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Am J Physiol Renal Physiol 283: F499-F508, 2002. First published April 10, 2002; doi:10.1152/ajprenal.00041.2002
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Vol. 283, Issue 3, F499-F508, September 2002

Leucokinin activates Ca2+-dependent signal pathway in principal cells of Aedes aegypti Malpighian tubules

Ming-Jiun Yu and Klaus W. Beyenbach

Department of Biomedical Sciences, Cornell University, Ithaca, New York 14853


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The role of Ca2+ in mediating the diuretic effects of leucokinin-VIII was studied in isolated perfused Malpighian tubules of the yellow fever mosquito, Aedes aegypti. Peritubular leucokinin-VIII (1 µM) decreased the transepithelial resistance from 11.2 to 2.6 kOmega · cm, lowered the transepithelial voltage from 42.8 to 2.7 mV, and increased transepithelial Cl- diffusion potentials 5.1-fold. In principal cells of the tubules, leucokinin-VIII decreased the fractional resistance of the basolateral membrane from 0.733 to 0.518. These effects were reversed by the peritubular Ca2+-channel blocker nifedipine, suggesting a role of peritubular Ca2+ and basolateral Ca2+ channels in signal transduction. In Ca2+-free Ringer bath, the effects of leucokinin-VIII were partial and transient but were fully restored after the bath Ca2+ concentration was restored. Increasing intracellular Ca2+ with thapsigargin duplicated the effects of leucokinin-VIII, provided that peritubular Ca2+ was present. The kinetics of the effects of leucokinin-VIII is faster than that of thapsigargin, suggesting the activation of inositol-1,4,5-trisphosphate-receptor channels of intracellular stores. Store depletion may then bring about Ca2+ entry into principal cells via nifedipine-sensitive Ca2+ channels in the basolateral membrane.

nifedipine; thapsigargin


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

HORMONES AND NEUROPEPTIDES are known to regulate transepithelial electrolyte transport in Malpighian tubules of insects (12, 39). So far, five classes of diuretic hormones have been identified: serotonin (8, 10, 29), the corticotropin-releasing factor (CRF)-like diuretic peptides (27), the leucokinins (20), the cardioacceleratory peptides (17), and the calcitonin-like diuretic peptides (18).

The leucokinins have been named after Leucophaea, the cockroach from which these neuropeptides were first isolated, sequenced, and synthesized in the laboratory of Holman et al. (23). Holman et al. considered all eight leucokinins myotropic peptides because they stimulate contractions of the cockroach hindgut. Curious that neuropeptides stimulating excretory functions in the gut might also stimulate epithelial transport in Malpighian tubules upstream, we discovered that leucokinins increased the rate of fluid secretion in isolated Malpighian tubules of the yellow fever mosquito, Aedes aegypti (20). Since then, the diuretic effects of leucokinins have been observed in the house cricket (14), locust (37), tobacco hornworm (5), fruit fly (31), and housefly (26). As many as 30 leucokinins have now been isolated and sequenced in 4 orders and 9 species of insects (24, 38, 39), including 3 leucokinin-like peptides of the yellow fever mosquito (40).

