Malpighian (renal) tubules are key components of the insect osmoregulatory system and show correspondingly great diversity in both number and length. Recently, the organisation of the Drosophila melanogaster tubule has been elucidated by enhancer trapping, and an array for functional properties has been shown to align with the functional domains. In Drosophila,there is a lower tubule domain, which coincides with expression of alkaline phosphatase and delineates the absorptive region of the tubule. Here, these observations are extended to three dipteran vectors of disease (Aedes aegypti, Anopheles stephensii and Glossina morsitans) and a non-dipteran out-group, Schistocerca gregaria (Orthoptera). Despite a huge range in cell number and size, alkaline phosphatase was found on the apical surface of the lower 10% of each of the dipteran tubules but nowhere within the orthopteran tubule. An alkaline phosphatase lower tubule domain is thus conserved among Diptera.

Cell counts are also provided for each species. As in Drosophila,stellate cells are not found in the lower tubule domain of Anophelesor Aedes tubules, confirming the unique genetic identity of this domain. As previously reported, we failed to find stellate cells in Schistocerca but, remarkably, also failed to find them in Glossina, the dipteran most closely related to Drosophila. The orthodoxy that stellate cells are unique to, and general among, Diptera may thus require revision.

Alkaline phosphatases (ALPs) are ubiquitous enzymes in all organisms. ALPs are abundant, high-pH metalloproteins known to play roles in phosphate uptake and in secretory processes in epithelia in mammals. However, the precise physiological role of ALPs remains unknown.

In insects, functional ALP has been shown to exist in Leptinotarsa decemlineata (Colorado potato beetle; Yi and Adams, 2001), Bombyx mori (silkworm) gut (Eguchi et al., 1990), Bemisia tabaci (whitefly) salivary glands(Funk, 2001), Drosophila melanogaster brain (Yang et al.,2000) and several mosquito species(Houk and Hardy, 1984; Igbokwe and Mills, 1982). However, given their role in secretory function, ALPs have also been studied in insect Malpighian tubules, which are critical osmoregulatory and secretory organs in insects; ALP expression has been documented in L. decemlineata and D. melanogaster(Yang et al., 2000) tubules. In particular, the D. melanogaster tubule is a genetically accessible transporting epithelium (Dow and Davies,2003) and is thus a useful model in which to study the functional role of ALP in vivo.

Previous work in D. melanogaster has shown that tubule ALP expression is confined to cells in the lower segment(Sozen et al., 1997; Yang et al., 2000), which plays a role in fluid re-absorption(O'Donnell and Maddrell,1995). Furthermore, a P-element mutation in the ALP gene, Aph-4, causes misexpression of Aph-4 in the tubules; expression is substantially reduced in the lower tubule but increased in the main segment. This misexpression results in reduced basal and stimulated fluid secretion by the tubules. Thus, ALP is likely to have a significant - although as yet unexplained - role in epithelial transport in D. melanogaster.

Insect Malpighian tubules are important to insect life and therefore constitute important targets for population control. The power of D. melanogaster as a genetic model organism has allowed significant steps in understanding tubule function (Dow and Davies, 2001, 2003), which can then be applied to those insect species with less-developed genomic resources but greater economic or medical significance; for example, tsetse fly or mosquito. Thus, we have begun to apply knowledge of tubule function in D. melanogaster to those of insect vector species with a view to advancing understanding of tubule physiology in the context of specific cell types and tubule regions in these animals. The species selected for the present study,in addition to D. melanogaster, are Anopheles stephensi (a malarial mosquito), Aedes aegypti (a mosquito vector for dengue fever) and Glossina morsitans (a vector for African sleeping sickness). Together, these provide a wide phylogenetic spread through the Diptera, the largest insect order. Additionally, we selected the locust Schistocerca gregaria as an out-group from the exopterygote order Orthoptera.

We show here that tubules from dipteran species - An. stephensi, Ae. aegypti and G. morsitans - share identical domains of ALP expression to D. melanogaster. However, tubules from Schistocerca gregaria do not show any significant staining for ALP. Thus, defined ALP expression in dipteran tubules may be an important feature of fluid transport mechanisms by these tubules and may constitute important control elements for fluid secretion/absorptive pathways.

