches) and many taxa are sympatric in this country. Drosophila sturtevanti (horizontal lines) is known from Mexico to Brazil and the West Indies. The range of D. emarginata (vertical lines) is from Mexico to Peru. Drosophila prosaltans (ovals) is known from southern Mexico to Brazil and Paraguay. Drosophila rectangularis and D. elliptica have only been collected in Mexico. Other species (not shown) fall within the ranges shown.

Figure 5.33. Distribution of species in the willistoni species group. The ranges of several species are shown: D. sucinea (forward slashes), D. fumipennis and D. paulistorum (dots), D. capricorni (horizontal lines), and D. pavlovskiana (crosshatches). Drosophila insularis is found in the West Indies. Several taxa are very widespread. Drosophila willistoni and D. nebulosa (not shown) both extend from the southern United States into the Caribbean, and throughout South America. Drosophila equinoxialis and D. tropicalis (not shown) both extend from El Salvador to Brazil, with the latter also being found in the West Indies and Bolivia.

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Figure 5.34. The affinis and pseudoobscura subgroups are endemic to the New World. Drosophila affinis (horizontal lines) is found from southern Canada into the central and eastern United States as far south as Georgia. Two other species, D. narragansett and D. algonquin (not shown), fall within the range of D. affinis, but are not found south of New England. Two species, D. azteca (dots) and D. tolteca (vertical lines), are found in both the Nearctic and Neotropical regions. The range of D. athabasca (ovals) extends south from Alaska and southwest Canada into the central United States and Pacific northwest. Within the pseudoobscura subgroup, D. pseudoobscura (backslashes) is most widespread. Several species including D. persimilis (forward slashes), D. miranda (not shown), and D. lowei (not shown) fall within the range of D. pseudoobscura in the western and southwestern United States. Many obscura and subobscura subgroup species are widespread in Europe (not shown). These include D. ambigua, D. obscura, D. subsilvestris, and D. tristis. Drosophila bifasciata is widespread in the Palearctic. Drosophila subobscura is cosmopolitan, having spread from its traditional range in the Palearctic into the Neotropical and Nearctic regions. One species, D. guanche, is found only on the Canary Islands.

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Figure 5.35. Distribution of species in the ananassae subgroup (melanogaster species group). Drosophila ananassae and D. malerkotliana (not shown) are both widespread in distribution with the former being mainly circumtropical and the latter considered subcosmopolitan. A number of other species are quite widely distributed in the Oriental and Oceanic regions. These include two species, D. bipectinata (horizontal lines) and D. pseudoananassae (forward slashes), that are widespread in India, southeast Asia, and several island groups in the Oriental region. Drosophila pseudoananassae is also found in Australia. Other taxa are more restricted, including D. parabipectinata (vertical lines) from southeast Asia, D. varians (dots) from the Philippines, D. pallidosa and D. phaeopleura from Fiji and surrounding islands, and D. ercepeae from Reunion Island.

Figure 5.36. Distribution of species in the suzukii subgroup (melanogaster species group). Drosophila mimetica (ovals) is known from Malaysia, D. lucipennis (dots) is disjunctly distributed in eastern India and Taiwan, D. biarmipes (forward slashes) is known from India and Sri Lanka to southeast Asia, and D. pulchrella (vertical lines) is found from India, China, and southeast Asia to Japan.

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Figure 5.37. Distribution of species in the takahashii subgroup (melanogaster species group). Drosophila takahashii (vertical lines) is most widespread, being found from India to Japan and into Micronesia. Drosophila lutescens (forward slashes) is known from Japan and Korea, D. paralutea (backslashes) from Thailand and Borneo, D. prostipennis (dots) is disjunctly distributed in Taiwan and India, and D. pseudotakahashii (horizontal lines) is known from Australia.

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Figure 5.38. Distribution of species in the melanogaster species group (melanogaster subgroup). Drosophila erecta (horizontal lines) is known from central Africa, D. orena (forward slashes) from Cameroon, D. yakuba (vertical lines) is widespread in Africa, and D. teissieri (dots) is known from both central and western Africa. Two members of this subgroup (not shown), D. melanogaster and D. simulans, are cosmopolitan. A number of other species (not shown) are island endemics including D. mauritiana (Mauritius Is.), D. seychellia (Seychelles Is.) and D. santomea (Santomea Is.).

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Figure 5.39. Distribution of selected African species in the montium subgroup (melanogaster species group). Drosophila seguyi (not shown) is widespread in Africa. A number of other species are sympatric over a large portion of central and western Africa, including D. diplacantha, D. greeni, D. nikananu, and D. tsacasi. Drosophila vulcana (forward slashes) is known from Zaire and southern Africa.

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Figure 5.40. Distribution of selected Oriental taxa in the montium subgroup (melanogaster species group). Drosophila auraria (forward slashes) is from Japan, Korea, and China, D. lacteicornis is from Okinawa Island, D. mayri is from New Guinea, D. baimaii (backslashes) is from southeast Asia, D. barbarae (dots) is from southeast Asia, Borneo, and the Philippines, and D. birchii (crosshatches) and D. serrata (horizontal) are both from New Guinea and Australia.

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Figure 5.41. Distribution of selected Oriental species in the montium subgroup (melanogaster species group). Drosophila jambulina (vertical lines) is widespread from India to southern China and southeast Asia, D. biauraria (forward slashes) is found in Japan and Korea, D. lini and D. quadraria are endemic to Taiwan, D. pennae is from New Guinea, D. kanapiae is found in the Philippines, D. bicornuta (dots) is from southeast Asia, the Philippines, and Borneo. Another species, D. kikkawai (not shown), is circumtropical in distribution.

Figure 5.42. Distribution of selected Oriental species in the montium subgroup (melanogaster species group). Drosophila punjabiensis (horizontal lines) ranges from India into southeast Asia, D. rufa (vertical lines) is widespread from India to Japan, D. triauraria (ovals) is from Japan and Korea, D. parvula (forward slashes) and D. orosa (dots) are both from southeast Asia, with the former also being found in Malaysia.

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Figure 5.43. Distribution of the elegans, eugracilis and ficusphila species subgroups (melanogaster species group). Drosophila elegans is from the Philippines and New Guinea, D. ficusphila is fairly widespread from India to Japan and Korea, and D. eugracilis is found in the Indian subcontinent, throughout southeast Asia, and into Australia.

References Bachli, G. and Rocha-Pite, M. T. (1981). Drosophilidae of the Palearctic Region. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3a, pp. 169–196. Academic Press, London. Bock, I. R. and Parsons, P. A. (1981). Species of Australia and New Zealand. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3a, pp. 291–308. Academic Press, London. Ehrman, L. and Powell, J. R. (1982). The willistoni species group. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3b, pp. 193–225. Academic Press, London. Grimaldi, D. A., Quinter, E. L. and Nguyen, T. (2000). Fruit flies as ecological indicators: species diversity and abundance of Drosophilidae (Diptera) along an altitudinal transect in the Parc National de Marojejy, Madagascar. Fieldiana Zool. 97, 123–135. Hardy, D. E. and Kaneshiro, K. Y. (1981). Drosophilidae of Pacific Oceania. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3a, pp. 309–348. Academic Press, London. Lachaise, D., Cariou, M.-L., David, J. R., Lemeunier, F., Tsacas, L. and Ashburner, M. (1988). Historical biogeography of the Drosophila melanogaster species subgroup. In: Evolutionary Biology (M. K. Hecht, B. Wallace and G. T. Prance, eds), pp. 159–225. Plenum Press, New York.

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181 Lakovaara, S. and Saura, A. (1982). Evolution and speciation in the obscura species group. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3b, pp. 1–59. Academic Press, London. Lemeunier, F., David, J. R., Tsacas, L. and Ashburner, M. (1986). The melanogaster species group. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3e, pp. 147–256. Academic Press, London. Okada, T. (1981). Oriental species, including New Guinea. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3a, pp. 261–290. Academic Press, London. Patterson, J. T. and Stone, W. S. (1952). Evolution in the Genus Drosophila. Macmillan, New York. Throckmorton, L. H. (1975). The phylogeny, ecology, and geography of Drosophila. In: Handbook of Genetics (R. C. King, ed.), Vol. 3, pp. 421–469. Plenum Press, New York. Throckmorton, L. H. (1982). The virilis species group. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3b, pp. 227–296. Academic Press, London. Tsacas, L., Lachaise, D. and David, J. R. (1981). Composition and biogeography of the Afrotropical Region. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3a, pp. 1–105. Academic Press, London. Val, F. C., Vilela, C. R. and Marques, M. D. (1981). Drosophilidae of the Neotropical Region. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3a, pp. 123–168. Academic Press, London. Wasserman, M. (1982). Evolution of the repleta group. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3b, pp. 61–139. Academic Press, London. Wheeler, M. R. (1981). Geographical survey of Drosophilidae: Nearctic species. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3a, pp. 99–122. Academic Press, London. Wheeler, M. R. (1982). Drosophilidae: a taxonomic overview. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3b, pp. 1–105. Academic Press, London. Wheeler, M. R. (1986). Additions to the catalog of the world’s Drosophilidae. In: The Genetics and Biology of Drosophila (M. Ashburner, J. N. Thompson and H. L. Carson, eds), Vol. 3e, pp. 395–409. Academic Press, London.

CHAPTER 6

Handling wild-caught specimens Contents

• • •

Living cultures Vouchering and preserving Drosophila References

Depending on the purpose for making collections of wild flies, specimens will be handled differently. Some investigators will be interested in having living material for experimental work, while others will want only DNA. In any case, if at all possible having both living and properly preserved material with standard documentation, from any collection, can be of great value to the research community as well as to the individual collector. Collecting wild specimens, especially from remote areas, is an expensive endeavor, and if live material can be transported and deposited in the Tucson Stock Center, a larger number of investigators will benefit from the effort. Similarly, if questions arise later as to species identification or geographic distribution they can be resolved if properly preserved, vouchered specimens also are available. Below we discuss approaches to handling living and preserved wild-caught specimens.

Living cultures When making living collections, the field notes should always include the species, the name of the collector, the date of the collection, detailed location of the collection (including GPS coordinates if possible), and what the flies were collected from – such as the natural substrate or the type of bait. Two types of living cultures are established from wild collections: multifemale and isofemale strains. Multifemale strains are established by pooling a number of flies, including males, of a given species from a given locality. The objective, when establishing a multifemale culture or strain, is to capture a high level of genetic variation. If possible, the number of females should be noted on the culture, along with the other collection information. A potential problem with starting multifemale strains in the field is obtaining the correct identification of the flies. Without a microscope, this might be impossible. For many related species, such as D. melanogaster and D. simulans, or D. pseudoobscura and D. persimilis, females are indistinguishable even with a microscope, so that simply pooling flies which look alike could produce mixed-species cultures. Flies could then

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potentially hybridize in the culture, or one of the species might be out-competed by the other. Ideally, females should be separated into isofemale lines (see below) and either the larval chromosomes or the male progeny used to verify species prior to pooling into a multifemale culture. If this is not feasible in the field – for example, if large numbers of vials are not available – isofemale cultures can be set up when the wild-caught flies are returned to the laboratory. In addition to their utility in validating species identification, isofemale strains are ideal for other purposes. By immediately separating wild-caught females into individual culture vials, strains are created which contain only the genetic variability carried by a single female, including however many different males she may have mated with. Isofemale strains then, as a rule, possess more limited nuclear genetic variability than found in a multifemale culture. Some investigators use isofemale strains or inbred lines descended from them in quantitative genetic studies of different traits. At the same time, however, isofemale strains should contain only one mitochondrial genome, which makes them indispensable for population genetic and phylogeographic studies. Isofemale lines should always be set up with field-caught flies, rather than with the descendants of flies transported en masse to the laboratory, as the F1 are more likely to be related and a set of strain derived from them would give lower estimates of genetic variability. Because there will be fewer F1 individuals to work the culture medium in a vial founded by a single versus multiple females, care should be taken to maximize oviposition when setting up isofemale lines. Often scratching the surface of the food with the end of a brush and providing a few grains of live yeast will stimulate females to lay eggs. Remember that yeast should not be used for mycophagous species, which require just a fresh mushroom (Chapter 9). For cactophilic Drosophila, adding a small cube of the cactus from which they were collected can promote oviposition and healthy larvae. If cactus is to be added, it should first be softened by cooking if possible. Wild-caught flies may carry problematic microbes, nematodes or mites. When returning from the field, wild cultures can be isolated and monitored for a generation. Chapter 11 provides tips for dealing with these problems should they arise.

Vouchering and preserving Drosophila Pinned, ethanol preserved, and/or cryopreserved voucher material is critical to, and notably absent from, many studies of Drosophila systematics and ecology. Having pinned voucher material allows verification of the species identity of study organisms at any time after the study has completed. With large studies involving the collection of many individuals, populations or species, proper vouchering and curation of pinned or otherwise preserved material is essential. Specimens are usually transported back to the laboratory alive, preserved in ethanol, or frozen. All relevant collection information should be recorded in the field notes (above), and these data should be linked to specimens with a collection number. Vials containing specimens should have this number marked on them in permanent ink. Keep in mind that many “permanent” markers will easily rub off when frozen, wet

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with condensation, or when ethanol is accidentally spilled on them. Plan to use ink that is suitable for your preservation and collection methods. Most Drosophila are small enough to be point-mounted, rather than pinned through the body. This is done by taking a standard paper insect point (see Chapter 12) and bending it slightly at the tip with a pair of forceps. An adhesive, either an insect-specific mounting material or clear nail polish, is then applied sparingly to the bent portion of the tip. The fly should be oriented on a clean piece of paper, right side up and with the head toward the top of the sheet. The tip of the point is then touched to the right pleurae of the fly such that the point of the bent tip is directed ventrally on the fly. The adhesive will bind the fly to the paper point so that the entire left side of the fly can be viewed. A number 3 insect pin is then positioned through the wide end of the point using an insect pinning block (Figure 6.1). Collection information is entered on a small label and placed on the same pin as the fly. The collection information should include the locality, date of collection, collecting method (baiting, aspirating, sweeping) and substrate, collector, and any collection numbers or stock numbers associated with the material (Figure 6.1). Once back in the lab, material is pinned and labels are printed, based on field note data. Many researchers are now using palm pilots or other portable computers to record field data. This is, in many ways, better than traditional field notes, but care should still be taken to back up the collection data. Many researchers, particularly those interested in molecular studies, preserve flies in ethanol or by freezing for further use. This is perfectly acceptable, provided that there is also a pinned voucher (above) linked to the spirit or frozen collection. After being frozen or pickled for a number of years, certain species identifications will be impossible and pinned material will be required for proper identification. It is probably most conservative to acquire the freshest, or best-preserved, samples possible (Dessauer et al., 1986, 1996; Prendini et al., 2002). Studies on ancient and old DNA samples suggest that denatured proteins or degraded DNA can hamper PCR amplification,

Fly on point Collection information Host/ecological information Collector information

Determination label Foam base Figure 6.1. A pinned Drosophila specimen, with labels for collection information. Standard labelling.

Handling wild-caught specimens

185

because of either inhibitory proteins or short templates (Soltis and Soltis, 1993; Handt et al., 1994; Thomas and Paabo, 1994). Cryopreservation in liquid nitrogen dry shippers is the preferred method to obtain high-quality tissue suitable for DNA and RNA work. However, for international collecting or extended domestic trips this method is untractable for a number of reasons, including difficulty in obtaining and maintaining liquid nitrogen, airport security, and shipping restrictions. Even collectors with the proper permits and airplane-safe dry shippers have had problems returning samples to the United States. Preservation in 70–100% ethanol is an acceptable alternative (Prendini et al., 2002). High-quality alcohol is readily available in all but the most remote locales, and can easily be transported by the researcher. Although tissue quality will be slightly degraded, transferring to cold storage or isolating nucleic acids as soon as possible after returning from the field will mediate these problems. Making considerations for long-term storage is also important, as DNA will degrade slowly over time. ETOH material stored at ambient temperature begins to degrade after six weeks (Reiss et al., 1995), and is probably best used within five years. Freezing ethanol-preserved material will slow degradation. Material stored at colder temperatures (
20°C and below) will degrade less and last longer. Vapor-phase liquid nitrogen material will preserve for over 25 years, although the preparation method will influence the quality and yield of the nucleic acids, and the use of a cryoprotectant is advocated (Prendini et al., 2002).

References Dessauer, H. C., Menzies, R. A. and Fairbrothers, D. E. (1994). Procedures for collecting and preserving tissues for molecular studies. In: Collections of Frozen Tissues: Value, Management, Field and Laboratory Procedures, and Directory of Existing Collections (H. C. Dessauer and M. S. Hafner, eds), pp. 21–24. Association of Systematics Collections, University of Kansas Press, Lawrence. Dessauer, H. C., Cole, C. J. and Hafner, M. S. (1996). Collection and storage of tissues. In: Molecular Systematics, 2nd edn (D. M. Hillis, C. Moritz and B. K. Mable, eds), pp. 29–47. Sinauer, Sunderland. Handt, O., Hoss, M., Krings, M. and Paabo, S. (1994). Ancient DNA – methodological challenges. Experientia 50, 524–529. Prendini, L., Hanner, R. H. and DeSalle, R. (2002). Obtaining, storing, and archiving specimens and tissue samples for use in molecular studies. In: Techniques in Molecular Systematics and Evolution (G. Giribet, W. C. Wheeler and R. DeSalle, eds), pp. 176–248. Birkhauser Press, Basel. Reiss, R. A., Schwert, D. P. and Ashworth, A. C. (1995). Field preservation of Coleoptera for molecular genetic analyses. Environ. Entomol. 24, 716–719. Soltis, P. S. and Soltis, D. E. (1993). Ancient DNA, prospects and limitations. NZ J. Botany 31, 203–209. Thomas, W. K. and Paabo, S. (1994). DNA sequences from old tissue remains. In: Methods in Enzymology, Vol. 224, Molecular Evolution: Producing the Biochemical Data (E. A. Zimmer, T. J. White and R. L. Cann, eds), pp. 406–419. Academic Press, San Diego.

CHAPTER 7

Life history variation Contents

• • • • •

Egg to adult development times Age at sexual maturity Fecundity and fertility differences Life history variation and ecology References

Although limited in scope and number, studies of life history characters in Drosophila have exposed considerable interspecific variability. Some of this variability has direct relevance to the ability of investigators successfully to rear and utilize different species in empirical studies. Two characters in particular – development time and adult age at reproductive maturity – are particularly important. In this chapter, we provide development times for a large number of the species maintained in the Tucson Drosophila Species Stock Center. For a subset of these species, we also provide ages at which flies become sexually mature. In Chapter 8, the implications of these species differences for rearing and for collecting virgin flies for experiments will be described. Issues of experimental design may be influenced by life history and behavioral variation, and these will be discussed in detail in Chapter 10.

Egg to adult development times Table 7.1 gives egg to adult development times for 198 species, with similar information for seven Hawaiian species being presented in Table 7.2. Most of the species are available from the Tucson Drosophila Species Stock Center, where the majority of the data were collected and where stocks are maintained at 18°C. For a number of species, however, development times are available from studies using higher temperatures. It is clear from these examples that development time is faster when flies are reared at higher temperatures. It should be kept in mind when reviewing these data that studies from different laboratories were likely to have used different larval densities, which may contribute to the variability in development time. Crowded cultures tend to exhibit delayed development (Santos et al., 1994; Borash et al., 2000). Overall development times of Drosophila species in Table 7.1 range from 8 days for D. ananassae to about 24 days for species of the virilis group. These differences are likely to reflect not only the evolutionary relationships of these species, but also their different geographic ranges and ecologies.