Ca2+ is widely believed to mediate the diuretic effects of leucokinin in Malpighian tubules (12). However, the relative roles of extra- and intracellular Ca2+ in signal transduction have not been clearly defined. In the present study of the effects of leucokinin-VIII on Malpighian tubules of the yellow fever mosquito, we report that principal cells mediate the signaling pathway, which involves the release of Ca2+ from intracellular stores and, more importantly, the entry of Ca2+ into the cell via nifedipine-sensitive Ca2+ channels in the basolateral membrane.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Mosquitoes. The mosquito colony was maintained in a controlled environment at 26°C, 45% humidity, and a 14:10-h light-dark cycle. Mark Brown (Univ. of Georgia, Athens, GA) kindly provided eggs of the yellow fever mosquito, A. aegypti. After the eggs were submerged (~200) in dechlorinated tap water, they were exposed to a vacuum of ~610 Torr for 20 min to induce hatching. The hatched larvae were transferred to a flat pan (30 cm × 22 cm) containing 1 liter of dechlorinated tap water and fed "mosquito chow" every day. Mosquito chow contained equal volumes of lactalbumin (Sigma, St. Louis, MO), yeast hydrolysate (ICN Biochemicals, Cleveland, OH), and RMH-3200 chow (Agway, Ithaca, NY). The larvae started to pupate after 5-7 days. The pupae were transferred to 200 ml of dechlorinated tap water in a flask equipped with a netted bucket to capture adult mosquitoes emerging from the water (eclosion). Adult mosquitoes were offered 3% sucrose ad libitum. On the day of experiments, a female mosquito (3-7 days posteclosion) was cold anesthetized and decapitated. Malpighian tubules were then removed from the abdominal cavity under Ringer solution. Only tubule segments near the blind end of the tubule, between 0.2 and 0.3 mm long, were used for study. Experiments on blood-fed mosquitoes were not conducted.

Ringer solution, leucokinin-VIII, and chemicals. Ringer solution contained the following (in mM): 150 NaCl, 3.4 KCl, 1.8 NaHCO3, 1.7 CaCl2, 1.0 MgSO4, 25 HEPES, and 5.0 glucose. The pH was adjusted to 7.1 with 1 M NaOH. CaCl2 was omitted in Ca2+-free Ringer, and, in addition, 1.0 mM EGTA was added to buffer trace Ca2+ in some experiments. The free Ca2+ concentration in Ringer solution was calculated using an online program (WEBMAXC v2.10. http://www.stanford.edu/~cpatton/webmaxc2.htm).

Synthetic cockroach leucokinin-VIII was a kind gift from Ronald Nachman (US Dept. of Agriculture, College Station, TX). Although the sequences of mosquito leucokinins are known (40), we preferred to study the cockroach leucokinin for the following reasons. As shown in Table 1, the cockroach leucokinin-VIII used in the present study shares high structural similarity with the mosquito leucokinins, Aedes leucokinin 1 and 3, in the last 5 amino acids that are required for bioactivity in all 30 leucokinins (24, 38, 39). In addition, the cockroach and mosquito leucokinins share close functional similarities (20, 40). The EC50 of the cockroach leucokinin-VIII on transepithelial voltage (Vt) of the mosquito Malpighian tubule is 2 × 10-9 M (20), comparable to that of the mosquito leucokinins (40). Similarly, low concentrations of both cockroach and mosquito leucokinin cause only partial, oscillating depolarizations of Vt in mosquito Malpighian tubules (20, 40). Stable, sustained depolarizations of Vt require high concentrations (1 × 10-6 M) of cockroach or mosquito leucokinins (20, 40). Moreover, diuretic effects are not observed at low concentrations of leucokinin, regardless of origin; they are observed only at concentrations >1 × 10-8 M in the case of cockroach and mosquito leucokinins (20, 40). In the absence of major physiological or pharmacological differences between cockroach and mosquito leucokinins, we continued to study the effects of the cockroach peptide in view of our previous studies of this peptide in mosquito Malpighian tubules (20, 28, 33, 41, 43). To study the electrophysiological correlates of diuresis, we used leucokinin-VIII at a concentration of 1 × 10-6 M, as in previous studies (28, 33, 41, 43) and as required of the native leucokinin in the mosquito (40).

                              
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  		Table 1.   Primary sequences of leucokinin-VIII from the cockroach and Aedes leucokinins from the yellow fever mosquito

Thapsigargin and nifedipine were dissolved in dimethylsulfoxide (DMSO) and diluted in Ringer solution to the desired concentration. The final concentration of DMSO in Ringer solution was <0.1%, which had no measurable effects on Vt and transepithelial resistance (data not shown). Chemicals were purchased from Sigma, Fisher Scientific (Fair Lawn, NJ), and VWR Scientific (Willard, OH).