Insects

D. melanogaster Meigen wild-type Oregon R (OrR) flies were maintained on standard D. melanogaster diet on a 12 h:12 h photoperiod at 55% humidity at 22°C. Aedes aegypti L. were obtained as non-infective adults from a colony maintained by Professor Eileen Devaney, Department of Veterinary Parasitology, University of Glasgow. Animals were used upon receipt. Non-infective Anopheles stephensi Liston adults were provided as a kind gift of Dr L. Ranford-Cartwright, Division of Infection and Immunity, IBLS, University of Glasgow. Animals were used upon receipt. If mosquitoes were not used immediately, they were maintained on a 12 h:12 h photoperiod at 55% humidity at 22°C on 5% sucrose (v/v) solution ad libitum for a maximum of three days before use in experiments. Non-infective Glossina morsitans Westwood adults were provided by the Centre for Tropical Veterinary Medicine, Royal School of Veterinary Studies,University of Edinburgh. These were also provided as a kind gift from a colony maintained by Professor D. Barry, Wellcome Centre for Molecular Parasitology,University of Glasgow. Animals were used immediately upon receipt. Schistocerca gregaria Forskal were obtained from Bugs Direct (Well Cottages, Devon, UK) and either used immediately or maintained on grass over a 12 h:12 h photoperiod at 55% humidity at 22°C for a maximum of 4 days.

All insects were cold-anaesthetised and decapitated prior to dissection to isolate intact tubules.

Histochemistry for alkaline phosphatase activity

The methods for staining of alkaline phosphate in tubules was essentially that of Yang et al. (2000). Briefly, intact tubules were dissected into wells containing phosphate-buffered saline (PBS) and treated with 4% (v/v) paraformaldehyde for 20 min. Tubules were washed for 3×10 min in PBS/0.1% Triton and then for 2×2 min in PBS. Tubules were incubated for 10-30 min with 75 mg ml-1 4-nitroblue tetrazolium chloride (NBT)/50 mg ml-15-bromo-4-chloride-3-indolyl-phosphate (BCIP) in dimethylfluoride (DMF) and used at 4.5 μl of NBT and 3.5 μl BCIP per 1 ml DIG kit detection buffer,pH 9.5 (Boehringer, Bracknell, UK). Control tubules were processed in the same way with the exception of NBT/BCIP addition. Tubules were washed for 3×2 min in PBS, viewed under a standard microscope and photographed with a digital camera.

Where required, tubules were stained for ALP activity and with 4′,6′-diamidino-2-phenylindole hydrochloride (DAPI; Sigma-Aldrich,Gillingham, UK). For these experiments, 1 μg ml-1 DAPI was added in PBS to ALP-stained tubules after the final 3×2 min PBS washes (see above) for 2.5 min. Tubules were washed in PBS for 3×2 min.

For all protocols, prior to mounting on slides, tubules were washed with glycerol as follows: 2×20% glycerol/PBS (v/v); 2×50% glycerol/PBS(v/v); 2×80% glycerol/PBS (v/v). Samples were viewed with the Axiocam imaging system (Zeiss, Welwyn Garden City, UK).

Statistical analysis

Where appropriate, the significance of differences was tested with Student's t-test (two-tailed), assuming unequal variances and taking the critical level to be P<0.05.

Comparative analysis of Malpighian tubule structure in Diptera

Results in Table 1 show the overall cell numbers of both principal and stellate cells in tubules of each of the insects used in this study.

Table 1.

Inter-species comparison of tubule structure

SpeciesTubule numberTubule length (mm)Principal cells per tubuleStellate cells per tubuleTotal cellsN
D. melanogaster 146±5 33±1 179±6 
Ae. aegypti 5, 3 46±1 10±1 54±2 25 
An. stephensi 5, 3 51±2 12±1 72±2 25 
G. morsitans 60 1016±10 1016±12 10 
S. gregaria ∼200 25 2024±10 2024±14 10 
SpeciesTubule numberTubule length (mm)Principal cells per tubuleStellate cells per tubuleTotal cellsN
D. melanogaster 146±5 33±1 179±6 
Ae. aegypti 5, 3 46±1 10±1 54±2 25 
An. stephensi 5, 3 51±2 12±1 72±2 25 
G. morsitans 60 1016±10 1016±12 10 
S. gregaria ∼200 25 2024±10 2024±14 10 

Intact tubules from D. melanogaster, Ae. aegypti, An. stephensi, G. morsitans and S. gregaria were dissected and stained with DAPI as described (representative images shown in Figs 2, 3, 4, 5). Stained nuclei were counted from several tubules from each species, and data for cell counts, to the nearest unit, are expressed ± s.e.m.