190

How to use living flies Table 7.1.

Egg to adult development time of non-Hawaiian Drosophilids

Species

Egg–adult

Temperature (°C)

Reference no.

D. acanthoptera D. affinis D. affinis D. albomicans D. algonquin D. americana D. ananassae D. ananasse D. anceps D. antillea D. arawakana D. arizonae D. athabasca D. auraria D. austrosaltans D. baimaii D. barbarae D. biarmipes D. biauraria D. bicornuta D. bifasciata D. bifurca D. bipectinata D. birchii D. borborema D. borealis D. bromeliae D. busckii D. busckii D. busckii D. buzzatii D. canadiana D. capricorni D. cardinoides D. caribiana D. crocina D. dacunhai D. diplacantha D. dunni D. elegans D. ellisoni D. eohydei D. equinoxialis D. ercepeae D. erecta D. eremophila D. ezoana

16 15.5 16 11.5 18 18 13 8 15 15 14.5 15.5 16 18 16.5 17.5 14 17.5 16 17 14.5 17.5 14.5 17 13 20 15 16.5 12 15 18 22 15 15 16 16 21 18.5 15 12 15.5 18 17 16.5 14.5 14.5 17

18 18 25 18 25 18 18 25 18 18 18 18 25 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 25 21 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18

1 1 4 1 4 1 1 8 1 1 1 1 4 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1

Life history variation Table 7.1.

191

(Continued)

Species

Egg–adult

Temperature (°C)

Reference no.

D. falleni D. flavomontana D. freckles D. fulvimacula D. fulvimaculoides D. funebris D. funebris D. gaucha D. greeni D. guanche D. guarani D. hexastigma D. huaylasi D. hydei D. hydei D. hydei D. immigrans D. immigrans D. jambulina D. kanapiae D. kanekoi D. kikkawai D. koepferae D. kohkoa D. lacertosa D. lacicola D. lacteicornis D. leonis D. limensis D. lini D. littoralis D. lucipennis D. lusaltans D. lutescens D. mainlandi D. malerkotliana D. martensis D. mayaguana D. mayri D. mediodiffusa D. mediopictoides D. mediostriata D. melanogaster D. melanopalpa D. mercatorum D. mercatorum

13 23.5 14 15 13 12.5 18 17.5 17.5 16 17 24 20.5 17.5 14 18 11 17 17 21 23.5 15.5 19 11.5 18.5 24 18 16.5 17 15 24.5 13.5 18 13 15.5 15 16.5 16.5 22 15 16 13 13 16.5 15.5 14.5

18 18 18 18 18 25 21 18 18 18 18 18 18 18 24 18 25 21 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18

1 1 1 1 1 10 6 1 1 1 1 1 1 1 5 6 10 6 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 (Continued)

192

How to use living flies Table 7.1.

(Continued)

Species

Egg–adult

Temperature (°C)

Reference no.

D. meridiana D. meridianalis D. mettleri D. micromettleri D. milleri D. mojavensis D. montana D. mulleri D. nannoptera D. narragansett D. nasuta D. nasutoides D. navojoa D. nebulosa D. neocardini D. neocordata D. neohypocausta D. neorepleta D. nigricruria D. nigrodumosa D. nigrodunni D. nikananu D. novamexicana D. orena D. ornatipennis D. orosa D. pachuca D. pallidosa D. pallidosa-like D. palustris D. parabipectinata D. para-like D. paralutea D. paranaensis D. parisiena D. parthenogenetica D. parvula D. paulistorum D. pavani D. pegasa D. peninsularis D. pennae D. persimilis D. persimilis D. phaeopleura D. phalerata D. polychaeta

15 17 18.5 17 21.5 16 24 15.5 17.5 17 11.5 19 16 13.5 15 17.5 13.5 21.5 18 17.5 15.5 13.5 21 15.5 16.5 17 20 18.5 17 14 13 14 15.5 16 17.5 15 16.5 16 21 15.5 16 23 15.5 13 13 12 15.5

18 18 18 18 18 18 18 18 18 18 18 20 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18 25 18 18 18

1 1 1 1 1 1 1 1 1 1 1 11 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 9 1 1 1

Life history variation Table 7.1.

193

(Continued)

Species

Egg–adult

Temperature (°C)

Reference no.

D. polymorpha D. ponera D. procardinoides D. propachuca D. prostipennis D. pseudoananassae D. pseudoobscura D. pseudoobscura D. pseudoobscura D. pulaua D. punjabiensis D. putrida D. quadraria D. repleta D. repletoides D. richardsoni D. robusta D. robusta D. rubida D. rufa D. saltans D. seguyi D. serrata D. simulans D. simulans D. sordidula D. spenceri D. sternopleuralis D. straubae D. subbadia D. subobscura D. subpalustris D. subsaltans D. sucinea D. sulfurigaster D. takahashii D. talamancana D. teissieri D. tolteca D. triauraria D. tropicalis D. tsacasi D. unipunctata D. varians D. venezolana D. virilis

15 13.5 15 20 12.5 14.5 14.5 13 17 13 18.5 14.5 18 16 16 19.5 15 21 14 19 17 13.5 18.5 12.5 8 15.5 17 12 14.5 16.5 23 17.5 16 17.5 11 12.5 16 12.5 12.5 15.5 13.5 19.5 15 15 23 20

18 18 18 18 18 18 18 25 21 18 18 18 18 18 18 18 25 21 18 18 18 18 18 18 25 18 18 18 18 18 20 18 18 18 18 18 18 18 18 18 18 18 18 18 18 18

1 1 1 1 1 1 1 9 6 1 1 1 1 1 1 1 3 6 1 1 1 1 1 1 1 1 1 1 1 1 7 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 (Continued)

194

How to use living flies Table 7.1.

(Continued)

Species

Egg–adult

Temperature (°C)

Reference no.

D. virilis D. vulcana D. willistoni D. yakuba S. lebanonensis S. lebanonensis S. pattersoni S. stonei Z. badyi Z. ghesquierei Z. sepsoides

18 16.5 15.5 12.5 19 18.5 19.5 18 13 15.5 16

24 18 18 18 18 18 18 18 18 18 18

2 1 1 1 1 1 1 1 1 1 1

1
Tucson Stock Center; 2
Alexander, 1976; 3
Carson, 1961; 4
Fogleman and Wallace, 1980; 5
Hess, 1976; 6
Kambysellis, 1968; 7
Maynard Smith and Maynard Smith, 1954; 8
Moriwaki and Tobari, 1973; 9
Poulson, 1934; 10
Royes and Robertson, 1964; 11
Zacharias, 1986.

Table 7.2. Development times of six species of Hawaiian Drosophila; ‘jump’ refers to the time at which larvae move to the sand to pupate Genus

Species

Strain

Egg–jump time

Jump– eclosure time

Egg– eclosure

Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila

gymnobasis crucigera soonae biseriata mimica grimshawi eurypeza

.0 .3 .0 .1 .0 .0 .0

12 14 10 12 10 14 13

14 14 13 14 15 15 16

26 28 23 26 25 26 29

Hawaiian Drosophila (Table 7.2) are reared differently from the species in Table 7.1 because pupation must be allowed to take place in sand jars (Chapter 8). For seven species of Hawaiian Drosophila, we partitioned development time into early development, which ends when larvae “jump” or “skip” from the culture medium into the sand, and the second half of their pre-adult development, which takes place in the sand and consists largely of the pupal stage. Development times for the Hawaiian species are between three and four weeks.

Age at sexual maturity Flies of most Drosophila species are rarely sexually mature on the day they eclose. In some species reproductive maturity is attained within a few days of emergence, while others may require weeks. Furthermore, males and females may mature at markedly different times. Age at sexual maturity is available for a number of Drosophila species (Table 7.3). In the majority of cases, the age reported is the age at which 80 percent of

Life history variation

195 Table 7.3.

Ages at reproductive maturity

Species

Female maturity in days (80)%

Male maturity in days (80%)

Reference no.

D. acanthoptera D. affinis D. americana D. anceps D. arizonae D. bifurca D. borealis D. buskii D. eohydei D. eremophila D. ezoana D. flavomontana D. guttifera D. hydei D. kanekoi D. lacicola D. littoralis D. lummei D. mayaguana D. melanica D. melanogaster D. mettleri D. micromelanica D. micromettleri D. mojavensis D. montana D. nannoptera D. navojoa D. nigrospiracula D. novamexicana D. pachea D. parisiena D. persimilis D. pseudoobscura D. recens D. robusta D. simulans D. subpalustris D. silvestris D. virilis D. wassermani

6 4 4 5 3 7 4 2 3 3 7 5 5 3 4 3 4 6 8 1 4 2 4 3 3 4 4 4 4 4 3 5 4 3 5 6 3 4 21 3 4

6 2 6 3 6 17 9 0 7 0 14 9 5 10 19 3 10 7 6 0 2 2 4 1 7 8 8 6 6 6 14 6 0 1 4 10 1 3 6 6 12

6 8 7 7 2 7 7 7 7 7 7 7 7 3, 5 7 7 7 7 7 7 7 2 7 7 2 7 6 7 2 7 4 7 8 8 7 7 7 7 1 7 7

1
Boake and Adkins, 1994; 2
Markow, 1982; 3
Markow, 1985; 4
Pitnick, 1993; 5
Pitnick and Markow, 1994a; 6
Pitnick and Markow, 1994b; 7
Pitnick et al., 1995; 8
Snook, 1995.

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How to use living flies

flies are observed to mate. For more than half of the species examined (Markow, 1996, 2002), if the sexes differ in the age at reproductive maturity it is the females that mature first. A notable exception is the Hawaiian species. In those species for which data are available, such as D. silvestris and D. grimshawi (Spieth, 1964; Boake and Adkins, 1994), females require nearly a month to become mature while conspecific males are ready less than a week after emergence. While these differences appear to be hard-wired in each species, environmental factors such as temperature and nutrition can influence maturation rates. Sexual maturation is typically faster in flies receiving adequate adult nutrition, and can be delayed by cold temperatures (Markow, unpublished).

Fecundity and fertility differences Flies of different Drosophila species also differ markedly in their reproductive potential, or number of progeny they can produce. While it is usually the case that female animals produce far fewer gametes than do males, Drosophila have provided an exception. In most species, the number of offspring is limited by the number of eggs a female produces. In several Drosophila species, however, male gamete production can limit potential productivity. For example, in species such as D. nannoptera and D. hydei (Pitnick and Markow 1994a, 1994b), males pass only 81 and 126 sperm per copulation, as opposed to 4600 in D. melanogaster (Gilbert, 1981). Unfortunately, comparative life tables for Drosophila species do not exist. Reproductive potential can, however, be inferred from characters such as ovariole number in females, which is associated with the number of eggs that can be produced. Ovariole number has been measured in only a subset of the species in the Tucson Stock Center (Table 7.4). Even from this sample, it is clear that potential egg production can vary several-fold among species. Ovariole number may be influenced by environmental factors (David, 1970; Bouletreau-Merle et al., 1982; Wayne and Mackay, 1998), but the large species differences are robust and will have major influences on maintenance and experimental activities as discussed in Chapters 8 and 10. Furthermore, oogenesis in some species, such as D. mulleri, is synchronous, with all ovarioles in a given female having oocytes of the same stage (Kambysellis, 1968), with the result that eggs are laid in clutches on given days, with days of no oviposition intervening. In D. melanogaster, on the other hand, oogenesis is asynchronous among ovarioles (Kambysellis, 1968), with a continual supply of mature eggs.

Life history variation and ecology Those cases in which phylogenic studies of the above life history characters were conducted (Pitnick et al., 1995, 1997) revealed that much of the observed interspecific variability is likely to be explained by ecological rather than long-term evolutionary relationships. While a good deal is known about Drosophila ecology, especially with respect to host associations, few studies have yet addressed the features of Drosophila resource ecology that may have driven the differences among species. The relationship between ovariole number (which exhibits incredible interspecific variation) and breeding

Life history variation

197 Table 7.4.

Ovariole number in selected species of Drosophila

Species

Ovariole no.

Reference no.

D. affinis D. aldrichi D. buskii D. cardini D. crucigera D. funebris D. gibberosa D. hydei D. immigrans D. melanogaster D. mettleri D. mimica D. mojavensis D. montana D. mulleri D. nigrospiracula D. pachea D. paramelanica D. persimilis D. pseudoobscura D. repleta D. robusta D. sechellia D. simulans D. spenceri D. virilis

27 36 44 40 40 42 36 46 64 43 36 24 26 38 40 40 28 24 36 34 36 50 16 35 49 34

7 3 3 3 4 3 3 3 3 6 2 4 2 3 3 2 5 3 7 7 3 3 1 1 2 3

1
Coyne et al., 1991; 2
Heed and Mangan, 1986; 3
Kambysellis, 1968; 4
Kambysellis and Heed, 1971; 5
Markow, 1996; 6
Robertson, 1957; 7
Snook and Markow, 2001.

site has been examined in a number of groups. Factors such as host specialization and predictability have been implicated in evolution of reproductive potential in Hawaiian Drosophila (Kambysellis and Heed, 1971; Montague et al., 1981), flower breeding Drosophila (Starmer et al., 1998, 2000), cactophilic species (Mangan, 1982; Heed and Mangan, 1986), and D. sechellia (Jones, 2004). Other studies have addressed the relationship between characters such as body size, sexual maturity, and gamete production (Pitnick and Markow, 1994a, 1994b; Pitnick et al., 1995; Pitnick, 1996), or the relationship between reproductive investment and sexual selection (Markow, 2002), but the ecological drivers of these relationships have not been examined. We hope that the wealth of data on variable life history traits among Drosophila will inspire additional research not only to test hypotheses regarding the forces responsible for their evolution, but also to understand their role in shaping the future evolutionary potential of these diverse species.

198

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References Alexander, M. L. (1976). The genetics of Drosophila virilis. In: The Genetics of Drosophila virilis (M. Ashburner and E. Novitski, eds), Vol. 1c, pp. 1265–1427. Academic Press, London. Boake, C. R. B. and Adkins, E. (1994). Timing of male physiological and behavioral maturation of Drosophila silvestris (Diptera: Drosophilidae). J. Insect Behav. 7, 577–583. Borash, D. J., Teotonio, H., Rose, M. R. and Mueller, L. D. (2000). Density-dependent natural selection in Drosophila: correlations between feeding rate, development time and viability. J. Evol. Biol. 13(2), 181–187. Bouletreau-Merle, J., Allemand, R., Cohet, Y. and David, J. R. (1982). Reproductive strategy in Drosophila melanogaster: significance of a genetic divergence between temperate and tropical populations. Oecologia 53(3), 323–329. Carson, H. L. (1961). Relative fitness of genetically open and closed experimental populations of Drosophila robusta. Genetics 46, 553–567. Coyne, J. A., Rux, J. and David, J. R. (1991). Genetics of morphological differences and hybrid sterility between Drosophia sechellia and its relatives. Genetic Res. 57, 113–122. David, J. R. (1970). Le nombre d’ovarioles chez la Drosophila: relation avec la fecondité et valeur adaptive. Arch. Zool. Exp. Genet. 111, 357–370. Fogleman, J. C. and Wallace, B. (1980). Temperature dependent development and competitive ability in three species of the Drosophila affinis subgroup. Am. Midland Naturalist 104, 341–351. Gilbert, D. G. (1981). Ejaculate esterase-6 and initial sperm use by female Drosophila melanogaster. J. Insect Physiol 27, 641–650. Heed, W. B. and Mangan, R. L. (1986). Community ecology of the Sonoran Desert Drosophila. In: The Genetics of Drosophila virilis (M. Ashburner and E. Novitski, eds), Vol. 3e, pp. 311–345. Academic Press, London. Hess, O. (1976). Genetics of Drosophila hydei Sturtevant. In: The Genetics of Drosophila virilis (M. Ashburner and E. Novitski, eds), Vol. 1c, pp. 1343–1363. Academic Press, London. Jones, C. D. (2004). Genetics of egg production in Drosophila sechellia. Heredity 92(3), 235–241. Kambysellis, M. P. (1968). Comparative studies of oogenesis and egg morphology among species of the genus Drosophila. Univ. Texas Publ. Genetics 4, 71–92. Kambysellis, M. P. and Heed, W. B. (1971). Studies of oogenesis in natural populations of Drosophilidae. Am. Naturalist 105, 31–49. Mangan, R. L. (1982). Adaptations to competition in cactus breeding Drosophila. In: Ecological Genetics and Evolution: The Cactus–Yeast–Drosophila Model System (J. S. F. Barker and W. T. Starmer, eds), pp. 257–272. Academic Press, New York. Markow, T. A. (1982). Mating systems of cactiphilic Drosophila. In: Ecological Genetics and Evolution: The Cactus–Yeast–Drosophila Model (J. S. F. Barker and W. T. Starmer, eds), pp. 273–287. Academic Press, New York. Markow, T. A. (1985). A comparative investigation of the mating system of Drosophila hydei. Animal Behav. 33, 775–781. Markow, T. A. (1996). Evolution of Drosophila mating systems. Evol. Biol. 29, 73–106. Markow, T. A. (2002). Female remating, operational sex ratio, and the arena of sexual selection in Drosophila. Evolution 59, 1725–1734. Maynard Smith, J. and Maynard Smith, S. (1954). Genetics and cytology of Drosophila subobscura. J. Genetics 52, 152–164. Montague, J. R., Mangan, R. L. and Starmer, W. T. (1981). Reproductive allocation in the Hawaiian Drosophilidae: egg size and number. Am. Naturalist 118, 865–871. Moriwaki, D. and Tobari, Y. N. (1973). Spontaneous male crossing-over of frequent occurrence in Drosophila ananassae from Southeast Asian populations. Japan. J. Genetics 48, 167–173.

