Metamorphosis of the central nervous system of Drosophila melanogaster Meigen (Diptera: Drosophilidae) during pupation*
Kusum Singh and R Naresh Singh
Molecular Biology Unit, Tata
Institute of Fundamental Research, Homi Bhabha Road,
Navy Nagar, Colaba,
Bombay 400 005, India
We studied the metamorphosis of the central nervous system (CNS) and neighbouring muscles of Drosophila melanogaster (Diptera: Drosophilidae) during pupation by transmission electron microscopy (TEM). The age of white pupa was assumed to be 0 h and the process of metamorphosis was monitored, onward between 6 and 96 h at 25°C. The profiles in the neuropil showed degeneration at 6 h and its extent increased by 12 h. The presence of glycogen in some of these profiles indicated their larval character. Between 1218 h, the neuronal profiles became separated from one another, the intervening space was filled with extracellular fluid, and some of the larval synapses degenerated. Synaptic vesicles started reappearing around 18 h and synapses were detectable by 24 h. Neuronal processes compactly filled the neuropil by 65 h and the maturation of synapses continued until 86 h. The degeneration of profiles in the neuropil was found to be bimodal, peaking at 12 and 42 h, and that of cortical cells was unimodal with a peak at 42 h. The number of neuronal profiles increased with the development time, indicating that more branching of neuronal profiles occurs in neuropils as the metamorphosis progresses. Average number of synapses per unit area (or volume) is minimum at 18 h and maximum at 72 h, when the average number of synapse per axon profile is 0·54. Because 2 axon profiles share one synapse, a value close to 0·5 for monad synapses shows that, on an average, each axon profile at least makes one synapse at this stage of development. Subsequently, there is more than 75% of reduction in the number of synapses during 73 and 78 h.
In muscles, vacuoles suggesting histolysis appeared by 6 h. Their ultrastructure became deranged between
1218 h and myoblasts were found to be present since 8 h. Except for a few muscles in the thorax, such as larval oblique muscles and pharyngeal muscles, most of the muscles in the head and thorax lost all the ultrastructural details and histolyzed by 18 h. Around 38 h, imaginal muscles were detectable, and well-developed muscles were found by 55 h. However, myofibrils continued to be added laterally to the preformed muscles even at 96 h.
Electron-dense mitochondria (EDMITs) were found in the neuropil, cortex and muscles of pupa, along with mitochondria of characteristic shape and normal appearance. These EDMITs often occurred in large clusters of more than 100, at times near the surface of the tissue. A few of these were enclosed in vacuoles and were darker than the rest of the EDMITs and normal looking mitochondria. Histochemistry with diaminobenzidine showed the presence of cytochrome c and marker enzyme cytochrome oxidase, both in EDMITs and normal mitochondria. EDMITs were not found to be present in any tissue of the adult Drosophila.