The numbers for D. melanogaster are in close agreement with those previously published (Sozen et al.,1997), with stellate cells constituting 21-24% of overall tubule cells. Interestingly, while in both Ae. aegypti and An. stephensi the five tubules are of different lengths, the overall cell number is similar in either long or short tubules. Furthermore, stellate cells constitute a similar percentage of tubule cells in both species: approximately 19-24% of total cells in Ae. aegypti and 21-26% in An. stephensi; these values are similar to those for D. melanogaster. The cell numbers obtained for Ae. aegypti are in close agreement with a previous study on tubules from other mosquitoes,including Ae. aegypti (Satmary and Bradley, 1984); the close agreement of cell numbers amongst mosquito species, including Aedes taeniorhyncus, Culex tarsalis and Culiseta inornata (Satmary and Bradley, 1984), support the An. stephensi cell numbers shown here. Interestingly, neither on the basis of nuclear size(Sozen et al., 1997) nor cell shape could stellate cells be observed in G. morsitans tubules. Although not explicitly addressed, previous work on G. morsitanstubule ultrastructure also failed to document stellate cells(Kongoro and Odhiambo,1988).

Amongst the dipteran species tested, cell number per mm of tubule was similar between the mosquitoes and G. morsitans. D. melanogaster,however, has vastly more cells per mm of tubule: nearly 80 in D. melanogaster versus 11-17 in the other Diptera.

We took the opportunity to perform a systematic cell count of each tubule in the two mosquito species (Fig. 1). This shows that while each of the tubules differs in cell count from the others, this is mainly attributable to differences in principal cell number. Both stellate cells and the numbers of cells in the ALP domain are almost invariant between tubules. It is also worth noting that, as in D. melanogaster, cell numbers are almost invariant. These data thus usefully show that the precision with which the D. melanogastertubule is specified (Sozen et al.,1997) is not unique to this genetic model organism.

Fig. 1.

Comparison of cell numbers in the five Malpighian tubules of mosquitoes. Total, principal and stellate cell counts, and cell count in alkaline phosphatase domain, for the longest (1) to the shortest (5) tubule in each insect. Data are means ± s.e.m.(N=10) for Ae. aegypti (left) and An. stephensi(right).

Fig. 1.

Comparison of cell numbers in the five Malpighian tubules of mosquitoes. Total, principal and stellate cell counts, and cell count in alkaline phosphatase domain, for the longest (1) to the shortest (5) tubule in each insect. Data are means ± s.e.m.(N=10) for Ae. aegypti (left) and An. stephensi(right).

Use of the locust as an out-group in this study shows that the notable differences between the S. gregaria tubule and those of the Diptera are numbers of tubules per insect and the lack of stellate cells (compared with D. melanogaster, Ae. aegypti and An. stephensi).

Alkaline phosphatase activity is confined to the lower tubule in dipteran insects

Previous work has shown that ALP expression in D. melanogastertubules is confined to the lower tubule and precisely matches the lower-main segment boundary (Yang et al.,2000). On close examination, combined with DAPI counter-staining of nuclei, it is clear that ALP is concentrated in the apical membrane(Fig. 2D).

Fig. 2.

Alkaline phosphatase domain in an intact adult D. melanogastertubule pair: (A) control tubules subjected to alkaline phosphatase staining protocol without chromogenic substrate; the junction with the ureter is shown with an arrow; (B) DAPI-stained; (C) as in B but subjected to alkaline phosphatase staining protocol in the presence of NBT/BCIP chromogenic substrate; the junction with the ureter is shown with an arrow; (D) higher magnification to show predominantly apical localisation of staining. Scale bars, 100 μm for A-C, 10 μm for D.