Life history variation

199 Pitnick, S. (1993). Operational sex ratios and sperm limitation in populations of Drosophila pachea. Behav. Ecol. Sociobiol. 33, 383–391. Pitnick, S. (1996). Investment in testes and the cost of making long sperm in Drosophila. Am. Naturalist 148, 57–80. Pitnick, S. and Markow, T. A. (1994a). Large-male advantages associated with costs of sperm reduction in Drosophila hydei, a species with giant sperm. Proc. Natl. Acad. Sci. 91, 9277–9281. Pitnick, S. and Markow, T. A. (1994b). Male gametic strategies: sperm size, testes size, and the allocation of ejaculate among successive mates by the sperm-limited fly Drosophila pachea and its relatives. Am. Naturalist 143, 785–819. Pitnick, S., Markow, T. A. and Spicer, G. S. (1995). Delayed male maturity is a cost of producing large sperm in Drosophila. Proc. Natl. Acad. Sci. 92, 10614–10618. Pitnick, S., Spicer, G. and Markow, T. A. (1997). A phylogenetic examination of female incorporation of ejaculate in Drosophila. Evolution 51, 833–845. Poulson, D. F. (1934). Times of development of the two races of Drosophila pseudoobscura. J. Exp. Zool. 68, 237–245. Robertson, F. W. (1957). Studies in quantitative inheritance. J. Genetics 55, 410–427. Royes, W. V. and Robertson, F. W. (1964). The nutritional requirements and growth relations of different species of Drosophila. J. Exp. Zool. 156, 105–135. Santos, M., Fowler, K. and Partridge, L. (1994). Gene–environment interaction for body size and larval density in Drosophila melanogaster: an investigation of effects on development time, thorax length and adult sex ratio. Heredity 72(5), 515–524. Snook, R. R. (1995). The evolution of sperm polymorphism in the Drosophila obscura group. PhD thesis, Arizona State University. Snook, R. R. and Markow, T. A. (2001). Mating system evolution in sperm-heteromorphic Drosophila. J. Insect Physiol. 47, 957–964. Spieth, H. T. (1964). Studies of the mating behavior of endemic Hawaiian Drosophila. Am. Zool. 4, 406. Starmer, W. T., Polak, M., Wolf, L. L. and Barker, J. S. F. (1998). Reproductive characteristics of the flower breeding Drosophila hibisci Bock (Drosophilidae) in eastern Australia: genetic and environmental determinants of ovariole number. Evolution 52, 806–815. Starmer, W. T., Polak, M., Wolf, L. L. and Barker, J. S. (2000). Reproductive characteristics of the flower-breeding Drosophila hibisci Bock (Drosophilidae) in eastern Australia: withinpopulation genetic determinants of ovariole number. Heredity 84, 90–96. Wayne, M. L. and Mackay, T. F. (1998). Quantitative genetics of ovariole number in Drosophila melanogaster. Genetics 148, 201–210. Zacharias, H. (1986). Tissue-specific schedule of selective replication in Drosophila nasutoides. Roux Arch. Devel. Biol. 195, 378–388.

CHAPTER 8

How to use ecological and life history information to rear flies Contents

• • • • • • •

The basics of starting and caring for cultures Life history and ecological considerations in culturing flies Setting up cultures to collect healthy virgin flies Collecting virgin flies • Collection schedule and protocol for species with rapidly maturing adults • Culture and collection schedule for flies with older ages at sexual maturity How to anesthetize flies Rearing Hawaiian Drosophila References

Regardless of the specific culture medium that is used (recipes are in Chapter 9), some husbandry practices will apply to most species. Here we describe, in the first section of this chapter, basic practices for routine maintenance and rearing for experiments. Subsequent sections of this chapter will provide suggestions for ways in which the life history and ecological differences among species should be taken into consideration for successful culturing. In addition, flies are most commonly reared for two different purposes; either simple maintenance of stocks, or setting up cultures for specific experiments. Depending upon the purpose, investigators may want to emphasize different practices when creating and caring for cultures. Thus another section of this chapter will suggest practices for collecting large numbers of virgin flies. Finally, in certain taxa, especially Hawaiian species, larvae pupate in sand or soil, and the final section of this chapter will describe rearing methods for these flies.

The basics of starting and caring for cultures Many laboratories that grow Drosophila melanogaster utilize disposable plastic bottles or vials. We do not recommend the use of plastic containers for two reasons: first, as the material is usually not as clear as glass, it is difficult to monitor the status of the culture; and second, cultures tend to dry out more rapidly in plastic containers. Suppliers of glass bottles and vials are listed in Chapter 12.

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A good practice is to set up culture bottles of any given stock, in sets of two or more, at the same time. Always, in addition to the name and collection number of the species and the initials of the individual starting the culture, write the date the culture was initiated on the bottle or vial. This will indicate when to expect adults of the next generation to emerge and when to plan to set up the next set of cultures. Several techniques should be employed to maximize culture success. First, although most food recipes include some microbial inhibitors, and unused food typically is refrigerated, it is still preferable to start cultures with food that is less than a week old. Even fairly fresh culture medium should be visually inspected before use. If food has begun to separate from the side of the container, flies can become stuck in the crack and die before they have a chance to reproduce. Second, for many species, cultures get off to a better start if the medium surface is rough rather than smooth. Females of many species prefer to oviposit on a rough surface, and first instar larvae find it easier to burrow and feed when the food surface is grooved. Scratching the surface of the food with a clean spatula or other long, pointed device usually will be sufficient. Third, inserting a folded tissue into the food (Figure 8.1) provides perching sites for adults as well as, later, pupation sites for late third instar larvae. Tissues will also absorb the excess moisture that can cause flies to get stuck. When flies (either those of the parental generation or emerging adults) are to be removed from the bottles, do not invert the bottles completely and pound them to shake out the flies (Figure 8.2). Rather, holding bottles slightly on one side will delay the loosening and crashing down of food – a situation that usually kills flies and greatly reduces the numbers of flies ultimately harvested. The start dates written on the cultures will indicate when to expect the next generation of adults. Egg to adult development times (Table 7.1), adjusted for temperature if rearing temperatures are different from 68°F (20°C), should be used as a guide. Thus, for D. affinis, at 68°F, the next generation would be expected to begin emerging roughly 15 days after the culture was started, while for D. novamexicana emergence would begin about 21 days after the culture was set up. At room temperature, development times will be from two to five days shorter for most species. Depending upon whether the cultures were established for routine stock maintenance or for the collection of virgins for an experiment, emerging flies will be handled differently as described in subsequent sections of this chapter.

Life history and ecological considerations in culturing flies Cultures should be initiated with several things in mind, including the type of food recipe and additional treatment of food (Table 9.1), age at sexual maturity (Table 7.3), reproductive potential (Table 7.4), and the length of time parental flies are left in the culture bottles. Some species will do equally well on more than one food type, while others can only be grown on one type of culture medium. Species that are widely distributed in nature and are ecological generalists, such as D. melanogaster, D. simulans, D. immigrans, D. buskii, and D. hydei (Patterson and Stone, 1952), tend to be successful on a wider

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(a)

(b)

Figure 8.1. The (a) correct and (b) incorrect ways to fold tissues in culture bottles.

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(a)

Figure 8.2. The (a) correct and (b, overleaf) incorrect ways to hold a bottle when removing flies.

range of food types compared to endemic species (such as the cactophilic D. pachea) or specialist species (such as the mycophagous species of the quinaria, testacea, and tripunctata groups). When starting to culture a new species, refer to the food types and special treatments listed in Table 9.1. If the species being cultured is not one that is shown on the list, find the most closely related species (see Chapter 1) that does appear and try that food type. Sexually mature adults should always be used. If cultures are started with sexually immature flies, the food will become dry, possibly growing harder or less effective in retarding microbial growth before females are ready to oviposit, and ultimately the culture may fail or be unhealthy. Refer to Table 7.3 to determine the best age for using flies to start the cultures. For species like D. melanogaster, in which flies of both sexes mature early, cultures can be maintained simply by transferring emerged adults, a few days after eclosion, from the previous culture bottle to the new one (Figure 8.3, top). On the other hand, if the species being cultured is one in which adults of one of the sexes

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(b)

Figure 8.2. (continued )

require a week or longer to become sexually mature, such as D. bifurca, D. pachea, or D. kanekoi, newly emerged flies should be aged in food vials, in groups of about twenty per vial, prior to starting the new culture (Figure 8.3, bottom). The vials should be changed every three to four days to maintain the adults in good condition while they are maturing (Figure 8.3). Flies can be aged in mixed-sex groups if they will be used to start new cultures. For most species, excepting the mycophagous and Hawaiian flies, a few grains of live yeast should be added to the surface of the food in the aging vials. Each bottle should be founded with enough flies to start the culture, but not so many as to create crowding. For most species, approximately twenty pairs is a good number. If stocks are reared in vials rather than bottles, fewer flies should be used. Maintaining stocks with too few flies will cause a loss of genetic variation. One way to maintain variation is by keeping more than one culture of a specific strain or species and then mixing flies (of the same stock, of course) from the different culture bottles when starting new generations. Initiating cultures with too many adults leads to crowding, which creates other problems, from delays in adult emergence to huge variations in adult size, and sloppy, runny cultures from which collecting is difficult to impossible. Uncrowded conditions will also yield robust third instar larvae in the event that salivary chromosome squashes are desired. Effects of crowding may be reduced by utilizing knowledge of the life history differences described in Chapter 7. Species differ several-fold in their reproductive output,

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Newly emerged adults

Day 1 – culture started

F1 adults emerge

Day 1 – culture started

Newly emerged adults

Day 1 – culture started

Day X – F1 adults emerge

Transfer adults every 3–4 days to fresh, yeasted vials while maturing

Day 1 – culture started

Figure 8.3. Serial transfer of adults while becoming sexually mature before starting cultures. Top indicates procedure for species with short times until sexual maturity, bottom for species in which sexual maturation is delayed.

as can be inferred from the differences in ovariole number shown in Table 7.4. D. melanogaster should not be used as the guide for species of the genus Drosophila. One approach to avoid crowding is to remove the founding adults once sufficient eggs have been laid to ensure a healthy culture. Reproductive potential data can be used to finetune the duration of time that adults are left in cultures. For example, in species with high ovariole numbers, such as D. immigrans, females are likely to lay more eggs and cultures will have more larvae after a shorter period of time than species such as D. sechellia, with lower reproductive potential. In the former case, crowding can be reduced by transferring the mature parental adults out of the cultures more quickly – say after two or three days – than for species that produce fewer eggs. Because the reproductive potentials of all species are not known, investigators may need to discover, on

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their own, the optimal number of days to leave adults in a culture. For routine maintenance, many investigators do not bother to remove parents from cultures or worry about the effects of crowding because it is extra work. If possible, however, attending closely to the cultures will provide consistent and healthy stocks. Several tricks can be used to increase oviposition. Scratching the surface of the food in the culture vial or bottle will often induce females to lay eggs, and will also enable first instar larvae to burrow more easily into the food. For specialist species, addition of their natural food (Chapter 9) to the culture is also useful. For example, for cactophilic species, place a small cube (less than 1 inch square) of autoclaved cactus onto the food surface before introducing the adults. For D. pachea the cactus must be senita (Lophocereus schottii), because of its nutritional dependence on this cactus (Heed and Kircher, 1965), but for other cactophilic species a piece of prickly pear cactus (Opuntia ficus-indica) will serve the purpose. For mycophagous flies, add a medium-sized grocery store mushroom.

Setting up cultures to collect healthy virgin flies If an experiment requires large numbers of virgin flies, several practices should be employed. First, establish a larger number of cultures each generation. Instead of starting cultures in duplicate, as for routine stock maintenance, prepare five or ten cultures. Second, completely clear cultures of parental flies three to five days after they are started. Removal of parents early will result in uncrowded cultures from which it will be easier to collect large numbers of healthy, uniform virgin flies. Use of an aspirator will facilitate removal of parents and improve the ability to collect virgins once they start to emerge. Removing parental flies by the standard technique of inverting the bottle and pounding will start loosening the food from the sides of the bottle. Collecting virgins later from these bottles will probably again involve pounding them, and this will eventually cause the food to crash down, killing or damaging flies that could have been collected. When removing flies with an aspirator, it is helpful temporarily to replace the paper cap with a foam plug that permits the aspirator to be inserted into the container without losing flies (Figure 8.4). Another technique that will maximize the life of the culture and the number of virgins obtained is to introduce a second piece of tissue (as in Figure 8.1) at the time the third instar larvae are beginning to pupate. This practice accomplishes three things: first, it increases the available pupation sites, thereby increasing the number of individuals emerging; second, it keeps the culture from becoming too mushy or “soupy”; and third, the food remains more tightly in place in the bottle and is more resistant to the destructive effects of the repeated upside-down pounding that takes place later, during the virgin-collecting phase. If adults are transferred out of a culture about five days after the culture is started, they can then be used to start another set of bottles by simply transferring them to new food bottles, creating a second “brood” from the same parents (Figure 8.5). Using parents for more than two broods, regardless of whether they are long-lived and reproductive, is not recommended, as potential parental age effects on the characters of interest may

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Figure 8.4. Aspirating flies from vials or bottles.

Day 1 Brood 1

X Days*

Begin collecting virgin flies, continue 4 days

X Days*

Begin collecting virgin flies, continue 4 days

Day 3: Transfer parental flies

Day 4 Brood 2

Day 10: Discard parental flies

Figure 8.5. Starting multiple broods for large-scale virgin collecting.

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come into play (see Chapter 10). For large-scale experiments, it is often desirable to keep a continuous supply of virgins by starting new sets of bottles every three to five days for an extended period of time. It is better to have too many flies than too few!

Collecting virgin flies One day before adults are expected to emerge, bottles should be inspected. Clearly, pupae should be present. In species with light or transparent pupa cases, dark, pharate adults may be visible inside. In many species, however, such as D. virilis, the pupa cases are pigmented and opaque, and the maturity of the pupae inside may not be ascertainable. Presumably, all parental adults were removed sometime earlier, and no older males will be lurking when the first virgin females appear. Because individuals vary in development time, a small number of new adults will appear early. These should be removed with an aspirator and discarded the first day they are detected. The following day, it should be possible to begin collecting and storing useful numbers of virgin flies from the cultures. There are two reasons for removing and discarding these first adults. First, it is always possible that a parental fly or flies went undetected when the bottles were cleared sometime earlier. Second, especially in species where flies reach sexual maturity early, older males may mate with newly emerging females (Markow, 2000), such that female virginity cannot be assumed. Once virgin adults start coming out, they can be collected daily for three to five days from these bottles. Collecting virgins from a given set of bottles for more than three to five days is unwise, because flies emerging later become smaller and more variable in size. Furthermore, because as larvae they have been ingesting medium that has been “worked” by predecessors for many days, they may be physiologically compromised in some way that could influence results of whatever experiment they will be used for. Some of these effects are more specifically addressed in Chapter 10. Because it is easy, with Drosophila, to obtain large numbers of healthy flies from fairly uniform culture conditions, there is no reason to compromise experiments. Two virgin-collecting approaches are common to all species. The first is to use cuticle color as an indicator of age. Teneral flies – that is, flies that have emerged within the last few hours – will be very light and translucent in color. In some species, a dark or greenish material in the digestive tract will be visible through the ventral abdominal wall of teneral individuals. Typical adult pigmentation patterns will not appear until tanning of the cuticle is complete. Thus, if a group of flies is harvested from a bottle in the early morning, flies that have eclosed overnight will exhibit the typical pigmentation patterns characteristic of more mature adults, while those having emerged much more recently will be quite pale and in some cases their wings will still be folded on their backs. If no sexually mature males remained in the vials, these very young flies are almost certainly virgins and can be accepted as such. At the same time, because they are not yet fully pigmented they are sometimes more difficult to separate by sex, and their external genitalia should be carefully examined under the microscope. If culture bottles are “cleared” regularly (every six hours in the case of rapidly maturing species, and every day in the case of more slowly maturing species, see below), all flies,

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Day 5

Day before eclosion expected

First day of eclosion

Remove parents

Check and clear any emerged flies

Collect, clear every 6 hours Early AM: Light flies only Later collect: All flies

Label vials with date, sex, initials

(a)

Allow to mature for 4 days

Figure 8.6. Procedure for culturing and virgin collecting in flies with (a) early and (b, overleaf) delayed sexual maturity.

regardless of how translucent they are, can be assumed to be virgin. Clearing means aspirating out all adults and killing those that appear to be incapacitated or stuck in the food, by pushing them into the food with the blunt end of a brush or a spatula. Once virgins begin emerging, the intervals at which they are collected will vary among species. Collection interval should be determined by the age at which adults become sexually mature (Table 7.3). Basically, two different collection schedules are recommended. In species in which flies, especially males, mature early, cultures should be cleared every six hours (Figure 8.6a). In species in which flies require many days or even weeks to mature, cultures can be cleared and collected daily (Figure 8.6b). Even in species where flies require weeks to become sexually mature, cultures should be cleared daily and virgins separated. Daily collection of virgins allows the ages of virgin flies used in experiments to be carefully controlled.

Collection schedule and protocol for species with rapidly maturing adults The sequence described below is recommended for species such as D. melanogaster and D. simulans, in which adults become sexually mature within a few days of emergence.

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How to use living flies Day 5

Day before eclosion expected

First day of eclosion

Remove parents

Check and clear any emerged flies

Collect, clear every 24 hours

Label vials with date, sex, initials

Allow to mature for 4 days and transfer (b)

Figure 8.6.

(continued )

Many other species will fall into this category (Table 7.3). The sequence assumes that cultures will be kept at room temperature rather than at 68°F. 1. Four days after starting the culture, remove all parents with an aspirator. Parental flies that are stuck in the food should also be removed or killed. Nine days after the particular set of cultures was started, examine the bottles to be sure all parental adults are gone. Because the generation time is about ten days at room temperature, it is possible that some flies of the new generation will come out a day early and they must be removed (with an aspirator) to be sure that all flies are virgins when collecting begins. Fold a new piece of tissue (Figure 8.1) and insert it down into the food in the corner or side of the bottle, between the culture medium and the glass. 2. The next day, virgins should be emerging and be ready to be collected. With D. melanogaster and other early maturing species, virgin males and females need to be collected and separated every six hours. This means that on day 10, collection from the bottles should be carried out first thing in the morning, and then again six hours later. Be absolutely certain that no flies remain in the bottles at each collection time. If at all possible, in the late evening of that day the bottles should again be cleared (with an aspirator), so that the flies found early in the morning on the second day of

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collecting will be virgins. If the bottles are not cleared during the evening, in the first collections made the following morning the only flies accepted as virgins should be the very light ones; any full pigmented flies should be discarded. These may no longer be virgin, and it is not worth the risk. All flies can be taken during the second collection on any day, provided the bottles were completely cleared during the morning collection. One means of ensuring that a culture is truly “cleared” of flies is to use an aspirator to poke about in the tissue a few minutes after the collection has been made; hiding flies will then either wander from their crevices or be discovered in their hiding places and removed. 3. Store females and males separately, with ten to fifteen flies per vial, in cornmeal vials with a few grains of yeast. Cornmeal vials are preferable for storage, as they are less moist and flies don’t get stuck. For mycophagous flies, do not use live yeast. For Hawaiian flies, follow the directions in the last section of this chapter. Before brushing the anesthetized flies into their storage vials, check them again to be sure they are all the correct sex. When anesthetized flies are first introduced into the storage vials, be sure no flies rest on the food or they could become stuck. Rest all vials on their sides until the flies have awakened. A good practice, after the flies have regained consciousness in their storage vials, is to hold each vial under the microscope and again inspect the flies to make sure they are all the same sex. Storing virgin flies at low densities may seem to be a needless use of extra storage vials, but it serves two important purposes: first, if there is contamination with a fly or flies of the wrong sex, fewer vials will need to be discarded; and second (as will be discussed in greater detail in Chapter 10), density during storage can affect behavior. Precautions taken early can prevent an aborted experiment. 4. Eighty per cent of the flies of species such as D. melanogaster and D. simulans will be sexually mature at three or four days old when kept at 75°F (24°C). While there is variability among individuals with respect to when sexual maturity is reached, and some flies may mature early, it may be best to wait until all flies are mature. In a species in which the majority of flies are mature at four days, waiting until the fifth day to perform crosses might be most appropriate if the experiment calls for observing courtship and mating. If flies are to be mated solely to set up crosses to obtain offspring, the crosses can be made and cultures started on the day the majority of flies become mature.