Compared with hemimetabolous insects in which the major features of the building plan of the adult central nervous system (CNS) are laid down during embryogenesis (Malzacher 1968; Edwards 1969; Bate 1976; Doe and Goodman 1985), a holometabolous insect, such as Drosophila, has 2 prominent phases of neurogenesis embryonic (Poulson 1950; Hartenstein and Campos-Ortega 1984; Campos-Ortega and Hartenstein 1985; Canal and Ferrus 1986; Hartenstein et al 1987; Jacob and Goodman 1989a, b), and postembryonic (Bodenstein 1950; Lawrence 1973). Further, in postembryonic period, 2 distinct phases of development of CNS exist (i) larval (Sonnenblick 1950; White and Kankel 1978; Thomas et al 1984; Truman and Bate 1988; Hartenstein 1988; Singh et al 1989; Prokop and Technau 1991), and (ii) post-larval, where metamorphosis of the CNS occurs during pupation leading to early adult stage of the fly (Hertweck 1931; Robertson 1936; Power 1943, 1948, 1952; Zalokar 1947; Bodenstein 1950; Kankel et al 1980; Technau and Heisenberg 1982; Truman 1990). Extensive studies have been done on the development of eyes (Lees and Waddington 1942; Power 1943; Meinertzhagen 1973; Kankel and Hall 1976; Ready et al 1976; Campos-Ortega and Hofbauer 1977; Kankel et al 1980; Zipursky et al 1984; Tomlinson and Ready 1987), and the peripheral nervous system of Drosophila (Bate 1978; Murray et al 1984; Hartenstein 1988; Singh et al 1989; Ghysen and Dambly-Chaudière 1990; Giangrande and Palka 1990) using light and electron microscopy. Except for the studies of White and Kankel (1978), Technau and Heisenberg (1982), and Truman (1990) the metamorphosis of the CNS in Drosophila during pupation has been rather neglected. Some prominent reasons for the lack of attention to the development of CNS during pupation could be that, right from the beginning, the puparium has a non-transparent exocuticle that is impervious to most of the aqueous fixative solutions. The high tergal pressure inside the puparium, is an added disadvantage, as the inner contents are likely to ooze out through any hole or slit made in the cuticle, for the facilitation of the entry of fixative. Because of these reasons, direct observation of the pupa by light microscope or the preservation of tissue structures for electron microscopy without recourse to injection of fixative through glass microcapillary electrode, becomes difficult.
We have felt the need to follow the development of the CNS and related structures such as glia (Fredieu and Mahowald 1989) and the neighbouring muscles (Sink and Whitington 1991), by transmission electron microscopy (TEM). TEM by virtue of the higher magnification compared with the light microscopy, was in our view better suited for monitoring the process of morphological development of the CNS and the neighbouring muscles during pupation in Drosophila. Although, at higher magnifications, the field of view becomes smaller in TEM this was suitably overcome by the light microscopic examination of 1 m m thick sections for a general view of the morphological development.
At the start of metamorphosis the larval CNS has both larval neurons and arrested imaginal neurons. From the patterns of neurogenesis the thoracic neuropil is likely to be enriched for imaginal neurons, whereas the posterior abdominal neuromere would consist mainly of larval processes (Truman 1990). It would be interesting, if it was possible to discriminate between these two types of processes. However, these processes could not be distinguished from each other by TEM.
The age of the white pupa was assumed to be 0 h. The development process was monitored from 6 to 96 h after white pupa formation (APF) at 4 to 8 (usually 6) hourly intervals at 25°C using mainly TEM. These observations were supplemented with light microscopic examinations.
Clusters of EDMITs, considered to be degenerating mitochondria, were discovered in the nervous tissue and muscles during pupation. These EDMITs were absent in the adult Drosophila tissues.
2. Materials and methods
2.1 Development of pupa at controlled temperature
Five to 6 day-old D. melanogaster Canton special (CS) flies were allowed to lay eggs at 25°C on Drosophila food medium containing yeast. The first 2 lots of eggs during first 2 h were discarded. Subsequent batches of eggs were harvested at hourly intervals on fresh food surface containing yeast. Eggs and the emerged larvae were allowed to grow at 25°C and the time of white pupa formation was regarded as 0 h for the sample. The white pupae were isolated with a soft hair-brush and grown for different lengths of time at 25°C in vials containing a piece of wet filter paper. The temperature during all these operations and especially during development was maintained at 25°C ± 1°C.
2.2 Transmission electron microscopy
At the end of the desired time, individual pupa was submerged under a drop of fixative (Karnovsky 1965). After nicking its anterior and posterior ends, the pupa was transferred to a vial containing approximately 5 ml of fixative and the fixation was done for 6 h at room temperature (RT). The exocuticle of the pupa was removed after 2 h of fixation in samples older than 18 h, whereas in younger samples the exocuticle was removed after full length 6 h of fixation. After several washings with 0·1 M sodium phosphate buffer of pH 7·4 during an hour, the specimens were postfixed with Daltons chrome-osmium tetroxide fixative (Dalton 1955) on ice for 2 h followed by 1 h fixation at RT. By this time, the entire specimen turned dark brown. The specimens were washed with sodium phosphate buffer, dehydrated with increasing concentrations of ethyl alcohol, followed by propylene oxide, infiltrated with Durcupan ACM (Fluka), oriented and embedded in the same resin mixture. Polymerization of the resin mixture was done at 60°C overnight. Silvery ultrathin sections were cut on a LKB Ultrotome III, mounted on Formvar coated grids, and stained with a saturated aqueous solution of uranyl acetate at 60°C for 10 min, washed with water and stained with lead citrate at RT for 5 min (Reynolds 1963). Subsequent examination of sections was done with a Jeol JEM 100 S electron microscope. Approximately 2800 photographs were taken for these studies. Comparative light microscopic observations were made with a Zeiss Universal RS III or a Leitz Orthoplan photomicroscope with plan-apo objectives.