Fig. 2.

Alkaline phosphatase domain in an intact adult D. melanogastertubule pair: (A) control tubules subjected to alkaline phosphatase staining protocol without chromogenic substrate; the junction with the ureter is shown with an arrow; (B) DAPI-stained; (C) as in B but subjected to alkaline phosphatase staining protocol in the presence of NBT/BCIP chromogenic substrate; the junction with the ureter is shown with an arrow; (D) higher magnification to show predominantly apical localisation of staining. Scale bars, 100 μm for A-C, 10 μm for D.

While the genetic determination of tubule sub-regions is currently unknown in other dipteran species, we show that in Ae. aegypti, An. stephensiand G. morsitans, staining is confined to the lower tubule (Figs 3, 4, 5). Interestingly, in spite of the different lengths of mosquito tubules, expression of ALP is observed in the lower domain to the same extent in all tubules in both Ae. aegypti and An. stephensi(Fig. 1).

Fig. 3.

Intact Ae. aegypti tubules. (A) Control tubules subjected to alkaline phosphatase staining protocol without chromogenic substrate; the junction with the gut is shown with an arrow; (B) as in A but subjected to alkaline phosphatase staining protocol in the presence of NBT/BCIP chromogenic substrate; the junction with the gut is shown with an arrow; (C) DAPI-stained;(D) as in C but also stained for alkaline phosphatase. Scale bars, 100μm.

Fig. 3.

Intact Ae. aegypti tubules. (A) Control tubules subjected to alkaline phosphatase staining protocol without chromogenic substrate; the junction with the gut is shown with an arrow; (B) as in A but subjected to alkaline phosphatase staining protocol in the presence of NBT/BCIP chromogenic substrate; the junction with the gut is shown with an arrow; (C) DAPI-stained;(D) as in C but also stained for alkaline phosphatase. Scale bars, 100μm.

Fig. 4.

Intact An. stephensi tubules. (A) Control tubules subjected to alkaline phosphatase staining protocol without chromogenic substrate; the junction with the gut is shown with an arrow; (B) as in A but subjected to alkaline phosphatase staining protocol in the presence of NBT/BCIP chromogenic substrate; the junction with the gut is shown with an arrow; (C) DAPI-stained;(D) as in C but also stained for alkaline phosphatase; (E) high-magnification view of lightly stained lower tubule, revealing predominantly apical staining. Scale bars, 100 μm for A-D; 10 μm for E.

Fig. 4.

Intact An. stephensi tubules. (A) Control tubules subjected to alkaline phosphatase staining protocol without chromogenic substrate; the junction with the gut is shown with an arrow; (B) as in A but subjected to alkaline phosphatase staining protocol in the presence of NBT/BCIP chromogenic substrate; the junction with the gut is shown with an arrow; (C) DAPI-stained;(D) as in C but also stained for alkaline phosphatase; (E) high-magnification view of lightly stained lower tubule, revealing predominantly apical staining. Scale bars, 100 μm for A-D; 10 μm for E.

Fig. 5.

Intact G. morsitans tubules. (A) Control tubules subjected to alkaline phosphatase staining protocol without chromogenic substrate; the junction with the ureter is shown with an arrow; (B) as in A but subjected to alkaline phosphatase staining protocol in the presence of NBT/BCIP chromogenic substrate; the junction with the ureter is shown with an arrow; (C) high-power magnification of B; (D) as in C but different tubule preparation showing lower tubule/ureter boundaries and normally close association of tubules at the lower segment (arrows); (E) DAPI-stained; (F) as in E but also stained for alkaline phosphatase. Lower tubule segments remain closely intertwined in tubules shown in E and F. Scale bars, 100 μm.

Fig. 5.

Intact G. morsitans tubules. (A) Control tubules subjected to alkaline phosphatase staining protocol without chromogenic substrate; the junction with the ureter is shown with an arrow; (B) as in A but subjected to alkaline phosphatase staining protocol in the presence of NBT/BCIP chromogenic substrate; the junction with the ureter is shown with an arrow; (C) high-power magnification of B; (D) as in C but different tubule preparation showing lower tubule/ureter boundaries and normally close association of tubules at the lower segment (arrows); (E) DAPI-stained; (F) as in E but also stained for alkaline phosphatase. Lower tubule segments remain closely intertwined in tubules shown in E and F. Scale bars, 100 μm.