Culture and collection schedule for flies with older ages at sexual maturity The sequence described here (Figure 8.6b) should be employed when setting up experiments with species such as D. bifurca or D. kanekoi, in which males require about two and a half weeks before they become sexually mature. 1. Remove parental flies with an aspirator about five days after starting the cultures. Flies that appear to be “stuck” in the medium should either be removed or killed. 2. Start checking for the new generation at about two days before adults are expected to emerge (based upon Tables 7.1 and 7.2, or preliminary data). Once virgins begin to appear, they only need to be collected once a day. Since these flies take considerably

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longer to become reproductively mature, they do not need to be cleared a second time during the day or collected every six hours. 3. As with the shorter maturation-time flies described above, brush male and female virgins into cornmeal vials (ten to fifteen flies per vial). Check them a second time to be sure all flies are the same sex. Rest vials on their sides, so the flies are off the food surface until they are awake. 4. In many slowly maturing species males and females differ dramatically in the onset of sexual maturity, and they will therefore be ready for use in experiments at different times. Flies requiring longer times to mature (most often the males) should be changed to fresh vials every three to four days to ensure that they are clean and healthy. While virgin flies may appear to be perfectly fine after storage for four days in the same vials, the absence of larvae in the storage vials will allow some microbial growth, reducing the health of the stored flies. Small decrements in health can produce detectable influences on experiments, especially those involving behavioral assays (see Chapter 10). In species in which females mature much earlier than males, there is no reason not to use the mature females in crosses or tests with sexually mature males from an earlier brood.

How to anesthetize flies Identification of wild-caught flies and routine culturing and collecting require that they be anesthetized. The two primary forms of anesthetizing flies are with ether or CO2. Both will work for most species, but each has its own advantages and disadvantages, and species may respond differently to them. Using cold treatments (ice) to knock out flies will not be discussed here, because flies can become stuck in any condensation on the slide or chamber. Etherization of flies requires that they be removed from their culture or collection container, either with an aspirator or by shaking directly into the etherizer, prior to being knocked out. There are several disadvantages of ether. One is that flies can easily become over-etherized, as evidenced by their wings sticking up at a 90° angle from their bodies. Flies over-etherized to this extent may already be dead. Even when the wings are not sticking up, flies may still be over-etherized and either may not wake up after they are transferred to new containers, or, while they may appear healthy, may be compromised in some way that alters behavioral (Gilbert, 1981; Seiger and Kink, 1993) or physiological (Krebs et al., 1995) processes that reduce fertility, fecundity or longevity. Another disadvantage to ether is the temporary nature of the anesthesia. Flies may wake up before they are sorted, leading to the loss of useful individuals. While species differ in the degree to which they are influenced by ether (Seiger and Kink, 1993), genetic variation for ether sensitivity/resistance also exists within species (Gamo et al., 1998). Most problems with ether anesthetization can be avoided by knocking out smaller groups of flies at a time, and by watching them as soon as they are in the etherizer. At the very instant that flies stop moving in the etherizer, they should be removed. Because this length of time will be different for different species, testing a few individuals of an unfamiliar species before full-scale work begins is recommended. Finally, not only is ether highly flammable, but also exposure to ether fumes causes some investigators to

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get headaches. Working in a well-ventilated area away from flames will reduce the latter problems. Carbon dioxide (CO2), while it avoids certain problems associated with ether, has its own disadvantages. Flies can be anesthetized before they are transferred out of their containers and viewed under the microscope in a CO2 chamber. The chamber allows flies to be immobilized longer, so that larger numbers can be sorted at a time. However, some species are very sensitive to the effects of CO2, and may either die simply from exposure or exhibit subsequent alterations in their behavior (Gilbert, 1985; Seiger and Kink, 1993) or physiology (Krebs et al., 1995). In addition to species differences, withinspecies variability in CO2 sensitivity has been known for some time and is associated with the presence of the sigma virus (L’Heritier, 1951; Fleuriet, 1996). Finally, flies immobilized with CO2 sometimes undergo some degree of temporary physical distortion. In some cases, their wings are slightly folded over their abdomens, potentially obscuring coloration patterns. In other cases, the male genitalia are more difficult to see. These two problems can complicate examination of features used in keying out species. A wide range of excellent laboratory and field portable CO2 devices are available through Genesee Scientific (www.geneseesci.com).

Rearing Hawaiian Drosophila In addition to the requirement for Wheeler–Clayton medium, Hawaiian flies prefer cool temperatures (68°F) and relative humidity of at least 35% (Spieth, 1972). In nature they live in cool, humid forests, and attempts to mimic their natural habitat are critical to successful rearing. While in nature they experience higher relative humidity, keeping them at higher humidity in the lab often leads to the growth of mold. Live yeast should not be added to the surface of the medium. Several life history features of Hawaiian Drosophila require that special procedures be employed for these flies. First, sexual maturity, especially in females, can be delayed by several weeks, and therefore adults should be transferred to new Wheeler–Clayton holding vials every four to seven days, until larvae are observed. While not essential for culturing these species, females can be encouraged to oviposit by placing a piece of tissue moistened with the juice from Clermontia leaves into the medium (see Chapter 9). For routine culturing, males and females can be aged together. Place a piece of tissue moistened with water into the vial to increase the humidity. Vials with adults should always be placed on their sides (Figure 8.7). Up until the time larvae are observed in a set of vials, empty holding vials can be discarded. Once larvae are present, vials should be kept and observed, and adults should continue to be transferred to new vials for several more weeks. Second, in nature mature larvae “jump” or “skip” in order to pupate in the soil near their larvae resource. In order to accommodate this behavior in the lab, vials with third instar larvae should be placed (without plugs) in sterilized gallon jars, one-fourth full of loose, moist sand, which will serve as pupation chambers (Figure 8.7). Approximately five vials can be placed in the jar. The gallon jars are sealed with muslin. Sand should be moistened and mixed with de-ionized water before vials are placed inside. Vials should be placed on their sides in the sand, with the vial mouths slanted slightly upward. Prior to having their plugs removed and being placed in the jars, the vials should be wiped

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Figure 8.7. Sand jars and vials for rearing Hawaiian Drosophila.

with 70% EtOH. Larvae will jump out of the vials and burrow into the sand to pupate. Adults will eclose two to three weeks later, depending upon the species. The adults should be aspirated out of the jars and, without anesthetization, transferred to new Wheeler– Clayton vials. Information on suppliers of appropriate sand is given in Chapter 12.

References Fleuriet, A. (1996). Polymorphism of the Drosophila-sigma virus system. J Evol. Biol. 9, 471–484. Gamo, S., Dodo, K., Matakatsu, H. and Tanaka, Y. (1998). Molecular genetic analysis of Drosophila ether sensitive mutants. Toxicol. Lett. 100–101, 329–337. Gilbert, D. G. (1981). Effects of CO2 vs. ether on two mating behavior components of Drosophila melanogaster. Drosophila Information Service 56, 45–46. Heed, W. B. and Kircher, H. W. (1965). Unique sterol in the ecology and nutrition of Drosophila pachea. Science 149, 758–761. Krebs, R. A., Loeschcke, V. and Hakansson, C. (1995). Resistance and acclimation to thermal stress in adult Drosophila buzzatii following collection under ether and CO2. Drosophila Information Service 76, 145–148. L’Heritier, P. (1951). The CO2 sensitivity problem in Drosophila. Cold Spring Harbor Symp. Quant. Biol. 16, 99–112. Markow, T. A. (2000). Forced matings of Drosophila females in natural populations. Am. Naturalist 156, 100–103. Patterson, J. T. and Stone, W. S. (1952). Evolution in the Genus Drosophila. Macmillan, New York. Seiger, M. B. and Kink, J. F. (1993). The effect of anesthesia on the photoresponses of four sympatric Drosophila. Behav. Genetics 23, 99–104. Spieth, H. T. (1972). Rearing techniques for the Hawaiian species Drosophila grimshawi and Drosophila crucigera. Drosophila Information Service 48, 155–157.

CHAPTER 9

Dietary considerations Contents

• •

• •

Adult and larval Drosophila diets Recipes • Standard cornmeal medium (makes approximately 28 cases of vials) • Standard banana–opuntia (makes approximately six cases of vials, or three cases of half-pint bottles) • Standard Wheeler–Clayton (makes approximately six cases of wide-diameter plastic vials) • Saguaro–potato (makes approximately 24 half-pint bottles) Special treatments and requirements References

Adult and larval Drosophila diets Of more than 1000 species of Drosophila, only about 300 are cultured successfully in the laboratory. Drosophila resource use in nature involves dietary adaptations ranging from dependency upon specific chemicals (Heed and Kircher, 1965) or the ability to detoxify specific substances (Jaenike et al., 1983) through the requirement for a diet either high or low in essential elements such as nitrogen and phosphorus (Markow et al., 1999). Establishing the correct nutrient composition is but one dimension of the rearing conditions in which a species can be cultured. The ecological niches of some species, such as D. carbonaria, which breeds in the flux of mesquite trees of the genus Prosopis (Patterson and Wheeler, 1942), D. carcinophila, a commensal with land crabs (Carson, 1967), and many flower breeders (Brncic, 1983), have seasonal and structural requirements in addition to nutritional specialization that make laboratory rearing exceedingly difficult or impossible. Creating the appropriate conditions for adult feeding, mating, and oviposition, as well as providing suitable larval habitats, requires an effort beyond that which most investigators are willing or able to invest. Considerable variation in the ease with which flies are cultured exists even among those species that are routinely maintained in laboratories or the stock centers. Some species can be grown easily on a range of recipes, while others require a specific recipe or nutritional treatment in order to be grown successfully (Table 9.1). In this chapter we provide recipes for the food types used at the Tucson Stock Center for the species maintained there. We also provide information for rearing a number of specialized species. The

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Recipes used by the Tucson Stock Center for rearing species in the

Genus

Species

Culture media

Chymomyza Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila

procnemis acanthoptera acutilabella affinis albomicans aldrichi algonquin ambigua americana ananassae anceps antillea aracatacas arawakana arizonae auraria austrosaltans azteca baimaii barbarae belladunni biarmipes biauraria bicornuta bifasciata bifurca bipectinata birchii biseriata borborema bromeliae busckii buzzatii camargoi canadiana canalinea canapalpa capricorni cardini cardinoides caribiana crocina crucigera dacunhai deflecta diplacantha

BO BO BO BO C BO, WC C BO C C BO, WC BO WC BO BO C BO BO C C BO C C C BO WC C C WC WC BO BO BO BO C BO BO C BO BO BO WC WC BO BO C

Dietary considerations

217 Table 9.1.

(Continued)

Genus

Species

Culture media

Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila

dunni elegans ellisoni emarginata eohydei equinoxialis ercepeae erecta eremophila eugracilis euronotus eurypeza ezoana falleni ficusphila flavomontana formosana fraburu freckles fulvimacula fulvimaculoides fumipennis funebris gaucha gibberosa greeni grimshawi griseolineata guanche guarani guttifera gymnobasis hamatofila hexastigma huaylasi hydei hypocausta immigrans iri jambulina kanapiae kanekoi kepulauana kikkawai koepferae

BO BP BO BO BO, C C C C BO C C WC C WC, M C C WC, C BO WC BO BO C C C WC C WC WC BO BO BO, WC, M WC WC BO BO C, BO WC C BO C C C C, BO C BO (Continued)

218

How to use living flies Table 9.1.

(Continued)

Genus

Species

Culture media

Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila

kohkoa lacertosa lacicola lacteicornis leonis limensis lineosa lini littoralis longicornis lucipennis lummei lusaltans lutescens macrospina mainlandi malerkotliana martensis mauritiana mayaguana mayri mediodiffusa mediopictoides mediostriata melanica melanogaster melanopalpa mercatorum meridiana meridianalis mettleri micromelanica micromettleri milleri mimica miranda mojavensis montana montana mulleri nannoptera narragansett nasuta navojoa nebulosa neocardini

C BO C C BO C C C C BO C C BO C BO BO C BO C, BO BO C WC WC WC BO C, BO BO BO C, BO BO BO, SP BO BO BO WC BO BO C C C, BO BO BO C BO C BO

Dietary considerations

219 Table 9.1

(Continued)

Genus

Species

Culture media

Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila

neocordata neohypocausta neorepleta nigricruria nigrodumosa nigrodunni nikananu novamexicana ornatipennis orosa pachea pachuca pallidipennis pallidosa pallidosa-like palustris parabipectinata para-like paralutea paramelanica paranaensis pararubida parisiena parthenogenetica parvula paulistorum pavani pegasa peninsularis pennae persimilis phaeopleura phalerata polychaeta polymorpha ponera procardinoides propachuca prosaltans prostipennis pseudoananassae pseudoobscura pseudotakahashii pulaua punjabiensis

BO BO C BO BO BO C C BO C BO
Senita BO BO C C BO, WC C C C BO C BO BO BO C C BO BO C C BO C WC, M C BO, WC WC BO BO BO C C BO C C C (Continued)

220

How to use living flies Table 9.1.

(Continued)

Genus

Species

Culture media

Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Drosophila Hirtodrosophila

putrida quadraria repleta repletoides richardsoni ritae robusta rubida rufa saltans sechellia seguyi serido serrata signata simulans soonae sordidula spenceri stalkeri sternopleuralis straubae sturtevanti subbadia subpalustris subsaltans sucinea sulfurigaster takahashii talamancana teissieri tolteca triauraria tripunctata tropicalis tsacasi unipunctata varians venezolana virilis vulcana wassermani wheeleri willistoni yakuba duncani

BO, M C BO BO BO BO BO C C BO C C BO C C C, BO WC BO BO BO BO BO BO BO BO BO C C C BO C BO C BO, WC, M C C BO, WC, M C BO C C BO BO C C BO

Dietary considerations

221 Table 9.1.

(Continued)

Genus

Species

Culture media

Scaptomyza Scaptomyza Scaptomyza Scaptomyza Scaptomyza Scaptomyza Scaptomyza Scaptomyza Zaprionus Zaprionus Zaprionus Zaprionus

anomala galloi latifasciaeformis lebanonensis leonensis palmae pattersoni stonei badyi ghesquierei inermis sepsoides

BO BO BO, WC BO WC WC BO BO BO BO BO BO

BO  banana–opuntia; C  cornmeal; WC  Wheeler–Clayton; SP  Saguaro–potato; M  mushroom.

Stock Center maintains a large number of species of the repleta group, and these species perform well on ordinary banana food. Because most of these species are cactophilic in nature, we find that the addition of opuntia powder, as given in the recipe below for banana–Opuntia medium, greatly enhances their performance at the same time as enabling the maintenance of many other, non-cactophilic, species. Therefore, the Stock Center uses this food for all species that might also perform well on plain banana food. Suppliers of ingredients and equipment used in preparing the recipes given below are listed in Chapter 12. Additional recipes for other Drosophila culture media can be found at http://flystocks.bio.indiana.edu/media-recipes.html.

Recipes When filling vials or bottles, regardless of the food recipe, care should be taken to avoid drips on the sides of the containers and to be sure a consistent amount of food is dispensed to each container. The Tucson Stock Center uses the following pouring dimensions:

• • • •

Glass vials: 10 ml/vial  1000 ml (1 l)/case of 100 vials Plastic vials: 10 ml/vial  1000 ml (1 l)/case of 100 vials Glass bottles: one-third cup/bottle
2 l/case of 24 bottles Plastic bottles: one-fourth cup/bottle
1.5 l/case of 24 bottles.

Standard cornmeal medium (makes approximately 28 cases of vials) Ingredients Agar: 270 g Cornmeal: 750 g

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Sugar: 1200 g Yeast: 420 g De-ionized water: 28 l Propionic acid: 106 ml Methylparaben mold inhibitor (optional): 20 g 95% Ethanol (optional): 200 ml

Cooking directions 1. Fill steam kettle with 17 l de-ionized water, depending on relative humidity (add less water when humidity is high). 2. Set kettle to power 5 for five minutes. 3. Turn kettle up to power 7.5 and bring to a RAPID BOIL (this usually takes about 15 minutes). 4. When the water has begun to boil rapidly, whisk dry mix with 6 l de-ionized water in a separate container. Turn the kettle mixer on to power 3, and then add the dry mix – 6 l of de-ionized water mixture to the 17 l of boiling water in the kettle. (Our dry mix is pre-weighed and stored at 20°C in a Ziploc bag, for up to six months in advance). 5. Turn the kettle mixer down to power 1.5–2 about a minute or two after adding the 6 l mixture to the boiling water. 6. WATCH CAREFULLY SO THAT THE CORNMEAL MIXTURE DOES NOT OVERBOIL. 7. Cook ingredients at a GENTLE BOIL for 15 minutes. 8. Gradually add an additional 5 l of de-ionized water while food cooks, as an overboil preventative. 9. After 15 minutes of cooking, turn the kettle power off but continue letting the kettle mixer mix the cornmeal food until cooled to pouring temperature. 10. Dissolve the mold inhibitor in ethanol (optional). 11. Cool food to 63°C and add propionic acid and ethanol–mold inhibitor solution. Continue letting food mix until cooled to 60°C. 12. Pour food mixture at 60°C.

Standard banana–opuntia (makes approximately six cases of vials, or three cases of half-pint bottles) Ingredients Dry mixture: Agar: 39 g Yeast: 165 g Methylparaben: 12 g Wet mixture: Blended bananas: 825 g Karo syrup: 285 g

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Malt: 180 g Powdered opuntia cactus: 12.75 g

Cooking directions 1. Bring 6 l de-ionized water to a boil in a large pot. (This recipe uses a conventional electrical stove top for our heat source. The temperature setting is HIGH). 2. Have pre-measured dry and wet mixtures ready for use. (Both mixes are prepared in advance and stored at 20°C in a Ziploc bag for up to six months). 3. Boil water vigorously for ten minutes and add the dry mix. Keep the temperature source setting at high heat. 4. Stir the mixture continuously. 5. Cook for ten minutes, without overboiling. Add the wet mixture to the pot. 6. Stir the mixture continuously for an additional ten minutes. 7. Remove pot from heat source and cool in a water bath. 8. Stir food continuously to prevent clumping. 9. Allow food to cool to 40°C and pour at exactly 40°C (pouring food below 40°C is too cold and food will clump; pouring food above 40°C is too hot and food will separate prior to plugging).

Standard Wheeler–Clayton (makes approximately six cases of wide-diameter plastic vials) Ingredients Top layer: Cornmeal: 201 g Special K: 67.5 g Wheat germ: 67.5 g Product 19: 37.8 g Brewer’s yeast: 135 g Hi-protein baby cereal: 102.6 g 3 medium bananas (300 g) Agar: 29.7 g Tap water: 6.75 l 95% Ethanol: 43.2 ml Propionic acid: 43.2 ml Bottom layer: Instant Drosophila medium: 1.5 cups Boiled tap water: 600 ml

Cooking directions 1. Mix the instant Drosophila medium with the 600 ml of boiled tap water in a small Ziploc bag. Allow closed Ziploc to sit and cool to a “touchable” temperature. Cut

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2.

3.

4. 5.