2.3 Histochemistry of cytochrome c and cytochrome oxidase in normal and electron-dense mitochondria
Cytochrome oxidase is one of the several marker enzymes among those present in normal mitochondria (Lehninger 1990). Seligman et al (1968), Reith and Schüler (1972), and Crammer and Moore (1973) clearly demonstrated that diaminobenzidine (DAB) can donate electrons to the oxidized form of cytochrome c in mitochondria. The reduced form is reoxidized by cytochrome oxidase in the presence of oxygen. Thus, the DAB localizes cytochrome oxidase and cytochrome c both inside the mitochondria.
Sixty-five hour-old pupae were cut lengthwise into 2 approximately equal parts. The cut end of right half was immersed for half an hour in a 30% solution of horseradish peroxidase (Sigma type VI) dissolved in water. Subsequently, the tissue was fixed with Karnovskys fixative for 4 h at RT. The tissue was washed several times with 0·2 M sodium cacodylate buffer of pH 7·2. The DAB reaction was carried out on the tissue in 0·05 M TRIS buffer of pH 7·6 according to Nässel (1983) followed by post fixation with 0·5% osmium tetroxide in 0·1 M cacodylate buffer for 1·5 h on ice. Further processing of the tissue and its TEM was done as described above.
TEM observations on the brain, ventral or thoracic ganglia and nearby muscles were recorded.
3.1 Metamorphosis of neuropils
In neuropils of brain and thoracic ganglia, signs of degeneration such as electron-dense profiles, pseudo-myelin figures and abnormal vacuoles, are predominantly present at 6 h and subsequent times until 50 h (figures 1AF, 3A, B). The presence of glycogen until approximately 12 h in some of the profiles in the neuropil, indicates their larval character (Singh et al 1991), (figures 1A, B, arrowhead). In the neuropil, glial profiles by being more electron-dense than the neuronal profiles, are easily distinguishable from each other by TEM. Neuronal profiles start separating from each other by 12 h and the intervening space so created is filled by extracellular fluid and vacuoles (figures 1CE; 3BD, arrow). Vacuoles are significantly present in the neuropil until 55 h (figures 1F, 2A, 3E marked v). The neuropil becomes compactly filled with neuronal profiles by 72 h (figure 2C). The larval synapses decrease in number during 12 to 18 h (figures 1AC; 3AC; 4C).
Neuronal growth cones and synaptic vesicles start appearing by 18 h (figure 1C, black star). Fasciculation of neuron profiles (figures 1D; 3D, marked f) and the formation of imaginal synapses were first detectable by 24 h (figures 1D; 3D, arrowhead). Morphological maturation of synapses is attained over a long period of time between 24 and 92 h (figures 2AF; 3DF). Around 65 h, synapses appear as large contiguous electron-dense regions (figures 2B, D; 3F, arrowhead) where only a few synaptic vesicles were visible. As the development progresses, the electron-dense material along the synaptic region condenses into discrete synaptic patches (figure 2C, E, arrow) and the synaptic vesicles were prominently visible. By 82 h, synapses appear fully mature with electron-dense synaptic thickenings and synaptic vesicles (figure 2F). At least 3 morphological types of synaptic vesicles were detectable: (i) electron-lucent vesicles (2560 nm in diam), (ii) electron-dense vesicles (5080 nm in diam) and (iii) dense core vesicles (80 nm in diam). In addition, neurosecretory granules were found in some neuron profiles. Neurosecretory profiles were found to be present throughout the period of metamorphosis (figures 1E; 3AC, marked s).