In tubules of the out-group species, S. gregaria, no staining for ALP was ever observed (Fig. 6). Incubation conditions were varied to maximise the chances of observing ALP staining, including increasing staining times to up to 24 h. However, specific staining was not noted under any experimental condition.

Fig. 6.

Single intact S. gregaria tubule. (A) Control tubules subjected to alkaline phosphatase protocol without chromogenic substrate; (B) as in A but with NBT/BCIP chromogenic substrate; (C) DAPI-stained; (D) high-magnification image of lower tubule, without chromogenic substrate; (E) high-magnification image of B. Scale bars, 100 μm.

Fig. 6.

Single intact S. gregaria tubule. (A) Control tubules subjected to alkaline phosphatase protocol without chromogenic substrate; (B) as in A but with NBT/BCIP chromogenic substrate; (C) DAPI-stained; (D) high-magnification image of lower tubule, without chromogenic substrate; (E) high-magnification image of B. Scale bars, 100 μm.

Alkaline phosphatase staining in the lower domain involves precise numbers of cells

Counterstaining for cell nuclei and ALP allowed the determination of cell numbers within the tubule lower domains for each species (Figs 2D, 3D, 4D,E, 5F). Table 2 shows the results for such analysis. Consistent with the fact that Ae. aegypti and An. stephensi tubules have fewer cells than either D. melanogasteror G. morsitans, there are fewer cells in the ALP domain of both mosquito species. Interestingly, while the absolute number of cells in the ALP domain may vary between species, the percentage of all tubule cells localised to the ALP domain in the dipteran species studied is virtually identical(∼10). Thus, the distribution and function of the cells in this lower domain must be specified by the same genetic determinants in each of the dipteran species studied here.

Table 2.

Cell counts in alkaline phosphatase (ALP) domain

SpeciesNumber of cells in ALP domainCells in ALP domain as a % of total number of tubule cellsN
D. melanogaster 22±1 12 
Ae. aegypti 20 
An. stephensi 25 
G. morsitans 104±1 10 12 
S. gregaria NA NA NA 
SpeciesNumber of cells in ALP domainCells in ALP domain as a % of total number of tubule cellsN
D. melanogaster 22±1 12 
Ae. aegypti 20 
An. stephensi 25 
G. morsitans 104±1 10 12 
S. gregaria NA NA NA 

Intact tubules from D. melanogaster, Ae. aegypti, An. stephensiand G. morsitans were dissected and stained with DAPI, then stained for alkaline phosphatase as described for Figs 2, 3, 4, 5. Stained nuclei in the alkaline phosphatase-stained region were counted from several tubules from each species, and data for cell counts, to the nearest units, are expressed± s.e.m. Errors of less than 1 are omitted. NA, not appropriate.

Furthermore, the relative numbers of stellate cells in D. melanogaster,Ae. aegypti and An. stephensi are the same: ∼18%. This further supports the view that the genetic determinants that set tubule domains are conserved across the Diptera.

Cell-specific in vivo roles for ALPs are still unknown, although ALP has functional significance in vertebrate epithelia including intestine,kidney, placenta and mammary gland(Fishman, 1990). Therefore,this enzyme has a primary function in transporting epithelia. Mutations in the D. melanogaster ALP gene, Aph-4, affect fluid transport by the Malpighian tubule; this suggests that the important role for ALP in epithelial transport is conserved across evolution. Furthermore, at least two ALP isoforms encoded by different genes are expressed in Bombyx morilarval midgut (Eguchi, 1995),supporting the view that ALP does have functional significance in insect epithelia. In Bombyx, membrane-bound and soluble isoforms of ALP are distributed differentially in midgut; mALP, for example, is located at the brush border membrane. Data presented here suggest that, in D. melanogaster and An. stephensi at least, the enzyme is localised to the apical membrane in tubules. This is consistent with the peptide sequence of Aph-4, the ALP gene expressed in Drosophilalower tubule (Yang et al.,2000); like its mammalian counterpart, it has a GPI anchor site(Eisenhaber et al., 2003). However, pronounced cytosolic staining in addition to staining at the apical membrane in Ae. aegypti, An. stephensi and G. morsitanstubules may suggest that, in these insects, a soluble ALP may also be involved. However, it is also possible that the cytoplasmic staining represents a pool of enzyme destined for the membrane. Several ALPs have been detected by biochemical analysis in Culex tarsalis(Houk and Hardy, 1984) and in Ae. aegypti (Igbokwe and Mills,1982). Furthermore, ALP isoforms have been demonstrated in European anopheline mosquitoes (Bianchi,1968).