6. 7. 8. 9. 10.

a corner out of the Ziploc bag and squeeze a small dab (about 1 ml) into the bottom of each vial. Take out the dry cereal mix (containing all of the cereals, blended to a powder – dry mixes are prepared ahead of time and stored at 20°C in a Ziploc bag for up to six months). In a large pot, add together 3 l of tap water and agar. Bring to a boil on the top of a stove. Stir the agar solution CONSTANTLY to prevent agar from burning to the bottom of the pot. Cook at a boil until agar is clear. This process can take up to thirty minutes. Blend the three medium bananas, the dry cereal mixes, and 1.75 l water in a large blender. Once the agar solution is clear, add in the blended wet mixture to the agar solution into the pot. Add an additional 2 l of tap water to the pot (using this additional 2 l to wash all of the cereal mix out of the blender and into the pot of agar solution). Lower the temperature slightly to prevent scalding. STIR CONSTANTLY; allow the mixture to simmer for ten minutes. Carefully add in the 95% ethanol, then propionic acid; mix thoroughly. Continue simmering for five more minutes. Remove from heat and pour into vials (it does not need to cool; the mixture can be poured immediately after removing from heat).

Saguaro–potato (makes approximately 24 half-pint bottles) Ingredients Saguaro cactus (autoclaved): 188 g De-ionized water: 1600 ml Potato buds (finely ground in blender): 282 g Propionic acid: 18.8 ml

Cooking directions 1. Put the finely ground potato buds in a large Ziploc bag. 2. Add autoclaved saguaro and de-ionized water to a blender. Blend at high speed for approximately sixty seconds. Make sure texture of blended solution is smooth and creamy, and that there are no chunks of saguaro. 3. Pour blended mixture into a medium-sized pot. Bring mixture to approximately 90°C (small bubbles, no boil). 4. Allow mixture to cool on a cold stove-top burner to 60°C (watch closely – this will happen quickly). Stir continuously. 5. Add propionic acid to mixture. 6. At 60°C, pour hot cactus suspension into large Ziploc bag containing the potato buds. 7. Knead/mix thoroughly to uniform consistency. 8. Cut a corner of the Ziploc bag and dispense by squeezing into glass bottles. Squeeze approximately 79 ml or one-third cup of mixture into each bottle.

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225

Special treatments and requirements Several examples exist in which a species or group of species requires a particular material or chemical in order to grow. For example, not only do mycophagous Drosophila species have the ability to tolerate alpha-amanitin (Jaenike et al., 1983), but also many of them can only be reared using fresh mushrooms (Grimaldi and Jaenike, 1982). For culturing species of the testacea group (testacea, neotestacea, orientacea, and putrida) and the mycophagous members of the quinaria group (guttifera, falleni, recens, subquinaria, tenebrosa, suboccidentalis, phalerata, innubila, munda, nigromaculata, and transversa), John Jaenike (personal communication) uses a mixture of instant food and fresh mushrooms. One teaspoon of Carolina instant food is mixed in a vial with 8 ml water. A piece of mushroom and a dental cotton roll are then added to the food surface and the flies are introduced. The mushrooms become liquefied quickly when the temperature is too high, so keeping cultures at 21–22°C works best for all species in the list. The Tucson Stock Center has been able to maintain D. falleni, D. guttifera, D. phalerata, and D. putrida on some of the standard media (Table 9.1), but even these will perform better if mushrooms can be provided. A similar situation exists with Drosophila pachea, a nannoptera group species that breeds exclusively in senita cactus, Lophocereus schottii. Not only is D. pachea the only Drosophila with the ability to tolerate the alkaloids found in this cactus species, but also it is nutritionally dependent upon a sterol unique to senita (Heed and Kircher, 1965). Some investigators find that adding powdered senita cactus to the medium will enable flies to be cultured. We find that using chunks of autoclaved cactus tissue is a better approach. Investigators interested in rearing D. pachea may be able to obtain plant material from their local botanical garden. The Tucson Stock Center is also a source of cactus material for culture medium. Investigators wishing to establish new cultures from wild-caught specimens of species not reared in the Stock Center may need to try a variety of recipes. If the species is suspected of being a specialist, we recommend incorporating any information about the natural feeding and breeding site of the species into the culturing approach. This may mean adding a piece of natural material or a few drops of some juice to the food surface.

References Brncic, D. (1983). Ecology of flower breeding Drosophila. In: The Genetics and Biology of Drosophila (M. Ashburner, H. L. Carson and J. N. Thompson, eds), Vol. 3d, pp. 333–377. Academic Press, London. Carson, H. L. (1967). The association between Drosophila carcinophila wheeler and its host, the land crab, Gecarcinus ruricola. Am. Midland Naturalist 78, 324–343. Grimaldi, D. A. and Jaenike, J. (1982). Laboratory culturing of mycophagous Drosophilids. Drosophila Information Service 58, 156–157.

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How to use living flies Heed, W. B. and Kircher, H. (1965). Unique sterol in the ecology and nutrition of Drosophila pachea. Science 149, 758–761. Jaenike, J., Grimaldi, D. A., Sluder, A. E. and Greenleaf, A. J. (1983). Alpha-amanitin tolerance in mycophagous Drosophila. Science 221, 165–167. Markow, T. A., Raphael, B., Breitmeyer, C. M., Dobberfuhl, D., Elser, J. and Pfeiler, E. J. (1999). Elemental stoichiometry of Drosophila and their hosts. Funct. Ecol. 13(1), 78–84. Patterson, J. T. and Wheeler, M. (1942). Description of new species of the subgenera Hirtodrosophila and Drosophila. Univ. Texas Publ. 4213, 67–109.

CHAPTER 10

Avoiding experimental artifacts: using information about life history and other differences in experimental design Contents

•

• •

Timing issues • Egg to adult development time • Sexual maturation • Diurnal factors • Female remating frequency • Copulation duration • Age effects Social influences on behavior References

From Chapters 7 and 8 it should be clear that Drosophila species can vary in many ways beyond their obvious physical phenotypes. These other differences can profoundly influence the outcome of experiments and, if not controlled for, lead to erroneous conclusions. Species differences should be considered when setting up all aspects of organismal and population studies, as well as for routine stock maintenance. In designing experiments, the particular species differences requiring control will depend upon the question being addressed. We are unable to anticipate all of the situations in which various factors will need to be controlled, but we can provide some specific examples related to common types of experiments.

Timing issues Here we will deal with several different categories of timing issues: egg to adult development time, sexual maturation, diurnal factors, female remating, copulation duration, and longevity.

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Egg to adult development time In experiments designed to compare two or more species for some behavioral or physiological trait, or a character for which behavior or physiology are underlying components, if the species differ in development time (Tables 7.1 and 7.2), cultures will need to be set up at different times. Cultures of species with longer development times should be set up earlier, such that adults of all species will emerge on the same day. Setting up cultures so that adults of the different species begin emerging on the same day will ensure that flies being compared are the same age. At the same time, other interspecific differences in the ages at which flies become sexually mature or die should be considered as they may influence the outcome of experiments.

Sexual maturation Depending upon the question the experiment is designed to answer, the investigator may need simultaneously to control for differences in age at reproductive maturity. Suppose, for example, that a comparative experiment is planned in which the courtship vigor of males of several species is to be compared. If the species differ in the age at which adult males become sexually mature (Table 7.3), courtship vigor should not be measured on all the species on the same day after they eclose. To do so could result in males of some species being scored as having low courtship vigor, when in reality they are simply sexually immature. A related (and very common) kind of experiment involves testing for sexual isolation between two related species using “choice” tests. In choice tests, males or females of a given species are presented with two potential mates – one from their own species and one from a related species – and the number of matings involving each type is scored and tested for departure from random mating. If females of two species differ, even by only a day or two, in the onset of sexual receptivity, females of the “slow” maturing species will be less interested in mating at earlier ages. A significant departure from random mating may be observed, but in this case low receptivity by females of the late maturing species would be due to sexual immaturity rather than to discrimination or true sexual isolation. Furthermore, the length of time virgin flies are stored prior to use in mating experiments can significantly alter their mating speed and discrimination among prospective mates (Long et al., 1980). Knowing the age at which flies become mature will enable the investigator to use them in a timeframe that reduces the potential for age-related artifacts to cloud experimental results. For the species listed in Table 7.3 the male and female sexual maturity ages are known and can be used in planning experiments, but for uninvestigated species the ages at which flies become mature should be determined prior to the experiments.

Diurnal factors Differences among species in their response to diurnal factors have not been well studied. Evidence exists, however, to suggest that diurnal factors be considered in comparative behavioral studies. For D. melanogaster, Hardeland (1972) reported that mating activity

Avoiding experimental artifacts

229

occurs at peak times in the morning and late afternoon, but that these times differ in other melanogaster group species. In the laboratory, Grossfield (1970) examined the dependence of courtship and mating on the presence of light in a number of Drosophila species. Some species were found to mate in darkness as well as in light, while others mate only in the light. Interestingly, D. pseudoobscura was found to be light independent, while its sibling species, D. persimilis, mated only in the light. These data suggest that closely related species may differ in the relative influence of light levels on their mating, and thus may also exhibit different sensitivities to diurnal effects in experiments – for example, one species may have a narrower or different window of mating activity than does a related species for which sexual isolation is being studied. Thus conducting sexual isolation experiments at different times of the day could yield different results. More recently, Drosophila has been shown to “sleep” and exhibit circadian cycles of “arousal” (Van Swinderen and Andretic, 2003; Zimmerman et al., 2004) and activity (Cobb et al., 1987), which may be found to influence a broad range of behaviors and their measurement.

Female remating frequency Drosophila species exhibit extreme differences in the incidence of female remating (Markow 1996, 2002), from species such as D. hydei in which up to five matings have been observed in a given morning, to species such as D. subobscura or D. acanthoptera in which females typically mate only once (Table 10.1). Differences in female remating frequency can have enormous impacts on studies of sexual selection, and reproduction. As discussed earlier, choice tests are a common means of testing for sexual isolation. Choice tests typically are scored either by direct observation of the matings of marked flies in observation chambers, or by scoring progeny produced by flies confined overnight to infer mating types. In these experiments the null hypothesis of random mating assumes that both types of females have equal probabilities of mating per se. A difference in remating frequencies between species, especially if the remating latency is within the experimental timeframe, will bias the data towards matings involving females of one of the species. Thus when differences in female remating exist, individual flies should be directly observed until they mate. Sperm utilization studies usually involve mating females to two genetically distinguishable types of males. The underlying question in these studies is how sperm of more than one male are used, relative to each other and/or the female’s genotype, in fertilization. A common assumption is that when a female’s progeny contains progeny of both types, that the female mated twice, and deviations from a 50:50 ratio of paternal markers in the progeny is attributable to some sort of sperm precedence or unequal sperm utilization. In sperm utilization experiments, females are mated to males of a known genotype and then remated to males of a different genotype. Because these sorts of experiments typically involve large numbers of rematings, and directly observing mating may be very time consuming, one common approach to obtaining remated females is to place them with males unobserved for 24 hours, and then remove the females to look for markers in their progeny as an indication of how sperm were utilized. If the progeny of a given female include both types of males she was tested with, it is assumed that she mated twice. However, in rapidly remating species a female could

230

How to use living flies Table 10.1. Frequency of female remating, as proportion of females having remated after 24 hours and the average number of days until females remate during a two-week period Species

% Remating in 24 h (n matings in 24 h)

Average number of days until remating

Reference no.

D. acanthoptera D. affinis D. aldrich D. arizonae D. biarmipes D. buzzatii D. guttifera D. grimshawi D. heteroneura D. hydei D. littoralis D. melanogaster D. mettleri D. mojavensis D. montana D. nannoptera D. nigrospiracula D. pachea D. persimilis D. pseudoobscura D. recens D. silvestris D. simulans D. subobscura

0.00 0.50 0.95 0.95 0.0 0.95 1.00 (1) 0.00 0.00 1.00 (4) 1.00 (3) 0.02 1.00 (1) 0.95 1.00 (3) 0.96 1.00 (4) 0.96 0.05 0.05 1.00 (1) 0.0 0.02 0.0

No remating 3 1 1 No remating 1 1 No remating No remating Same day Same day 5 Same day 1 Same day 1 Same day 1 3 4 1 No remating 4 No remating

1 2 3 1 4 3 5 6 7 8 9 8 5 5 9 1 1 1 2 2 5 7 1 10

1
Markow, 1982; 2
Snook and Markow, 2001; 3
Markow, unpublished; 4
McRobert et al 1997; 5
Markow, 2002; 6
Schwartz and Boake, 1992; 7
Wisotsky, 1987; 8
Markow, 1985; 9
Aspi and Lankinen, 1992; 10
Maynard Smith, 1956.

have remated three or more times. Progeny counts thus could suggest that the sperm of the second male were more successful in siring progeny, whereas in fact the female may simply have mated more times to the second male and have a greater number of his sperm type. This situation will be more common in species such as D. hydei, in which females remate up to several times a day, but could occur in almost any species.

Copulation duration Drosophila species exhibit enormous differences in copulation duration (Table 10.2). Drosophila melanogaster is by no means representative. In fact, in some species, such as D. mulleri, copulation is so fast that it might easily be missed. In other species, like D. acanthoptera, flies may remain in copula for two hours. Although copulation duration has never been associated with any other reproductive character, such as the

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231

Table 10.2. Copulation durations of a subset of Drosophila species maintained at the Tucson Stock Center (Grant, 1983; Pitnick et al., 1991; Pitnick and Markow, 1994) Species

Minutes ( seconds)

Species

Minutes ( seconds)

D. acanthoptera D. affinis D. aldrichi D. algonquin D. americana D. ananassae D. arizonae D. auraria D. azteca D. buskii D. buzzatii D. canapalpa D. equinoxalis D. funebris D. fumipennis D. gibberosa D. guttifera D. hamatofila D. hydei D. immigrans D. lacicola D. longicornis D. macrospina D. melanica D. melanogaster D. melanopalpa D. mercatorum D. micromelanica D. miranda D. mojavensis

137.0 1.5 1.3 5.3 2.3 4.1 1.4 6.4 5.2 1.2 2.0 2.2 16.0 17.0 4.2 6.4 7.0 9.3 2.1 45.0 3.1 2.2 32.5 7.0 18.0 1.5 2.0 7.0 8.5 3.0

D. montana D. mulleri D. nannoptera D. nebulosa D. neocardini D. nigrohydei D. novamexicana D. pachea D. paramelanica D. paulistorum D. paulstris D. peninsularis D. persimilis D. polychaeta D. prosaltans D. pseudoobscura D. repleta D. ritae D. robusta D. rufa D. simulans D. sturtevanti D. sucinea D. takahashi D. tripunctata D. tropicalis D. virilis D. wassermani D. willistoni

3.2 0.3 7.5 2.1 21.2 7.3 2.4 39.3 4.4 15.3 7.2 1.4 6.0 1.0 11.3 7.0 2.1 9.0 0.4 10.2 13.0 20.0 17.0 17.3 38.1 3.0 12.2 14.1 17.3

number of sperm transferred or female remating incidence, it can still greatly confound the outcome and interpretation of certain experiments, especially choice tests for sexual isolation (discussed above). For example, males of most Drosophila species are ready to court and remate immediately after copulating. If two species differ significantly in copulation duration and they are in the same choice test, males of the species with the shortest copulation will be ready to re-enter the pool of courting males more frequently than males involved in longer copulation. The potential will then exist for more matings to occur with these short copulation duration males because there will be more of them at any one time, and not necessarily because they are “preferred”. These are problems that can be controlled by the experimental design. Rather than using mating chambers with large numbers of flies for choice tests, we recommend using single vials with either one female and two males (“female choice test”),

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one male and two females (“male choice test”) or two females and two males (multiple choice tests) so that individual flies and rematings can be tracked.

Age effects Just as species differ in their development times and the ages at which they attain sexual maturity, they differ in how long they live (Yoon et al., 1990), making it likely that age-dependent metabolic changes observed in D. melanogaster (Collins and Kiefer, 1979; Promislow and Hselkorn, 2002) will be found. Resistance to stress, such as that caused by desiccation (Hoffmann and Harshman, 1999), heat (Goto and Kimura, 1998; Stratman and Markow, 1998), starvation (Hoffmann and Harshman, 1999), and cold (Goto and Kimura 1998), has been found to be age-dependent. At present, insufficient data exist to determine the relationship, if any, between longevity and physiological processes. Therefore in comparative studies it is advisable to control for such differences, perhaps by examining the character of interest in each species across several ages rather than assuming flies of all species will be the same at, say, five days of age.

Social influences on behavior Sexual behavior in Drosophila is particularly sensitive to social interactions among individuals prior to their use in experiments. Social interactions are either intra- or intersexual. For practical purposes, we will confine our comments to the conditions under which adult flies are stored prior to their use in experiments. Most investigators store virgin flies in vials with others of the same sex, whether they are to be used to study sexual or non-sexual behaviors. The density at which flies of either sex are stored or tested can influence their performance in behavioral studies (Connolly, 1968; Knoppien, 1986; Crossley and Wallace, 1987). Higher-density storage can greatly reduce male courtship vigor, putting males stored at higher-density conditions at a disadvantage in studies of sexual isolation. Higher storage densities can also lead to reduced levels of general activity in flies of both sexes (Markow, unpublished observation). Intersexual social interactions also can influence mating propensities. Since flies are typically stored in same-sex groups, intersexual social interactions are a function of contamination by a member of the opposite sex. When a male is found in a vial thought to house virgin females, the entire vial typically is discarded, as all females may have mated. However, when a female is found in a vial of virgin males, because it is less likely in many species that all of the males will have mated with the female, it is tempting simply to remove the female and retain the males for the experiment. Males that have been kept with a mated female, however, even if they are still virgin, tend to be significantly slower to court and mate (Siegel and Hall, 1979; Reif et al., 2002). Therefore, we recommend discarding vials of males in which a female has been found. We also recommend that, for use in behavior experiments, flies be stored at densities of no more than ten per vial. Not only does this control density effects, but also, in the event that a contamination is found, fewer flies have to be discarded.