Estimates of degenerating neuron profiles in the neuropil and degenerating cells in the cortex were made by scoring them on electron micrographs covering 50,000 sq. m m area of ultrathin sections of tissue-samples at various times of pupal development. Results so obtained are shown in figure 4A. Increase in the number of neuronal profiles per unit area of ultrathin sections of brain-neuropil were similarly estimated from electron micrographs of known enlargement (figure 4B). It was observed that the neuronal profile counts continuously increase until 54 h of development, then tend to stabilize by 85 h (figure 4B). The number of synapses per unit area of ultrathin sections during the course of development were also estimated from electron micrographs and are given in figure 4C. The average number of synapse per neuronal profile in the brain-neuropil was estimated at a given time by dividing the number of synapses given in figure 4C with number of axon profiles read from the curve in figure 4B. This was considered necessary to overcome the scattering of observed points in figure 4B. Results so obtained are plotted in figure 4D.
3.2 Changes in the rind/cortex
Electron-dense cells showing signs of degeneration
were found in significant numbers in the cortex right from 6 h
through 60 h (figures 4A; 5AC). It was not possible to distinguish between, the degenerating neuronal or glial cells of the cortex by TEM. The number of degenerating cells present with progressing time is given in figure 4A.
3.3 Changes in muscles
Muscles of larval origin in pupa develop vacuoles by
6 h (figures 6A, B; 7A, B) and they start showing definite signs of degeneration
(figures 6A, B; 7B). These muscles lose most of their ultrastructural details between
1218 h. By 18 h, no trace of organized larval muscle is detectable by TEM
in the head, while in the thorax, except for a few known muscles such as larval oblique
muscles (Fernandes et al 1991) (figure 7C), most of the muscles histolyze.
Myoblasts are detectable from 8 h onward (figure 6D, 38 h sample) and
progressive development of the muscle precursor structure proceeds in them (figure 6C).
Gradually, these myoblasts form characteristic muscle structure, and by 8096 h,
the muscles are almost fully formed (figures 6E, F; 7E, F). However, a few myofibrils
continue to be added laterally to the preformed muscles even during
8696 h (figure 7D, 86 h sample, short thick arrow).
3.4 Electron-dense mitochondria
In addition to the normal-looking mitochondria of
characteristic shape, electron-dense mitochondria (EDMITs)
are detectable in neuropil (figure 1F, asterisk), cortex (figures 5C, D; 8B) and muscles during pupal development (figures 6F, 7A; 8A, C, asterisk). These EDMITs are at times found in isolation together with normal mitochondria (figures 5C; 6F) and are often found in large clusters, with more than 100 mitochondria (figures 5D, 7A, 8B, C). Usually, these clusters are near the surface of the cortex or muscle, but they also occur in the interior of the tissue (figure 5C). A few of them, referred to as electron-opaque mitochondria, are found enclosed in a vacuole and appear darker than the rest of EDMITs and normal-looking mitochondria (figures 5D, 7A, 8A arrow). These EDMITs were not found in the tissues of adult Drosophila.
Among several marker molecules known to be present in normal mitochondria (Lehninger 1990), cytochrome c and cytochrome oxidase were selected for histochemical characterization of EDMITs. Horseradish peroxidase-DAB reaction shows darkly stained small particles (1030 nm in diam) inside the normal-looking mitochondria (figure 8C, arrowhead). Similar darkly stained particles occur also in EDMITs (figure 8C, arrowheads), indicating the presence of cytochrome oxidase and cytochrome c in EDMITs.
For a general view of the progress of morphological development of the CNS of Drosophila,photomicrographs of 1 m m-thick sections of the brain region are shown in figure 9. The time course of major events during the development of the imaginal CNS of Drosophila during pupation at 25°C is given in figure 10.