We have shown for the first time that ALP activity marks a conserved lower tubule domain of dipteran insects. While it is impossible to assert absolutely that this tightly conserved expression domain represents a conserved vital function, the closeness of conservation across even a wide range of dipteran taxa suggests strongly that there is a functional role required of the lower 10% of dipteran tubules. It will be interesting (though technically very demanding) to establish in the future whether the reabsorptive function of this domain is also conserved beyond Drosophila. In D. melanogaster, the ALP expression pattern is identical to that marking the lower tubule domain by enhancer trapping(Yang et al., 2000). Data in Table 2 also show that the proportion of cells in the ALP domain compared with overall cell number in the tubule is similar in the dipteran species tested. However, the tsetse tubule can be differentiated from those of the other Diptera: it lacks stellate cells. To our knowledge, this is the first time that stellate cells have been found to be absent in a dipteran.

A mutation in the D. melanogaster ALP gene results in reduced basal rates of fluid transport (Yang et al., 2000) without affecting stimulated transport via the neuropeptides CAP2b (Davies et al., 1995) or drosokinin(Terhzaz et al., 1999). Thus,this suggests that ALP has a specific role in maintaining resting rates of transport via cellular events in the lower domain. Given that the lower domain has been shown to be involved in fluid reabsorption(O'Donnell and Maddrell,1995), the possibility exists that ALP influences the overall rate of fluid transport by controlling fluid reabsorption. By extension, this situation may also exist in the mosquito and tsetse tubule. Developmental work in Schistocerca americana has demonstrated anchoring of epithelial ALP to the plasma membrane by glycosyl-phosphatidylinositol and involvement of phospholipase C in release of ALP from the anchoring sites(Chang et al., 1993). Given that fluid transport in D. melanogaster tubules is under control by several signalling pathways (Dow and Davies, 2001), including that involving phospholipase C (PLC; Pollock et al., 2003), it is entirely possible that signalling processes are an important control mechanism for fluid reabsorption involving ALP action.

In this work, we show that the ALP/lower domain is conserved in dipteran species but not in an out-group, S. gregaria. Does S. gregaria express ALP at all? Published work has shown that both fat body(George and Eapen, 1959) and alimentary canal (Navqi, 1981)contain ALP. However, expression in tubules has not been documented. Thus, the important role for ALP in the lower tubule may not be a feature of orthopteran insects.

There are also intriguing comparative aspects to our study(Fig. 7). The tsetse fly is the most closely related to D. melanogaster; consistent with this, it has four tubules, arranged in two pairs, each sharing a common ureter. However, it lacks stellate cells, even though these are found in the much more distantly related mosquito species, although they each have five tubules that do not have shared ureters. Stellate cells have also been demonstrated in a species of intermediate distance, Calliphora erythrocephala(Berridge and Oschman, 1969). It is thus reasonable to suppose that the unique lifestyle of G. morsitans (it is an obligate blood-feeder and gives birth to live young)has led to secondary loss of the dipteran stellate cell. There may thus be useful discrimination built into these simple functional morphological techniques; it will be interesting to extend them to other insects in the future.

Fig. 7.

Taxonomy to genus of the insects used in this study. Taxonomy is that of NCBI(http://www.ncbi.nlm.nih.gov/Taxonomy/).

Fig. 7.

Taxonomy to genus of the insects used in this study. Taxonomy is that of NCBI(http://www.ncbi.nlm.nih.gov/Taxonomy/).

This work was supported by the Biotechnology and Biological Sciences Research Council (UK) Genomics in Animal Function (GAIN) initiative.

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