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References Aspi, J. and Lankinen, P. (1992). Frequency of multiple insemination in natural populations of Drosophila Montana. Hereditas 117, 169–177. Cobb, M. J., Connolly, K. and Burnet, B. (1987). The relationship between locomotor activity and courtship in the melanogaster species subgroup of Drosophila. Animal Behav. 35, 705–713. Collins, M. and Kiefer, B. I. (1979). Age dependent changes in RNA metabolism in Drosophila melanogaster. Genetics 91, s23–24. Connolly, K. (1968). The social facilitation of preening behavior in Drosophila melanogaster. Animal Behav. 16, 385–391. Crossley, S. and Wallace, B. (1987). The effects of crowding on courtship and mating success in Drosophila melanogaster. Behav. Genetics 17, 513–522. Goto, S. G. and Kimura, M. T. (1998). Heat and cold shock responses and temperature adaptation in subtropical and temperate species of Drosophila. J. Insect Physiol. 44, 1233–1239. Grant, B. (1983). On the relationship between average copulation duration and insemination reaction in the genus Drosophila. Evolution 37, 854–856. Grossfield, J. (1970). Species differences in light influenced mating behavior in Drosophila. Am. Naturalist 104, 307–309. Hardeland, R. (1972). Species differences in the diurnal rhythmicity of courtship behavior within the melanogaster group of the genus Drosophila. Animal Behav. 20, 170–174. Hoffmann, A. and Harshman, L. (1999). Desiccation and starvation resistance in Drosophila: patterns of variation at the species, population and intrapopulation levels. Heredity 83, 637–643. Knoppien, P. (1986). Low density storage enhances mating speed in Drosophila melanogaster. Drosophila Information Service 63, 76–77. Long, C., Markow, T. A. and Yaeger, P. (1980). Relative male age, fertility, and competitive mating success in Drosophila melanogaster. Behav. Genetics 10, 163–170. Markow, T. A. (1979). Phototactic behavior of Drosophila species at different temperatures. Am. Naturalist 114, 884–892. Markow, T. A. (1982). Mating systems of cactophilic Drosophila. In: Ecological Genetics and Evolution: The Cactus–Yeast–Drosophila Model System (J. S. F. Barker and W. T. Starmer, eds), pp. 273–287. Academic Press, New York. Markow, T. A. (1985). A comparative investigation of the mating system of Drosophila hydei. Animal Behav. 33, 775–781. Markow, T. A. (1996). Evolution of Drosophila mating systems. Evol. Biol. 29, 73–106. Markow, T. A. (2002). Female remating, operational sex ratio and the arena of sexual selection. Evolution 59, 1725–1734. Maynard Smith, J. (1956). Fertility, mating behavior and sexual selection in Drosophila subobscura. J. Genetics 54, 261–279. McRobert, S. P., Adams, C. R., Wutjke, M., Frank, J. and Jackson, L. L. (1997). A comparison of female post-copulatory behaviour in D. melanogaster and D. biarmipes. J. Insect Behav. 10, 761–770. Pitnick, S. and Markow, T. (1994). Male gametic strategies: sperm size, testes size and the allocation of ejaculate among successive mates to the sperm-limited fly, Drosophila pachea and its relatives. Am. Naturalist 143, 785–819. Pitnick, S., Markow, T. and Reidy, M. (1991). Transfer of ejaculate and incorporation of malederived substances by females in the nannoptera species group of Drosophila. Evolution 45, 774–780.

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How to use living flies Promislow, D. E. and Haselkorn, T. S. (2002). Age-specific metabolic and mortality rates in the genus Drosophila. Ageing Cell 1, 66–74. Reif, M., Linsenmair, K. F. and Heisenberg, M. (2002). Evolutionary significance of courtship conditioning in D. melanogaster. Animal Behav. 63, 143–155. Schwartz, J. M. and Boake, C. R. B. (1992). Sexual dimorphism in remating in Hawaiian Drosophila species. Animal Behav. 44, 231–238. Siegel, R. W. and Hall, J. C. (1979). Conditioned responses in the courtship behavior of normal and mutant Drosophila. Proc. Natl. Acad. Sci. 76, 3430–3434. Snook, R. R. and Markow, T. A. (2001). Mating system evolution in sperm-heteromorphic Drosophila. J. Insect Physiol. 47, 957–964. Stratman, R. and Markow, T. A. (1998). Heat tolerance of Sonoran Desert Drosophila. Funct. Ecology 8, 965–970. Van Swinderen, B. and Andretic, R. (2003). Arousal in Drosophila. Behav. Processes 64, 133–144. Wisotsky, R. G. (1987). Sperm competition in Drosophila silvestris. MSc thesis, University of Hawaii, Honolulu. Yoon, J. S., Gagen, K. P. and Zhu, D. L. (1990). Longevity of 68 species of Drosophila. Ohio J. Sci. 90, 16–32. Zimmerman, J. E., Mackiewicz, M., Galante, R. J. et al. (2004). Glycogen in the brain of Drosophila melanogaster: diurnal rhythm and the effect of rest deprivation. J. Neurochem. 88, 32–40.

CHAPTER 11

Troubleshooting Contents

• • • • • • •

Cultures too dry? Cultures too wet? Food crashes down when collecting? Cultures contaminated with mold, bacteria, or mites? Flies that won’t mate? Females won’t oviposit? References

When working with different species, the likelihood of encountering problems in rearing flies or getting flies to cooperate in behavioral studies is increased greatly over those found with D. melanogaster. The best antidote to such frustrations is familiarity with the biology of the species. In general, knowing about a species’ natural history and reproductive biology will facilitate working with it in the laboratory. Besides finding and reading the published literature about a particular species, directly contacting investigators with previous experience and asking them about peculiarities and how they overcame them can save considerable time and effort. There are some general problems encountered in culturing and experimenting with flies, however, that we will discuss in this chapter.

Cultures too dry? Dry, hard cultures can result from one or more factors. If the laboratory is in a part of the world with low relative humidity, or work is being done in a dry season of the year, the food recipe may simply need to have the percentage of water increased (starting with 10%). Another explanation is that the food is old (more than four to five days) and it could have dried out, providing a substrate that is either unattractive to ovipositing females or too difficult for the mouthparts of first instar larvae to penetrate. Sealing flats of vials or bottles of food tightly in plastic wrap until used will reduce loss of moisture from the food. Finally, if the adults are introduced to fresh culture medium before they are reproductively mature, the food may dry out before females are ready to oviposit. Be familiar with the reproductive biology of the species in order to start cultures with mated, gravid females.

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Cultures too wet? Larvae of some species, such as D. hydei, tend to create very “soupy” cultures. With any species, too many larvae working the medium can cause runny food. Using fewer adults to start the cultures or leaving the adults in the cultures for a shorter period of time will help avoid this problem. Once cultures have become runny, a piece of tissue (in addition to the one used when the culture was created) can be folded twice and inserted down into the medium. The tissue will absorb extra moisture as well as providing pupation sites for late third instar larvae. Extra tissue can be placed in the cultures several times, a day or two apart, if problems continue during virgin collecting.

Food crashes down when collecting? Collecting virgin flies involves repeatedly inverting cultures and “pounding” out newly emerged adults for sorting. The continual pounding can quickly cause the food to loosen from the container wall and crash down, often smashing flies in the process. Several practices can reduce this problem. Figure 8.2 demonstrates how to hold the inverted bottle to the side, to reduce the stress on the culture medium while removing flies. Another approach is to reduce the amount of pounding per se. Bottles are pounded to be sure all flies are removed, especially those hiding in crevices. By first injecting the bottle with carbon dioxide, flies will be immobilized and thus not able to hide. If there are not huge numbers of flies in the bottle a foam plug can be inserted into the mouth, temporarily replacing the cap or other rigid type of plug, to allow an aspirator to be inserted into the bottle to remove the flies. Remember, when flies are excited (which they will be when their culture bottle or vial is picked up) they exhibit negative geotaxis and positive phototaxis and will, if the culture is not inverted, fly or run out of the open space through which the aspirator is inserted. The foam plug will help by expanding into the space, but the vial should be held as shown in Figure 8.4.

Cultures contaminated with mold, bacteria, or mites? In general, keeping the work area and tools (such as brushes, sorting plates, and forceps) clean will cut down on problems from fungi and bacteria. These tools, as well as the bench tops near the microscopes, should be cleaned weekly with either 70% ethanol or 10% household bleach. Obviously the use of appropriate levels of mold inhibitors (see Chapter 9) is critical and can be adjusted in problem cases, but other practices can also help to control mold. Once cultures are contaminated, using adults from moldy cultures to start new cultures can overpower the potency of the mold inhibitors. Using clean flies for propagation typically is sufficient to control microbial contamination. Removing emerging adults from moldy cultures and transferring them daily for one week to fresh food containers prior to staring new cultures allows the flies to clean themselves so that they are not vectoring microbes. Fungal contamination

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may be addressed by adding the fungal inhibitor p-hydroxy-benzoic acid methyl ester to the medium after it is cooked, as described in Chapter 9. A few grains of live baker’s yeast added to the surface of a culture also tends to inhibit the growth of undesirable fungi, but should only be used for non-mycophagous species. Drosophila cultures also can become contaminated by bacteria, which in some cases are brought into the lab by wild-caught flies. When failed cultures are associated with a layer of slime on the surface of the food, there is a good chance that bacterial contamination is responsible. Females may refuse to oviposit on surfaces with bacterial material, but if they do their eggs may not develop or the larvae could become trapped in the slime. Normally the presence of larvae overcomes the effects of the bacteria but, as discussed in Chapter 8, cultures of species with delayed reproductive maturity or low fecundity may be more susceptible to the effects of bacteria. Several adjustments to the culture medium can be tried in order to control bacteria. One approach is to lower the pH of the food to about 5. Another is to try adding PenStrep, a combination of penicillin and streptomycin, to the medium. Since each species is likely to differ with respect to its response to antibiotics, attaining the appropriate levels of these drugs may require some experimentation. Mites of several types pose serious threats to Drosophila labs, and unfortunately species other than D. melanogaster, because of their longer generation times, are more vulnerable to being wiped out by laboratory mites. The most detailed review of mites that infest Drosophila cultures is found in Ashburner (1989). Some mite species cause their harm by simply out-competing the larvae in the culture medium. Other mite species are predacious on eggs and larvae. The longer a culture must be kept because of fly development time, the greater the opportunity for mites to develop. Reservoirs of mites can exist in forgotten culture bottles sitting in corners in the laboratory. No culture bottle, whether mites have been experienced in a laboratory or not, should be kept around any longer than necessary to harvest the next generation of adults. Furthermore, cultures should be examined for the presence of mites, not just on the container walls or plugs but in the food as well. If a culture is discovered to have mites, any living adults should be transferred to a clean culture vial and the contaminated culture discarded, frozen, or autoclaved immediately. Adults from mite-infested cultures should be transferred daily for a week before they are used to start new cultures. All the transfer vials should be discarded immediately. Wiping all surfaces with 10% benzyl benzoate in 95% ethanol helps prevent mites from moving between cultures, as does using benzyl benzoate-soaked plugs. If plugging plastic vials or bottles, test the containers first to be sure that they are not of a type of plastic dissolved by this substance. In our experience, most mite contaminations result from the receipt of mite-infested fly cultures from other laboratories, rather than from wild-caught flies. We recommend that all fly shipments not only be inspected for mites but also quarantined for at least two generations after they are received. If contamination problems with mold, bacteria or mites persist, another solution is to start with axenic cultures, using clean, dechorionated embryos (Starmer and Gilbert, 1982; Ashburner, 1989). Embryos can be collected from the oviposition substrate and treated with 5.25% liquid household bleach (sodium hypochlorite) for three to five minutes and then washed with Ringers solution. Embryos can then be transferred to fresh medium to develop.

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Flies that won’t mate? One of the most frustrating experiences encountered occurs when a huge amount of effort has gone into collecting virgin flies for a mating behavior experiment or cross, and the flies won’t mate. If this happens there are several things to check. The ages at which reproductive maturity is attained should be checked to be sure that the flies are old enough, but not too old. Another problem is that many Drosophila species lack the obvious sexual dimorphisms of D. melanogaster (Markow, 2002), so it is easier to make mistakes when collecting virgins. As pointed out in Chapter 10, a single female accidentally placed in the same vial with males during storage will cause those males to exhibit reduced or even a complete lack of interest in courtship (Siegel and Hall, 1979; Tompkins et al., 1983). Females, even in species exhibiting frequent female remating, if confined with just one male for several days will also be uninterested in mating. If these potential sources of failed matings can be ruled out, two other environmental factors that can prevent or reduce courtship should be considered. The time of day that the experiment is set up can be critical (Hardeland, 1972), as many species have a very narrow window of time during the day during which they mate. This window is controlled by the diurnal light cycle. The timeframes for most species have not been worked out in detail, so this could be a situation in which some preliminary work will be required. In the absence of such data, however, we recommend storing flies to be used for mating experiments on a strict light:dark cycle of 12:12, 14:10 or 16:8, and setting up the experiments (i.e. placing the males and females together) immediately after “lights-on”. Maintaining cultures in environmental conditions that best mimic those encountered in the flies’ natural habitat is a recommended practice. The other environmental factor that impacts mating activity in many species is outside barometric pressure. Documented originally in D. pseudoobscura (Ankney, 1984), we have observed that flies of many species show little interest in courtship during storms or stormy seasons of the year. Unfortunately, unlike the situation with diurnal light cycles, we have not found anything that can control the impact of weather systems on mating experiments other than trying to time the experiments to take place during optimal periods. Obtaining F1 hybrids between two species is often prevented by sexual isolation. Manipulating the experimental conditions can sometimes increase the likelihood of getting an interspecific cross to work. Within species, for example, social facilitation of mating is well known (Crossley and Wallace, 1987). In groups of flies, when one mating takes place the others in the chamber are stimulated to mate. By placing large numbers of males of one species together with many females of a different species, social facilitation may lead to an increase in the number of interspecific matings obtained. Another trick worth trying is to use either extremely young virgin females or virgin females that are several days past the time when they become reproductively mature. Also, using virgin males that are about a week past their maturity age may lead, in the very least, to more persistent courtship.

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Females won’t oviposit? Sometimes everything seems to be going well, but females don’t seem to want to lay eggs. The first thing to verify is that the females have in fact been inseminated. Observing a mating doesn’t guarantee that females will carry or use sperm. Several females could be dissected to verify that sperm are present in their storage organs. Not all species of females use both the ventral receptacle and spermathecae to store sperm (Pitnick et al., 1999), so be sure, if dissections are performed, to examine both types of storage organs. In some species females may be inseminated but will not utilize sperm from closely related males (Markow, 1997). Here, providing females with more than one male would be useful – unless performing sibmatings, in which case the failure rate may be high. If females are full of sperm, they might be induced to oviposit by increasing the attractiveness of the substrate. This could be as simple as scratching the medium surface with the end of a brush, as females prefer surfaces that are easily penetrated by early instar offspring. For specialist species, females may be stimulated by the natural substrate – such as Morinda fruit in the case of D. sechellia (Amlou et al., 1998), cactus in the case of repleta and nannoptera group species, or a tissue soaked in Clermontia juice (Chapter 10) and inserted into the medium in the case of Hawaiian Drosophila.

References Amlou, M., Moreteau, B. and David, J. R. (1998). Genetic analysis of Drosophila sechellia specialization: oviposition behavior toward the major aliphatic acids of its host plant. Behav. Genetics 28, 455–464. Ankney, P. F. (1984). A note on barometric pressure and behavior in D. pseudoobscura. Behav. Genetics 14, 315–317. Ashburner, M. (1989). Drosophila: A Laboratory Handbook. Cold Spring Harbor Press, New York. Crossley, S. and Wallace, B. (1987). The effects of crowding on courtship and mating success in Drosophila melanogaster. Behav. Genetics 17, 513–522. Hardeland, R. (1972). Species differences in the diurnal rhythmicity of courtship behavior within the melanogaster group of the genus Drosophila. Animal Behav. 20, 170–174. Markow, T. A. (1997). Assortative fertilization in Drosophila. Proc. Natl Acad. Sci. 94, 7756–7760. Markow, T. A. (2002). Operational sex ratio, female remating and the arena of sexual selection in Drosophila. Evolution 59, 1725–1734. Pitnick, S., Markow, T. A. and Spicer, G. (1999). Evolution of multiple kinds of female sperm storage organs in Drosophila. Evolution 53, 1804–1822. Siegel, R. W. and Hall, J. C. (1979). Conditioned responses in the courtship behavior of normal and mutant Drosophila. Proc. Natl. Acad. Sci. 76, 3430–3434. Starmer, W. T. and Gilbert D. (1982). A quick and reliable method for sterilizing eggs. Drosophila Information Service 58, 170–171. Tompkins, L., Siegel, R.W., Gailey, D. A. and Hall, J. C. (1983). Conditioned courtship in Drosophila and its mediation by association of chemical cues. Behavior Genetics 13, 565–578.

CHAPTER 12

Links to sources for supplies and equipment Contents

• • • • •

Field work and equipment Media supplies and equipment, laboratory supplies • Ingredient brands • Cooking equipment Flies for sale Species sequencing and BAC library projects Other Drosophila resources and databases

Field work and equipment Aspirators: Bioquip, http://www.bioquip.com Wards Scientific, http://www.wardsci.com Tubes for ethanol or liquid nitrogen preserved samples: Herman J. Phaff Yeast Collection at the University of California at Davis, http://www.phaffcollection.org./home.asp Tanglefoot, http://www.tanglefoot.com VWR International, http://www.vwrsp.com Collecting Permits (USA): Import Permits for the United States, United States Department of Agriculture import permits, http://www.aphis.usda.gov/ppq/permits/ International Organizations of Biological Field Stations, http://www.iobfs.org United States National Parks and Monuments, http://www.nps.gov Nets Genesee Scientific (http://www.geneseesci.com) Anesthetizers Genesee Scientific (http://www.geneseesci.com)

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Media supplies and equipment, laboratory supplies Ingredient brands Agar: Fine ground agar. Moorhead & Company (1-800-423-3170), http://www.moorhead-agar.com/ Yeast: Red Star Nutritional Yeast (inactive). Le Saffre Yeast Corporation (toll free 1-800-641-4615), http://www.redstaryeast.com/ (This is the yeast that is mixed into the food for cooking.) Mold inhibitor: Methylparaben powder. Carolina Biological Supply Co. (toll free 1-800-287-9989), http://www.carolina.com Karo syrup: Light. ACH Food Company (purchased at any major grocery store). Malt: Original John Bull Bulldog Blend Light. Paines Malt Ltd, http://www.homebrewers.com/ Yeast sprinkles (optional): Fleischmann’s Active Dry Yeast for Baking. Fleischmann Yeast (these are the yeast sprinkles used on the cornmeal food once the food has jellied and solidified; they can be purchased at any major grocery store). Cornmeal: Degermed Yellow Corn Product. Iowa Corn Processors (found in local grocery stores). Baby cereal: Nestle Carnation Baby Cereal with Formula Rice and Banana Flavor and FOS (FOS is a chemical used for the easy digestion of food for newborns; cereal can be purchased at any major grocery store).

Cooking equipment Sundries: Cottonballs buzzplugs for glass and plastic vials paper lids for glass and plastic bottles glass vials glass bottles plastic vials plastic bottles

• • • • • • •

can all be purchased through Genessee Scientific (toll free 1-800-789-5550), www.geneseesci.com: Steam kettle: Groen Model TDB/7/TA/2-40 (toll free 1-888-371-6048), www.groen.com Mixer: Groen Model TA2 (toll free 1-888-371-6048), www.groen.com Fly vial filler: Fisher Scientific (1-800-766-7000), www.fishersci.com Also available by catalog order as Applied scientific fly vial filler AS-770 (narrow diameter) or AS-780 (wide diameter).