Prior to undergoing metamorphosis, mature non-feeding larvae attain maximum carbohydrate reserves, which are mainly present as glycogen in the fat body and trehalose in the haemolymph (Chippendale 1978). The formation of imaginal CNS from the larval CNS takes place inside the confined space of puparium without intake of any nutrients, except air from the external surroundings. This necessitates that the system has sufficient energy reserves and be able to generate utilizable energy for the metamorphosis to be complete. During this period the biochemical decomposition of glycogen and trehalose supply glucose, as energy source for the formation of pupal and adult tissues (Chippendale 1978). At the initial stage of metamorphosis (612 h, figure 1A, B), many profiles within the neuropil contain glycogen deposits (Singh et al 1991), (figure 5AC). Since such glycogen deposits are not found in the neuropils of adult Drosophila, it is inferred that the glycogen present in the neuropils of pupa is utilized during metamorphosis as an energy source, in addition to other sources of energy such as: glycogen present in fat bodies, trehalose present in haemolymph and products of processes such as, cell degeneration. Though the energy derived from glycogen in the neuropil may be small compared to other mentioned processes, yet being located right within the neuropil, it is likely to have a significant role in the development of the neuropil and the processes within it.
Significant degeneration of cortical cells and their profiles in the neuropils, is seen during the first half of metamorphosis (up to 50 h) (figures 1AF, 3A, B). It is seen from figure 4A, where estimates of degenerating cells in the cortex and the profiles in the neuropil, are given that: (i) the cell bodies in the cortex show degeneration broadly peaking around 42 h and, (ii) at least 2 types of profiles exist in the neuropil, one that is maximally susceptible to degeneration at 12 h and the other whose degeneration peaks around 42 h. This bi-modal distribution can be understood in the light of the fact that the cortex in the CNS of Drosophila does not contain sensory neurons, which are on the periphery or away from CNS. The cortex consists of cell bodies of interneurons, motor neurons, glial, and neurosecretory cells. We suggest that the 42 h degeneration peaks in figure 4A correspond to some of these cells and their profiles in the neuropil, whereas the 12 h peak (figure 4A) corresponds to the degeneration of sensory profiles, whose cell bodies are not present in the cortex. We are aware that there could be other explanations. For example it may be possible to explain these results by assuming that there are 2 types of neuron profiles namely inhibitory and excitatory.
It is observed from figure 4B that the number of axon profiles in a given area of a section, or for that matter in a given volume, increases with the progress of development. These results are comparable with the increase in the number of Kenyon cell axons during pupation studied by Technau and Heisenberg (1982). Contrary to the expectation, the size of axon profiles does not increase appreciably; it is their number that increases. This suggests that branching of axon profiles increases during the development.
In addition to the neurosecretory granules, we found only 3 prominent types of synaptic vesicles in the CNS: electron-lucent, electron dense and dense core vesicles. We were not able to detect flattened vesicles in significant numbers. These synaptic vesicles are comparable to those described earlier by Shepherd (1979) for vertebrate nervous system: (i) electron-lucent, spherical (4060 nm in diam) vesicles containing acetylcholine or amino acids, (ii) electron-lucent flattened vesicles (3060 nm long) having g -amino butyric acid or glycine, (iii) electron-dense or dense-core (40100 nm in diam) vesicles containing catecholamines and (iv) large electron-dense vesicles (100160 nm in diam) known to be neurosecretory granules. All the above types of vesicles were prominently present in Drosophila neuron profiles, except electron-lucent flattened vesicles.
The number of synapses per unit area of sections initially decreases during the development and is minimum at 18 h (figure 4C). By this time, some of the synapses of larval origin, histolyze. Then there is a 2-step increase in the number of synapses per unit area until a peak value is reached at 72 h; this suggests that developmentally, at least 2 classes of synapses are formed during this period. Soon after, the number of synapses decrease sharply until 80 h (figure 4C). The observed decrease of 7075% in the total number of synapses, within a span of 6 h out of 100 h of pupal development could be dismissed as fictitious, if only there was 300400% increase in the volume of CNS, during this 6 h. Certainly, this does not seem to be the case. Drosophila, like Musca domestica (Fröhlich and Meinertzhagen 1983), makes synapses in excess, and finally only 2530% of the maximum number of synapses formed are retained and the rest eliminated.