In vitro microperfusion of Malpighian tubules. Figure 1A illustrates the method for measuring transepithelial and membrane voltages and resistances in isolated perfused Malpighian tubules (7). The tubule lumen was cannulated with a double-barreled perfusion pipette with an outer diameter of ~10 µm (Theta-Borosilicate glass, no. 1402401; Hilgenberg, Germany). One barrel of this pipette was used to perfuse the tubule lumen with Ringer solution and to measure the Vt with respect to ground in the peritubular Ringer bath. The other barrel was used to inject current (I = 50 nA) into the tubule lumen for the measurement of the transepithelial resistance (Rt) by cable analysis (21). The peritubular bath (500 µl) was perfused with Ringer solution at a rate of 6 ml/min. Vt was recorded continuously, and Rt was measured periodically when of interest.


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  		Fig. 1.   A: electrophysiological measurements in Malpighian tubules of Aedes aegypti perfused in vitro by the method of Burg et al. (7). Transepithelial voltage (Vt) is measured through 1 barrel of the perfusion pipette lodged in the tubule lumen. The other barrel serves to inject current (I = 50 nA) into the tubule lumen for measurement of the transepithelial resistance (Rt) by cable analysis (21). The membrane voltage (Vbl) and the fractional resistance of the basolateral membrane are measured with a conventional microelectrode impaling a principal cell. Vl is the voltage measured in the collecting pipette. All measurements are referenced to ground in the peritubular bath. B: electrical equivalent circuit of transepithelial electrolyte secretion. V, voltage; R, resistance; E, electromotive force. Subscript letters: t, transepithelial; a, apical membrane; bl, basolateral membrane; sh, shunt.

To measure voltage and the fractional resistance of basolateral and apical membranes of principal cells, we impaled one principal cell of the perfused tubule with a conventional microelectrode (Fig. 1A). Referenced to ground in the peritubular Ringer bath, this microelectrode measured the basolateral membrane potential (Vbl). Because apical and basolateral membranes of principal cells are in series, the apical membrane voltage (Va) can be calculated as the difference between Vt and Vbl. Current injected into the tubule lumen gives rise to membrane voltage deflections across both apical and basolateral membranes (Delta Va and Delta Vbl) that are proportional to their respective resistances (Ra and Rbl). The fractional resistance of the basolateral membrane of the principal cell (fRbl) is therefore
 f<IT>R</IT><SUB>bl</SUB><IT>=R</IT><SUB>bl</SUB><IT>/</IT>(<IT>R</IT><SUB>a</SUB><IT>+R</IT><SUB>bl</SUB>)<IT>=&Dgr;V</IT><SUB>bl</SUB><IT>/</IT>(<IT>&Dgr;V</IT><SUB>a</SUB><IT>+&Dgr;V</IT><SUB>bl</SUB>)<IT>=&Dgr;V</IT><SUB>bl</SUB><IT>/&Dgr;V</IT><SUB>t,x</SUB> (1)
where Delta Vt,x is the change in Vt at the site where the microelectrode impales the principal cell "x" cm away from the opening of the current pipette. Delta Vt,x decreases along the length of the tubule according to Eq. 2
&Dgr;V<SUB>t,x</SUB><IT>=&Dgr;V</IT><SUB>t</SUB>{cosh[(<IT>l−x</IT>)<IT>/&lgr;</IT>]<IT>/</IT>cosh(<IT>l/&lgr;</IT>)} (2)
where l is the length of the perfused tubule segment, lambda  is the tubule length constant, and cosh is the hyperbolic function of cosine.

All voltage measurements were made with custom-made high-impedance amplifiers (Burr-Brown, 1011 Omega ). A permanent record of voltages was produced with a strip-chart recorder (model BD 64; Kipp and Zonen, Bohemia, NY). Data were also collected in a digital form using a Macintosh computer (model 7300) equipped with data-acquisition hardware (Multifunction I/O Board PCI-1200 and Termination Board model SC-2071) and software (LabView for Macintosh, v. 4.1; National Instruments Manufacturer, Austin, TX).