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Chemicals (methylparaben, benzyl benzoate, propionic acid): Sigma, http://www.sigmaaldrich.com/suite7/Local/SA_Splash.html Lab supplies and ingredients: The Drosophila Company, http://drosophila.herpetology.com/ Forceps (Dumont high precision): Ted Pella, http://www.tedpella.com/tweezers.htm#anchor490590 Plugs, cheesecloth, media ingredients: SciMart (Drosophila Catalogue), http://www.scimart.com/ Instant culture medium: www.carolina.com Additional culture medium recipes: http://flystocks.bio.indiana.edu/media-recipes.htm

Flies for sale Bloomington Stock Center (United States), http://flystocks.bio.indiana.edu/ Carolina Biological Supply (United States), www.carolina.com Drosophila Genetic Resource Center (Japan), http://www.shigen.nig.ac.jp/fly/nigfly/ Ehime Species Stock Center of Japan, http://kyotofly.kit.jp/ehime/ The Drosophila Company (United States), http://drosophila.herpetology.com/ Szeged Stock Center (Hungary), http://gen.bio.u-szeged.hu/servlet/jate.genetics. servlet.Welcome Tucson Stock Center (United States), http://stockcenter.arl.arizona.edu/

Species sequencing and BAC library projects Drosophila pseudoobscura genome project (Baylor College of Medicine), http://www.hgsc.bcm.tmc.edu/projects/drosophila/ Drosophila species genome sequencing projects, http://genome.gov/page.cfm?pageID
10002154 Drosophila species BAC library projects, http://www.genome.gov/10001852

Other Drosophila resources and databases Drosophila Genomics Resource Center (United States), http://dgrc.cgb.indiana.edu/ Drosophila Information Service, http://www.ou.edu/journals/dis/ Drosophila Polymorphism Database (Spain), http://bioinformatica.uab.es/dpdb/ Flybase, http://flybase.net/ Flybrain, http://flybrain.neurobio.arizona.edu/ TaxDros, http://taxodros.unizh.ch/

Index

Abdomen, 65, 66, 75–8 Acrostichal setulae, 69, 72, 86 Adh, 26, 27 Aedeagus (penis), 76 Africa, 160, 169, 176, 177 Age at reproductive maturity, 194–6, 201, 204, 228, 238 culture method adjustment rapidly maturing adults, 210–11 slowly maturing adults, 211–12 Hawaiian Drosophila, 213 setting up crosses, 211 Age-dependent metabolic changes, 232 Allozyme investigations, 152 Alpha-amanitin, 34, 225 Amanita mushroom bait, 147 Amazon Basin, 155 amd, 14 Amiota, 65 phylogenetic relationships, 10 Anesthetizing methods, 212–13 Antennae, 65, 68 third segment chaetotaxy, 65 Arista, 65 Asia, 158, 159, 166, 169, 170, 174, 175, 178, 179, 180 Aspirators, 145–6 removal of parent flies, 206, 207 virgin flies collection, 209, 210, 211, 236 Ateledrosophila, 37 Australia, 174, 175, 178, 180 Axenic culture, 237 BAC library projects, 245 Bacterial contamination, 183, 236–7 Baits: containers, 148–9 Hawaiian Drosophilidae collecting, 150 placement, 148–9 preparation, 147–8 Banana bait baby food bait, 150 preparation, 147

Banana food, 221 Banana-opuntia medium, 221 recipe, 222–3 Barometric pressure, 238 Basiphalus, 76 Beating bags, 151 Behavioural studies diurnal factors, 228–9 fly storage density, 232 social factors, 232 Benzyl benzoate, 237 Bolivia, 161, 172 Borneo, 169, 175, 178, 179 Bottle baits, 149 Brazil, 155, 167, 172 Bunostoma, key to species, 88 Cactophilic species, 203, 206 dietary considerations, 221, 225 natural substrates, 149 oviposition stimulation, 239 Cactus bait preparation, 147–8 Cameroon, 176 Camilidae, 3 Canary Islands, 173 Carbon dioxide anesthesia, 212, 213 Caribbean, 163, 167, 168, 171, 172 Carina, 66 Central America, 155, 156, 157, 159, 161, 162, 163, 164, 165, 166, 167, 168, 170, 171, 172, 173 Cerci, 76 China, 166, 174, 178, 179 Choice tests for sexual isolation, 228, 229, 231–2 Chymomyza, 4, 44 key to species, 90 morphological characters, 66, 72 phylogenetic relationships, 11–13 species groups, 12 Chymomyza aldrichii, 12, 13 Chymomyza amoena, 68, 89, 90 Chymomyza costata, 12

248

Index Chymomyza fuscimana, 12 Chymomyza obscuta, 12 Chymomyza procnemis, 12 Ciliated tarsus species group, 38 Cladistics, 5–6 Cleaning, 236 Clermontia, 213, 239 Climate change, 154 Collecting methods, 145–52 baiting, 147–9 Hawaiian Drosophilidae, 150–1 larvae from decaying material, 149–50 from host plant material, 151 natural substrates, 149–50 seasonal factors, 149 tools/equipment, 145–7 Collecting permits, 151–2 Collecting tools/equipment, 145–7 aspirators, 145–6 equipment list, 145 holding vials, 145–6, 147 nets, 147 Colombia, 171 Compound eye, 69 Copulation duration, 230–2 Cornmeal medium, 221–2 Costa Rica, 171 Courtship vigor, 232 experiments, 228 Coxa, 72 Crowding, 204–5, 206 influence on behviour, 232 in storage vials, 211 Cryopreserved wild-caught Drosophila, 183, 184, 185 Cuba, 171 Culture medium, 201 antibiotic additives, 237 moisture content, 235 pH reduction, 237 recipes, 216–24 special requirements, 225 supply sources, 243–5 transport of wild flies, 152 Culture methods, 200–1 age of adult flies, 203, 204 aging in food vials, 204 anesthetizing methods, 212–13 collected larvae, 149–50, 151

dietary considerations, 215, 221 food types, 201, 203 recipes, 216–24 documentation, 201 ecological considerations, 201 embryo culture, 237 fly density, 204–5, 206 food surface characteristics, 201 Hawaiian Drosophilidae, 151, 213–14 life history considerations, 201 oviposition maximization, 206 perching/pupation sites provision (use of tissue), 201, 202 problems, 235–9 dry food, 235 loose food, 236 mating failure, 238 oviposition failure, 239 runny food, 236 removing flies, 201, 203, 204 reproductive potential influence, 205–6 setting up crosses, 211 setting up cultures, 201 species with older ages at sexual maturity, 211–12 species with rapidly maturing adults, 209–11 virgin flies: collection, 208–9 starting culture, 206–8 storage in vials, 211, 212 Curtonotidae, 3 Cut wood, 13 Databases, 245 Dcd, 14 Dettopsomyia, 14, 16, 55 Development time, 189–94, 201, 228 Diastatidae, 3 Dietary considerations, 215–25 recipes, 216–24 special requirements, 225 see also Culture medium Dissection techniques, 85 Distiphallus, 76 Distribution, 154–80 data sources, 154–5 maps, 155–80 sampling bias, 155 Diurnal activity patterns, 228–9, 238

Index

249 DNA investigations, 152, 184–5 Dorsilopha, 13–14, 28 key to species, 96 phylogenetic relationships, 13–14 Dorsocentral setae, 69 Dorsolateral plate, 75 Drosophila endemic Hawaiian species see Hawaiian Drosophilidae key to species, 85–142 morphological characters, 66, 69, 72 phylogenetic relationships, 13–41 cladistics, 13, 16 molecular analysis, 14, 16 subgenera, 13 see also Dorsilopha; Drosophila subgenus Drosophila; Sophophora wild-caught see Wild Drosophila Drosophila acanthoptera, 20, 130, 131, 159, 229, 230 Drosophila acutilabella, 32 Drosophila adiastola, 40, 75 Drosophila adunca, 68 Drosophila aenicta, 78 Drosophila afer, 158 Drosophila affinis, 99, 173, 201 Drosophila affinis (species group), 49, 51 distribution, 173 Drosophila albomarginata, 156 Drosophila albomicans, 34, 169 Drosophila albostrigata, 34 Drosophila aldrichi, 163 Drosophila alexanderi, 168 Drosophila algonquin, 98, 173 Drosophila altiplanica, 158 Drosophila ambigua, 173 Drosophila americana, 26, 167 Drosophila amlerkotliana, 174 Drosophila ananassae, 174, 189 morphological characters, 73 Drosophila ananassae (species subgroup), 49 distribution, 174 Drosophila anceps, 162 Drosophila angularis, 170 Drosophila annularis, 156 Drosophila annulimana, 155 morphological characters, 69 Drosophila annulimana (species group), 19 distribution, 155 key to species, 132–4

phylogenetic relationships, 14, 16, 17 species, 17 Drosophila antillea, 32 Drosophila antopocerus (species group), 37, 38, 65, 68 Drosophila aracatacas, 128, 133, 155 Drosophila araucana, 168 Drosophila arizonae, 163 Drosophila asper, 159 Drosophila athabasca, 173 Drosophila attigua, 40 Drosophila auraria, 178 Drosophila australosaltans, 51 Drosophila austrosaltans, 103 Drosophila azteca, 173 Drosophila baimaii, 117, 178 Drosophila barbarae, 123, 178 Drosophila bedichecki, 31, 167 Drosophila belladunni, 167 Drosophila biarmipes, 49, 106, 174 Drosophila biauraria, 179 Drosophila bifasciata, 173 Drosophila bifurca, 161, 204 Drosophila bifurcata, culture methods, 211–12 Drosophila bilimbata, 34 Drosophila bipectinata, 116, 174 Drosophila birchii, 125, 178 Drosophila bisetata, 158 Drosophila boliviana, 157 Drosophila borealis, 26, 167 Drosophila boroborema, 165 Drosophila brachynephros, 35, 170 Drosophila breuerae, 155 Drosophila briegeri, 157 Drosophila brncici, 158 Drosophila bromeliae, 128, 156 Drosophila bromeliae (species group) distribution, 156 key to species, 127 phylogenetic relationships, 14, 16, 17, 18 species, 17 Drosophila bromelioides, 156 Drosophila bromelioides (species group), 17 Drosophila busckii, 13, 96, 201 Drosophila buzzatii, 22 Drosophila calloptera, 75, 166 Drosophila calloptera (species group) key to species, 137–8 phylogenetic relationships, 28

250

Index Drosophila calloptera (species group) (Contd) species, 30 wing morphology, 74 Drosophila camargoi, 157 Drosophila canalinea, 156 Drosophila canalinea (species group), 14 distribution, 156 key to species, 132 phylogenetic relationships, 18–19 species, 17 Drosophila canalinoides, 156 Drosophila capricorni, 172 Drosophila carbonaria, 149, 215 Drosophila carbonaria (species group) distribution, 156 phylogenetic relationships, 14, 19 species, 17 Drosophila carcinophila, 161, 215 Drosophila cardini (species group) chromosome phylogeny, 32 distribution, 167, 168 key to species, 141 morphological characters, 75 phylogenetic relationships, 28, 31–2, 58 species, 30 Drosophila cardinoides, 31, 167 Drosophila carsoni (species group) distribution, 156 phylogenetic relationships, 14, 19 species, 17 Drosophila cheda, 159 Drosophila coffeata (species group), 14 distribution, 157 phylogenetic relationships, 19 species, 17 Drosophila colorata, 22, 166 Drosophila cordata, 51, 52, 171 Drosophila coroica, 161 Drosophila curvispina, 35 Drosophila cyrtoloma, 40 Drosophila dacunhai, 171 Drosophila deflecta, 35, 170 Drosophila denticulata, 105 Drosophila dentissima, 44 Drosophila desertorum, 164 Drosophila diplacantha, 177 Drosophila dispar, 44 Drosophila dreyfusi, 157 Drosophila dreyfusi (species group), 14 distribution, 157

key to species, 132 phylogenetic relationships, 19 species, 17 Drosophila dunni, 28, 32, 168 Drosophila elegans, 108 Drosophila elegans (species subgroup), 49 distribution, 180 Drosophila elliptica, 51, 52, 53, 172 Drosophila emarginata, 52, 54, 103, 172 Drosophila eohydei, 161 Drosophila equinoxalis, 54, 172 Drosophila ercepeae, 174 Drosophila erecta, 176 Drosophila eremophila, 22, 163 Drosophila eugracilis, 105 Drosophila eugracilis (species subgroup), 49 distribution, 180 Drosophila euronotus, 20 Drosophila ezoana, 166 Drosophila falleni, 34, 35, 170, 225 Drosophila fasciola, 161 Drosophila fasciola (species subgroup), distribution, 161 Drosophila ficusphila, 111 Drosophila ficusphila (species subgroup), distribution, 180 Drosophila fima, 44, 45 Drosophila flavomontana, 167 Drosophila formosana, 169 Drosophila fulvalineata, 161 Drosophila fulvimacula, 162 Drosophila fulvimaculoides, 162 Drosophila fumipennis, 172 Drosophila funebris, 139 Drosophila funebris (species group), 4, 14 key to species, 138 Drosophila fungiperda, 40 Drosophila fuscipennis, 157 Drosophila fuscolineata, 157 Drosophila gaucha, 158 Drosophila gibberosa, 133, 155 Drosophila glabriapex, 40 Drosophila greeni, 124, 177 Drosophila grimshawi, 40 Drosophila griseolineata, 33, 168 Drosophila guanche, 173 Drosophila guaraja, 168 Drosophila guaramunu (species group), 32–3 Drosophila guarani, 168

Index

251 Drosophila guarani (species group) distribution, 168 phylogenetic relationships, 28, 32–3 species, 30 Drosophila guarau, 168 Drosophila guayllambambae, 161 Drosophila guttifera, 34, 35, 167, 225 Drosophila haleakalae (species group), 37, 38, 40, 43, 58, 150 Drosophila hamatofila, 164 Drosophila heteroneura, 40, 65, 75 Drosophila hexastigma, 165 Drosophila hirtipes, 160 Drosophila huilliche, 168 Drosophila hydei, 161, 196, 201, 236 female remating frequency, 229, 230 Drosophila hydei (species subgroup) distribution, 161 phylogenetic relationships, 22, 25 Drosophila hypocausta, 169 Drosophila immigrans, 140, 201, 205 morphological characters, 69, 72, 73, 76, 86, 90, 97 Drosophila immigrans (species group) distribution, 169 key to species, 138–40 phylogenetic relationships, 28, 33–4 species, 30–1 Drosophila immigrans-Hirtodrosophila radiation, 4, 5, 27–8 Drosophila immigrans-tripunctata radiation, 14, 27–35 key to species, 137–42 phylogenetic relationships, 29 species groups, 30–1 Drosophila inca, 161 Drosophila inca (species subgroup), distribution, 161 Drosophila innubila, 35, 170, 225 Drosophila insularis, 54, 172 Drosophila jambulina, 121, 179 Drosophila kallima, 166 Drosophila kanapiae, 116 Drosophila kanekoi, 166, 204 culture methods, 211–12 Drosophila kepulauana, 169 Drosophila kikkawai, 179 Drosophila koepferae, 165 Drosophila kohokoa, 169 Drosophila krugi, 157

Drosophila kuntzei, 35 Drosophila lacertosa, 22, 159 Drosophila lacicola, 167 Drosophila lacteicornis, 120, 178 Drosophila leonis, 162 Drosophila limbata, 35 Drosophila limbinervis, 168 Drosophila limensis, 162 Drosophila linearepleta, 162 Drosophila lini, 118, 179 Drosophila longicornis, 164 Drosophila longipedis, 40 Drosophila longiserrata, 158 Drosophila lowei, 173 Drosophila lucipennis, 49, 174 Drosophila lugubripennis, 157 Drosophila lusaltans, 51, 101, 102, 171 Drosophila lutea, 112 Drosophila lutescens, 175 Drosophila macroptera, 28 Drosophila macrospina, 139 Drosophila magniquinaria, 167 Drosophila mainlandi, 164 Drosophila malerkotliana, 115 Drosophila maracaya, 166 Drosophila martensis, 22, 165 Drosophila mathisi, 164 Drosophila mauritiana, 176 Drosophila mayaguana, 163 Drosophila mayri, 122, 178 Drosophila mediobandes, 170 Drosophila mediodiffusa, 171 Drosophila mediopictoides, 171 Drosophila mediostriata, 171 Drosophila melanica, 157 Drosophila melanica (species group) distribution, 157, 158 key to species, 130–1 phylogenetic relationships, 14, 19–20 species, 17 Drosophila melanissima, 157 Drosophila melanogaster, 104, 107, 176, 182, 196, 201, 203, 230, 232 culture methods, 209–11 diurnal influences on mating activity, 228–9 morphological characters, 69, 72, 73, 76 wild fly collecting from natural substrates, 149

252

Index Drosophila melanogaster (species group) distribution, 174, 175, 176, 177, 178, 179, 180 key to species, 104–26 phylogenetic relationships, 44, 45, 46, 49, 50, 51, 58 species, 48 Drosophila melanura, 20 Drosophila mercatorum, 161 Drosophila mercatorum (species subgroup), 22 distribution, 161 Drosophila meridiana, 22, 162 Drosophila meridionalis, 162 Drosophila mesophragmatica, 158 Drosophila mesophragmatica (species group), 14 distribution, 158 key to species, 132 phylogenetic relationships, 20, 21 species, 17 Drosophila mettleri, 163 Drosophila microlabis, 51 Drosophila micromelanica, 20, 157 Drosophila micromettleri, 163 Drosophila milleri, 171 Drosophila mimetica, 174 Drosophila miranda, 173 Drosophila mojavensis, 22, 163 Drosophila mojuoides, 161 Drosophila montana, 167 Drosophila montium (species subgroup), 49 distribution, 177, 178, 179 Drosophila moriwakii, 22, 159 Drosophila mulleri, 196, 230 Drosophila mulleri (species subgroup), 22 distribution, 162, 163, 164, 165 Drosophila munda, 35, 170, 225 Drosophila nannoptera, 129, 159, 196 Drosophila nannoptera (species group) distribution, 159 key to species, 128–30 phylogenetic relationships, 14, 20–1 species, 17 Drosophila narragansett, 173 Drosophila nasuta, 33–4, 169 Drosophila navojoa, 163 Drosophila nebulosa, 54, 172 Drosophila neocardini, 32, 167 Drosophila neocordata, 52

Drosophila neohydei, 161 Drosophila neohypocausta, 169 Drosophila neomorpha, 31, 168 Drosophila neonasuta, 34 Drosophila neookadai, 159 Drosophila neopicta, 40 Drosophila neorepleta, 162 Drosophila neotestacea, 35, 167, 225 Drosophila nigella, 40 Drosophila nigra, 40 Drosophila nigricruria, 164 Drosophila nigrodunni, 32 Drosophila nigrohydei, 161 Drosophila nigromaculata, 35, 170, 225 Drosophila nigromelanica, 20 Drosophila nigrosaltans, 101 Drosophila nigrospiracula, 162 Drosophila nikananu, 98, 106, 107, 177 Drosophila niveifrons, 34 Drosophila novamexicana, 167, 201 Drosophila novemmarista, 161 Drosophila nudidrosophila (species group), 38 Drosophila obscura, 72, 173 Drosophila obscura (species group) key to species, 98–100 phylogenetic relationships, 44, 45, 49, 51, 52, 58 species, 48 Drosophila occidentalis, 35, 170 Drosophila okadai, 159 Drosophila orena, 176 Drosophila orientacea, 35, 225 Drosophila orientalis, 35 Drosophila orkui, 158 Drosophila ornatipennis, 137, 166 Drosophila orosa, 121, 179 Drosophila pachea, 20, 159, 203, 204, 206, 225 Drosophila pachuca, 164 Drosophila pallidipennis centralis, 34 Drosophila pallidipennis pallidipennis, 34 Drosophila pallidipennis (species group) distribution, 170 key to species, 140 phylogenetic relationships, 34 species, 31 Drosophila pallidosa, 111, 174 Drosophila palustris, 34, 35, 167 Drosophila paraaensis, 161 Drosophila parabipectinata, 174