The average number of synapse per axon profile in the neuropil (figure 4D) has a close resemblance with the average number of synapses per unit area during the course of development (figure 4C). Because 2 neuron profiles share one monad synapse, a value of 0·5 or more for an average number of synapse per axon profile indicates that approximately every neuron profile is engaged in making a synapse at 72 h (figure 4D).
The metamorphosis of larval muscles to imaginal muscles goes through a more drastic change than the CNS. In CNS during metamorphosis, the neuronal profiles do not vanish completely; rather they become loose and the space so generated is filled with extracellular fluid, most probably produced by histolysis. In muscle metamorphosis, however, most of the muscles in head and all muscles, except dorsal thoracic muscles and some pharyngeal muscles (Bodenstein 1950; Fernandes et al 1991) in thorax, histolyze. New muscles are probably formed from the molecules generated by histolysis. What sort of changes these molecules undergo before being incorporated into imaginal muscles, is unknown. The control processes operating at any of these steps are worth investigating.
The presence of a large number of EDMITs is a positive indication that the imaginal structures, in need of readjustment of their energy requirement, discard a number of such organelles. For the following reasons, these EDMITs are likely to be degenerating mitochondria: (i) presence of a delimiting double-membrane in the EDMITs, (ii) lack of ultrastructural details such as cristae, and (iii) occurrence of some EDMITs inside vacuoles, indicating that they are going to be histolyzed or ejected out of the cell.
The possibility of EDMITs being degenerating mitochondria, raises the likelihood that the cellular energy requirement in pupa and adult are quite different. Because of differences in the habitats of the pupa and the adult, the pupa proportionally requires many more mitochondria than the adult or the mitochondria in adult are qualitatively different from the pupal mitochondria. In the latter case, old mitochondria are discarded and the new adult mitochondria are regenerated during pupation. In the former case, only a proportion of pupal mitochondria will be converted to EDMITs and discarded, and the rest would be utilized during pupation for the formation of adult tissue. In either case, the energy requirements of pupa and adult Drosophila, seem to be significantly different.
We thank Mrs Shubha Shanbhag and Dr Rajashekhar Patil for their helpful suggestions.
Bate C M 1976 Embryogenesis of an insect nervous system. I. A map of the thoracic and abdominal neuroblasts in Locusta migratoria; J. Embryol. Exp. Morphol. 35 107123
Bate C M 1978 Development of sensory systems in arthropods, in Handbook of sensory physiology (ed.) M Jacobson (Berlin: Springer-Verlag) vol. 9, pp 153
Bodenstein D 1950 The postembryonic development of Drosophila; in Biology of Drosophila (ed.) M Demerec (New York: Hafner Publishing Company) 2nd edition 1965, pp 275367
Crammer W and Moore C L 1973 Oxidation of 3,3¢ -diaminobenzidine by rat liver mitochondria; Biochemistry 12 25022509
Campos-Ortega J A and Hofbauer A 1977 Cell clones and pattern formation: on the lineage of photoreceptor cells in the compound eye of Drosophila; Wilhelm Rouxs Arch. Dev. Biol. 181 227245
Campos-Ortega J A and Hartenstein V 1985 The Embryonic development of Drosophila melanogaster (Berlin: Springer-Verlag)
Canal I and Ferrus A 1986 The pattern of early neuronal differentiation in Drosophila melanogaster; J. Neurogen. 3 293319
Chippendale G M 1978 The functions of carbohydrates in insect life processes; in Biochemistry of insects (ed.) M Rockstein (New York: Academic Press) pp 154