Index

253 Drosophila paracanalinea, 156 Drosophila paraguttata, 161 Drosophila parakuntzei, 170 Drosophila paralutea, 115, 175 Drosophila paramelanica, 20, 157 Drosophila pararubida, 169 Drosophila parasaltans, 53 Drosophila parisiena, 163 Drosophila parthenogenetica, 31, 168 Drosophila parvula, 117, 179 Drosophila paulistorum, 54, 172 Drosophila pavani, 158 Drosophila pavlovskiana, 54, 172 Drosophila pegasa, 165 Drosophila peninsularis, 161 Drosophila pennae, 125 Drosophila persimilis, 173, 182, 229 Drosophila peruensis, 168 Drosophila peruviana (species group) distribution, 159 phylogenetic relationships, 14, 21 species, 17 Drosophila phaeopleura, 99, 109, 110, 174 Drosophila phalerata, 34, 35, 225 Drosophila picticornis, 40 Drosophila pictilis, 161 Drosophila pictura, 161 Drosophila planitibia, 40 wing morphology, 74 Drosophila planitibia (species group), 40, 42 Drosophila polita, 40 Drosophila polychaeta, 127 Drosophila polychaeta (species group) distribution, 159, 160 key to species, 127 phylogenetic relationships, 14, 16, 21–2 species, 17 Drosophila polymorpha, 32, 168 Drosophila primaeva, 40 Drosophila procardinoides, 31, 167 Drosophila promeridiana, 162 Drosophila propachuca, 164 Drosophila prosaltans, 51, 54, 102, 172 Drosophila prostipennis, 113, 114, 175 Drosophila pseudoananassae, 174 Drosophila pseudoobscura, 173, 182, 229, 238 Drosophila pseudoobscura (species group), 51 distribution, 173

Drosophila pseudosordidula, 22, 159 Drosophila pseudotakahashii, 175 Drosophila pulaua, 169 Drosophila pulchella, 171 Drosophila pulchrella, 112, 174 Drosophila punjabiensis, 120, 179 Drosophila putrida, 35, 138, 167, 225 Drosophila quadraria, 179 Drosophila quinaria, 167, 203 Drosophila quinaria (species group), 58, 225 distribution, 167, 170 key to species, 141–2 phylogenetic relationships, 28, 34–5 species, 31 Drosophila radiation, 4 Drosophila recens, 34, 35, 167, 225 Drosophila rectangularis, 172 Drosophila rellima, 170 Drosophila repleta, 162 morphological characters, 75 Drosophila repleta (species group), 14, 58, 221 chromosome phylogeny, 22, 23 distribution, 161, 162, 163, 164 key to species, 134–7 molecular phylogeny, 22, 24, 25 phylogenetic relationships, 22 species, 17–18 Drosophila repletoides, 14, 25, 166 Drosophila rhopaloa, 49 Drosophila richardsoni, 163 Drosophila ritae, 164 Drosophila robusta, 166 Drosophila robusta (species group) chromosome phylogeny, 25 distribution, 159, 166 key to species, 132 phylogenetic relationships, 14, 22 species, 18 Drosophila rubifrons, 28 Drosophila rufa, 119, 179 Drosophila saltans, 101, 171 Drosophila saltans, (species group) distribution, 171, 172 key to species, 100–4 phylogenetic relationships, 44, 45, 51–4, 58 species, 48 Drosophila santomea, 176 Drosophila seguyi, 123, 177

254

Index Drosophila septentriosaltans, 171 Drosophila serido, 165 Drosophila serrata, 125, 178 Drosophila seychellia, 176, 205, 239 Drosophila signata, 169 Drosophila simulans, 97, 176, 182, 201 culture methods, 209–11 morphological characters, 69 wild fly collecting from natural substrates, 149 Drosophila sonorae, 164 Drosophila sordidula, 22, 159 Drosophila spenceri, 165 Drosophila stalkeri, 22, 163 Drosophila starmeri, 165 Drosophila straubae, 163 Drosophila sturtevanti, 51, 52, 53, 54, 100 Drosophila subabdia, 32 Drosophila subgenus Drosophila, 13 key to species, 127–42 phylogenetic relationships, 14–41 Drosophila subobscura, 51, 173, 229 Drosophila suboccidentalis, 35, 170, 225 Drosophila subpalustris, 34, 35 Drosophila subquinaria, 35, 225 Drosophila subsaltans, 51, 52 Drosophila subsilvestris, 173 Drosophila sucinea, 172 Drosophila suffusca, 170 Drosophila sulfurigaster, 34, 169 Drosophila suzukii (species subgroup), 49 distribution, 174 Drosophila takadai, 170 Drosophila takahashii, 49, 104, 114, 175 Drosophila talamanca, 134 Drosophila teissieri, 176 Drosophila tenebrosa, 170, 225 Drosophila testacea, 203, 225 morphological characters, 72 Drosophila testacea (species group), 225 distribution, 167, 170 key to species, 138 phylogenetic relationships, 28, 35, 58 species, 31 Drosophila texana, 26 Drosophila tolteca, 99, 173 Drosophila transversa, 35, 225 Drosophila triauraria, 124, 179 Drosophila trilutea, 113

Drosophila tripunctata, 171, 203 morphological characters, 71, 75 Drosophila tripunctata radiation, 5 Drosophila tripunctata (species group) distribution, 171 key to species, 142 phylogenetic relationships, 28, 35 species, 31 Drosophila tristis, 173 Drosophila tropicalis, 54, 172 Drosophila tsacasi, 122, 177 Drosophila tsigana, 158 Drosophila tumiditarsus (species group) distribution, 166 phylogenetic relationships, 14, 16, 25 species, 18 Drosophila unipunctata, 171 Drosophila uniseta, 165 Drosophila unispina, 35, 170 Drosophila varians, 110, 174 Drosophila venezolana, 165 Drosophila venusta, 40 Drosophila virilis, 208 Drosophila virilis (species group) distribution, 166, 167 key to species, 131–2 phylogenetic relationships, 14. 25–7, 58 species, 18 Drosophila virilis-repleta radiation, 4, 5, 14–27 key to species, 127–37 species groups, 17–18 Drosophila vulcana, 119, 177 Drosophila wassermani, 20, 129, 159 Drosophila wheeleri, 163 Drosophila willistoni, 172 morphological characters, 72 Drosophila willistoni (species group) distribution, 172 key to species, 126–7 phylogenetic relationships, 44, 45, 54, 58 species, 49 Drosophila wingei, 157 Drosophila yakuba, 176 Drosophilidae, 3 Hawaiian endemic see Hawaiian Drosophilidae origins, 3–4 phylogenetic relationships, 4–57 cladistics, 3, 5–6

Index

255 Drosophilinae, 11–57 molecular studies, 6–7, 8 radiations, 4–5, 13, 14 Steganinae, 10, 11 taxonomic relationships, 7, 9 Drosophilinae, 10, 16 phylogenetic relationships, 11–57 genus Chymomyza, 11–13 genus Drosophila, 13–41 genus Hirtodrosophila, 54–5 genus Liodrosophila, 55, 56 genus Samoaia, 55, 56 genus Scaptodrosophila, 57 genus Sophophora, 43–54 genus Zaprionus, 57 Ecological aspects, 196–7 Egg production, 196 Egg to adult development time, 189–94, 228 El Salvador, 161 Electrophortica, 4 Embryos collection/culture, 237 Engiscaptomyza, 43 Epandrial lobes, 76 Epandrium, 76, 77 Ephydridae, 3 Ephydroidea, 3 phylogenetic relationships, 3, 4 Equipment list for collecting, 145 sources, 243 Ethanol-preserved wild-caught Drosophila, 183, 184, 185 Etherization, 212–13 Europe, 158 Eyes, 69 Face, 66 Fecundity, 196 Female choice test, 231 Female remating frequency, 229–30 Femoral spines, 72, 73 Femoral tubercles, 72, 74 Femur, 72 Fertility, 196 Field notes, 182, 183, 184 Field stations, 151–2 Field work equipment sources, 243 Fiji, 174 Flies for sale, 245

Flower breeders, 215 natural substrates, 149 Fly density see Crowding Flybase, 65 Food vials, transport of wild flies, 152 Foreleg modifications, 73 Fronto-orbital plates, 66 Fronto-orbital setae, 66, 68, 69 Fruit bait preparation, 147 natural substrates, 149 Fungal inhibitors, 236–7 Gena (cheeks), 69 Genetic variability in living cultures, 182–3 Genitalia, 76, 78 Genome sequencing, 245 Gitona, 65 Glass containers, 200 Gonopods, 76 Grimshawomyia, 38, 42 Haiti, 171 Halteres (ballancers), 72 Harvesting methods from baits, 149 removing flies, 201, 203, 204, 236 Hawaiian Drosophilidae, 6, 14, 28, 35–8 antopocerus species group, 37, 38, 65, 68 ciliated tarsus species group, 38 culture methods, 151, 211, 213–14 development times, 194 haleakalae species group, 37, 38, 40, 43, 58, 150 modified mouthparts species group, 37, 38, 151 modified tarsus species group, 37, 38, 73 morphological characters, 65, 66, 68, 70, 72–3, 74, 75 nudidrosophila species group, 38 oviposition stimulation, 213, 239 phylogenetic relationships, 6, 35–41 molecular analysis, 37, 38, 39 picture wing species group, 37–8, 40, 41, 58, 74, 75, 151 planitibia species group, 40, 42 reproductive maturity, 213 reproductive potential, 197 sand chamber pupation, 194, 213–14

256

Index Hawaiian Drosophilidae (Contd ) transport of wild flies, 151 wild fly collecting, 150–1 Head, 65–9 Hirtodrosophila, 6, 27–8, 36 key to species, 90 Old World radiation, 4, 5, 28 phylogenetic relationships, 54–5 Hirtodrosophila duncani, 55, 91 Holding vials, 145–6, 147 Hawaiian Drosophila, 213 Humeral callus, 69 Humidity Hawaiian Drosophila culture requirements, 213 transport of wild flies, 152 Hydroxy-benzoic acid methyl ester, 237 Hypandrium, 76, 77 Hypercephaly, 65, 67, 68 Idiomyia, 37, 40 Import permits, 152 India, 174, 175, 179 Indonesia, 169 Inner vertical setae, 66 International Organization of Biological Field Stations, 151 Interspecific crosses, 238 Isofemale lines, 182, 183 oviposition stimulation, 183 Jamaica, 167, 171 Japan, 166, 170, 174, 175, 178, 179, 180 Katepisternal setae, 72 Katepisternum, 72 Key to species, 85–142 Korea, 175, 178, 179, 180 Labellum, 66 Land crab commensalism, 215 Larvae collection from decaying material, 149–50 from host plant material, 151 Leg segments, 72 Legs, 72 Life history variables, 189–97, 227–32 age at sexual maturity, 194–6 culture method adjustments, 205 ecological aspects, 196–7

egg to adult development times, 189–94 fecundity, 196 fertility, 196 Hawaiian Drosophila, 213 Liodrosophila, 16 key to species, 91 morphological characters, 66 phylogenetic relationships, 55, 56 Liodrosophila aerea, 55 Lissocephala, 3 Living cultures, 182–3 isofemale lines, 182, 183 multifemale strains, 182, 183 Longevity, 232 Lophocereus schottii (seinita), 206, 225 Lordiphosa, 44 Madagascar, 169 Malaysia, 174, 179 Male choice test, 232 Mating failure, 238 Mauritius Islands, 176 Mesonotum, 69 Mesothorax, 69 Mesquite (Prosopis), 19, 149, 215 Metathorax, 69 Mexico, 155, 159, 167, 171, 172 Micronesia, 175 Mites, 183, 236–7 Modified mouthparts species group, 37, 38, 151 Modified tarsus species group, 37, 38, 73 foreleg modifications, 73 Moisture, transport of wild flies, 152 Mold contamination, 236–7 Morinda, 239 Morphological characters, 65–83 glossary of terms, 78–83 Mouthparts, 66, 70 Multifemale strains, 182, 183 Multiple choice test for sexual isolation, 232 Mushroom, 206, 225 baits placement, 148 baits preparation, 147 natural substrates, 149 “tea” bait, 150 Mycodrosophila, 54, 55 Mycophagous species, 28, 29, 203, 206, 225 culture methods, 211 dietary requirements, 225

Index

257 Natural substrates oviposition stimulation, 239 wild fly collecting methods, 149–50 Nearctic species, 157, 166, 167 Nematodes, 183 Neotanygastrella, 44 Nets, 147, 149 for sweeping rainforest vegetation, 151 New Guinea, 178, 179, 180 North America, 156, 157, 166, 167, 168, 170, 173 Nudidrosophila species group, 37, 38 Oceanic region, 174 Ocellar setae, 66 Oogenesis, 196 Opuntia ficus-indica, 206 cactus bait preparation, 148 Opuntia powder, 221 Oral setae, 69 Orbital setae, 66, 68, 69 Outer vertical setae, 66 Ovariole number, 196, 197, 205 Oviposition failure, 239 Oviposition stimulation, 206, 239 Hawaiian Drosophila, 213 isofemale lines, 183 Ovipositor, 78 Ovisensilla, 78 Palps, 69 Paraguay, 172 Parascaptomyza, 88 key to species, 88 Penicillin, 237 Penis (aedeagus), 76 Peru, 159, 172 Philippines, 169, 174, 178, 179, 180 Phloridosa, 14 Pholadoris, 57 Phylogenetic relationships, 3–58 Drosophilinae, 11–57 genus Chymomyza, 11–13 genus Drosophila, 13–41 genus Hirtodrosophila, 54–5 genus Liodrosophila, 55, 56 genus Samoaia, 55, 56 genus Scaptodrosophila, 57 genus Scaptomyza, 42–3

genus Sophophora, 43–54 genus Zaprionus, 57 Hawaiian Drosophilidae, 6, 35–41 molecular studies, 6–7 Picture wing species group, 37–8, 40, 41, 58 culture, 151 wing morphology, 74, 75 Pilosity, 69 Pinned specimens, 183, 184 Plastic containers, 200 Plumose arista, 65 Point-mounting, wild-caught Drosophila, 184 Postocellar setae, 66 Prensisetae, 76 Prescutellar setae, 69 Preservation of wild Drosophila, 183–4 collection data labels, 184 long-term storage, 185 Presutural setae, 72 Proclinate orbital setae, 66 Prosopis (mesquite), 19, 149, 215 Prothorax, 69 Pubescent arista, 65 Puerto Rico, 171 Rearing see Culture methods Reclinate orbital setae, 66 Removing flies, 201, 203, 204, 236 Reproductive maturity, 194–6, 203, 204, 228 see also Age at reproductive maturity Reproductive potential, 196, 201 culture density influence, 205–6 Reunion Island, 174 Saguro–potato medium, 224 St Vincent Islands, 171 Samoaia, 28 key to species, 89–90 phylogenetic relationships, 55, 56 Samoaia afiamalu, 55 Samoaia attenuata, 55 Samoaia leonensis, 55, 87 Sand chambers, 194, 213–14 Santomea Islands, 176 Scaptodrosophila, 4, 6 key to species, 94–6 morphological characters, 69 phylogenetic relationships, 57 Scaptodrosophila latifasciaformis, 94 Scaptodrosophila lebanonensis, 94

258

Index Scaptodrosophila radiation, 4 Scaptomyza, 14, 28, 35, 36, 38, 54 key to species, 86–9 morphological characters, 76 phylogenetic relationships, 42–3 Scaptomyza adusta, 86 Scaptomyza anomala, 87 Scaptomyza palmae, 89 Scutellar setae, 72 Scutellum, 72 Seinita (Lophocereus schottii), 206, 225 Setae, 66, 68, 69 abdomen, 76 thorax, 69, 71, 72 Sex combs, 72, 73 Sexual isolation studies, 228, 229, 231, 232, 238 Sexual selection studies, 229 Seychelles Islands, 169, 176 Siphlodora, 16, 28 Social facilitation of mating, 238 Social interactions, 232 Sophophora, 4, 13 key to species, 96–142 phylogenetic relationships, 43–54 species groups, 48–9 Sophophoran radiation, 4 Sordophila, 14 South America, 155, 156, 157, 158, 159, 161, 162, 163, 164, 165, 167, 168, 170, 171, 172, 173 Southeast Asia, 169, 174, 175, 178, 179 Sperm production, 196 storage confirmation, 239 utilization studies, 229–30 Sphaerogastrella, 16, 55 Sri Lanka, 174 Standard banana-opuntia medium, 222–3 Standard cornmeal medium, 221–2 Stegana coleoptrata, 74 Steganinae, 4, 65 phylogenetic relationships, 10, 11 Steganinae radiation, 4 Sternites (ventral plates), 75, 76 Sterno index, 72 Sternopleuron, 72 Storage conditions, 185 Storage vials, 211, 212 Streptomycin, 237

Styloptera, 55 Subvibrissal setae, 69 Supply sources, 243–5 Surstyli, 76 Sweeping techniques, Hawaiian Drosophilidae collection, 150–1 Taiwan, 169, 174, 175, 179 Tarsal segments, 72 Tarsus, 72 Taxodros, 85 Temperature conditions development time influence, 189 Hawaiian Drosophila culture, 213 transport of wild flies, 151, 152 Tergites, 75, 76 Terminalia, 78 Territorial behavior, 65, 68 Thorax, 65, 66, 69, 71–5 Tibia, 72 Timing issues (experimental design), 227–32 age effects, 232 copulation duration, 230–2 diurnal factors, 228–9 egg to adult development time, 228 female remating frequency, 229–30 sexual maturation, 228 Titanochaeta, 38, 42 Tools, 145–7 cleaning, 236 Transport of wild flies, 152 Hawaiian Drosophilidae, 151 import permits, 152 Trinidad, 161, 167 Trochanter, 72 Tubercles, 72, 74 United States National Parks and Monuments, 151 Ventral epandrial lobes, 76 Ventral plate (sternite), 75 Vertical setae, 66 Vibrissae, 69 Virgin flies: collection, 208–9 aspirator use, 236 from species with older ages at sexual maturity, 211–12

Index

259 from species with rapidly maturing adults, 210–11 setting up culture, 206–8 multiple broods, 206, 207, 208 pupation sites provision (use of tissue), 206 removal of parents, 206, 207 storage in vials, 211 Voucher material, 183–5 storage, 185 Wheeler–Clayton medium, 213, 214 recipe, 223–4 Wild Drosophila: collecting methods, 145–52 distribution, 154–80 documentation, 182, 183 handling specimens, 182–5

living cultures, 182–3 microbes/mites/nematodes, 183 preservation, 183–5 long-term storage, 185 voucher material, 183–5 Wings, 73–4 Xdh, 53, 54 Zaire, 177 Zaprionus, 14, 28 key to species, 93–4 morphological characters, 72, 74 phylogenetic relationships, 57 Zaprionus ghesquierei, 91, 93 Zaprionus sepsoides, 74, 92, 93 Zaprionus tuberculatus, 72, 92 Zygothrica, 55, 66