Crammer W and Moore C L 1973 Oxidation of 3,3¢ -diaminobenzidine by rat liver mitochondria; Biochemistry 12 25022509
Doe C Q and Goodman C S 1985 Early events in insect neurogenesis. I. Developmental and segmental differences in the pattern of neuronal precursor cells; Dev. Biol. 111 193205
Dalton A J 1955 A chrome-osmium tetroxide fixative for electron microscopy; Anat. Rec. 121 281A
Edwards J S 1969 Postembryonic development and regeneration in the insect nervous system; Adv. Insect Physiol. 6 97137
Fernandes J, Bate M and. Vijayraghavan K 1991 Development of the indirect flight muscles of Drosophila; Development 113 6777
Fredieu J R and Mahowald A P 1989 Glial interactions with neurons during Drosophila embryogenesis; Development 106 739748
Fröhlich A and Meinertzhagen I A 1983 Quantitative features of synapse formation in the flys visual system. I. The presynaptic photoreceptor terminal; J. Neurosci. 11 23362349
Ghysen A and Dambly-Chaudiere C 1990 Early events in the development of Drosophila peripheral nervous system; J. Physiol. (Paris) 84 1120
Giangrande A and Palka J 1990 Genes involved in the development of the peripheral nervous system of Drosophila; Semin. Cell Biol. 1 197209
Hartenstein V 1988 Development of Drosophila larval sensory organs: spatiotemporal pattern of sensory neurons, peripheral axonal pathways and sensilla differentiation; Development 102 869886
Hartenstein V and Campos-Ortega J A 1984 Early neurogenesis in wild-type Drosophila melanogaster; Wilhelm Rouxs Arch. Dev. Biol. 193 308325
Hartenstein V Rudloff E and Campos-Ortega J A 1987 The pattern of proliferation of the neuroblasts in the wild-type embryo of Drosophila melanogaster; Wilhelm Rouxs Arch. Dev. Biol. 196 473485
Hartweck H 1931 Anatomie und Variabilität des Nervensystem und der Sinnesorganne von Drosophila melanogaster (Meigen); Z. Wiss. Zoöl. 139 559663
Jacobs J R and Goodman C S 1989a Embryonic development of axon pathways in the Drosophila CNS. I. Role of glia; J. Neurosci. 9 24012411
Jacobs J R and Goodman C S 1989b Embryonic development of axon pathways in the Drosophila CNS. II. Behavior of pioneer growth cones; J. Neurosci. 9 24122422
Kankel D R and Hall J C 1976 Fate mapping of nervous system and other internal tissues in genetic mosaics of Drosophila melanogaster; Dev. Biol. 48 124
Kankel D R, Ferrus A, Garen S H, Harte P J and Lewis P E 1980 The structure and development of the nervous system; in The Genetics and biology of Drosophila (eds) M Ashburner and T R F Wright (London: Academic Press) vol. 2d, pp 295368
Karnovsky M J 1965 A formaldehyde-glutaraldehyde fixative of high osmolarity for use in electron microscopy; J. Cell Biol. 27 137A
Lawrence P A 1973 Polarity and patterns in the postembryonic development of insects; Adv. Insect Physiol. 7 197266
Lees A D and Waddington C H 1942 The development of the bristles in normal and some mutant types of Drosophila melanogaster; Proc. R. Soc. London Ser. B. 131 87110
Lehninger A L 1990 Principles of Biochemistry (Delhi: CBS Publishers and Distributors) pp 435460
Malzacher P 1968 Die Embryogenese des Gehirns paurometaboler Insekten, Untersuchungen an Carausius morosus und Periplaneta americana; Z. Morphol. Okol. Tier. 62 103161
Meinertzhagen I A 1973 Development of the compound eye and optic lobe of insects; in Developmental neurobiology of arthropods (ed.) D Young (Cambridge: Cambridge University Press) pp 51104
Murray M A, Schubiger M and Palka J 1984 Neuron differentiation and axon growth in the developing wing disc of Drosophila melanogaster; Dev. Biol. 104 259273
Nässel D R 1983 Horseradish peroxidase and other heme proteins as neuronal markers; in Functional Neuroanatomy (ed.) N J Strausfeld (Berlin: Springer-Verlag) pp 4491
Poulson D F 1950 Histogenesis, organogenesis and differentiation in the embryo of Drosophila melanogaster Meigen; in Biology of Drosophila (ed.) M Demerec (New York: Hafner Publishing Company) 2nd edition, 1965, pp 168274
Power M E 1943 The brain of Drosophila melanogaster; J. Morphol. 72 517559
Power M E 1948 The thoracico-abdominal nervous system of an adult insect, Drosophila melanogaster; J. Comp. Neurol. 88 347409
Power M E 1952 A quantitative study of the growth on the central nervous system of a holometabolous insect, Drosophila melanogaster; J. Morphol. 91 389411
Prokop A and Technau G M 1991 The origin of postembryonic neuroblasts in the ventral nerve cord of Drosophila melanogaster; Development 111 7988
Ready D F, Hanson T E and Benzer S 1976 Development of the Drosophila retina, a neurocrystalline lattice; Dev. Biol. 53 217240
Reith A and Schüler B 1972 Demonstration of cytochrome oxidase activity with diaminobenzidine. A biochemical and electron microscopic study; J. Histochem. Cytochem. 20 583589
Reynolds E S 1963 The use of lead citrate of high pH as an electron opaque stain in electron microscopy; J. Cell Biol. 17 208212
Robertson C W 1936 Metamorphosis of Drosophila melanogaster, including an accurately timed account of
the principal morphological changes; J. Morphol. 59 351399
Seligman A M, Karnovsky M J, Wasserkrug H L and Hanker J S 1968 Nondroplet ultrastructural demonstration of cytochrome oxidase activity with a polymerizing osmiophilic reagent, diaminobenzidine (DAB); J. Cell Biol. 38 114
Shepherd G M 1979 Neurobiology: The synapse (New York: Oxford University Press) pp 7583
Singh K, Singh R N and Kankel D R 1991 How do the fine structures of neuropils of larva and adult differ in Drosophila?; Drosophila Inform. Service 70 212214
Singh R N, Singh K and Kankel D R 1989 Development and fine structure of the nervous system of lethal (1)
optic ganglion reduced visual mutants of Drosophila melanogaster; in Neurobiology of sensory systems (eds)
R N Singh and N J Strausfeld (New York: Plenum Press) pp 203218
Sink H and Whitington P M 1991 Pathfinding in the central nervous system and periphery by identified embryonic Drosophila motor axons; Development 112 307316
Sonnenblick B P 1950 The early embryology of Drosophila melanogaster; in Biology of Drosophila (ed.) M Demerec (New York: Hafner Publishing Company) 2nd edition 1965, pp 62167
Technau G M and Heisenberg M 1982 Neural reorganization during metamorphosis of the corpora pedunculata in Drosophila melanogaster; Nature (London) 295 405407
Thomas J B, Bastiani M J, Bate M and Goodman C S 1984 From grasshopper to Drosophila: a common plan for neuronal development; Nature (London) 310 203207
Tomlinson A and Ready D F 1987 Neuronal differentiation in the Drosophila ommatidium; Dev. Biol. 120 366376
Truman J W 1990 Metamorphosis of the central nervous system of Drosophila; J. Neurobiol. 21 10721084
Truman J W and Bate M 1988 Spatial and temporal patterns of neurogenesis in the CNS of Drosophila melanogaster; Dev. Biol. 125 145157
White K and Kankel D R 1978 Patterns of cell division and cell movement in the formation of the imaginal nervous system in Drosophila melanogaster; Dev. Biol. 65 296321
Zalokar M 1947 Anatomie du thorax de Drosophila melanogaster; Rev. Suisse. Zool. 54 1753
Zipursky S L, Venkatesh T R, Teplow D B and Benzer S 1984 Neuronal development in the Drosophila retina: monoclonal antibodies as molecular probes; Cell 36 1526
MS received 31 May 1999; accepted 23 June 1999
Corresponding editor: Veronica Rodrigues
BACK TO CONTENTS