Transepithelial Cl- diffusion potentials. The amplitude of transepithelial Cl- diffusion potentials was measured as the change in Vt in response to the 10-fold replacement of Cl- with isethionate in the peritubular Ringer as in previous studies (43). The Cl- concentration in the tubule lumen was held constant by perfusion of the tubule lumen with normal Ringer at rates <5 nl/min. In the presence of a constant luminal Cl- concentration, the 10-fold step reduction of the peritubular Cl- concentration drove Cl- to diffuse from the tubule lumen to the peritubular bath, generating lumen-positive transepithelial potentials with magnitudes proportional to the shunt Cl- conductance.

Equivalent electrical circuit of transepithelial ion transport in Malpighian tubules. Because the secretion of NaCl and KCl by Malpighian tubules of A. aegypti generates voltages (2), transepithelial ion transport can be modeled with an electrical equivalent circuit, illustrated in Fig. 1B. The circuit distinguishes between the active transport pathway through principal cells and the passive transport pathway through a shunt (Rsh) located outside principal cells (Fig. 1B). The active transport pathway is further defined by electromotive forces (E) and resistance of the apical (a) and basolateral (bl) membranes. Vbl is the sum of the electromotive force of the basolateral membrane (Ebl) and the voltage drop (Ioc × Rbl) across the basolateral membrane resistance, where Ioc is the intraepithelial current during epithelial transport under "open-circuit" conditions. Likewise, the Va is the sum of Ea and Ioc × Ra.

Current passing through principal cells is carried largely by Na+ and K+ in a secretory direction from the peritubular bath to the tubule lumen. Current passing through the shunt is carried by Cl-, also in a secretory direction (Fig. 1B). Accordingly, the active transport pathway through principal cells is electrically coupled to the passive transport pathway of the shunt, such that an anion (Cl-) is secreted into the lumen for every cation (Na+ and K+) transported through principal cells. Indeed, rates of transepithelial cation secretion come close to, or equal, rates of transepithelial Cl- secretion in Aedes Malpighian tubules (33).

Statistical evaluation of data. Each tubule served as its own control. Accordingly, the data were analyzed for the differences between paired samples, control vs. experimental, with the use of the paired Student's t-test.


    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Leucokinin-VIII decreases the Vt and Rt in Malpighian tubules. Isolated Malpighian tubules generated lumen-positive Vt when the same Ca2+-containing Ringer solution was present on both sides of the epithelium (symmetrical perfusion). In the representative experiment shown in Fig. 2A, Vt was 70.7 mV (lumen positive), and the Rt was 18.0 kOmega · cm. After 1 µM leucokinin-VIII was added to the peritubular bath, Vt depolarized sharply to 5.0 mV together with the drop of Rt to 2.9 kOmega · cm. These effects were fully reversible on washout of leucokinin-VIII. In a total of seven Malpighian tubules, leucokinin-VIII significantly depolarized Vt from 40.6 ± 9.0 to 0.5 ± 1.3 mV and significantly decreased Rt from 9.9 ± 1.9 to 2.1 ± 0.2 kOmega · cm (Fig. 2B).


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  		Fig. 2.   Effects of leucokinin (LK)-VIII on the Vt and Rt in isolated perfused Malpighian tubules of the yellow fever mosquito, A. aegypti. The tubule was bathed and perfused with normal Ringer solution containing 1.7 mM Ca2+. A: reversible effects of LK-VIII in a representative Malpighian tubule. LK-VIII was washed into and removed from the peritubular Ringer as indicated by arrows. B: means ± SE of 7 tubule experiments.* P < 0.01.

Nifedipine reverses the effects of leucokinin-VIII. To explore the effects of leucokinin-VIII and nifedipine on principal cells of the tubule, we measured Vbl and fRbl with a conventional microelectrode impaling a principal cell (Fig. 1A). Data from 8-18 tubule experiments are summarized in Fig. 3. Under control conditions, Vt was 42.8 ± 5.3 mV, Vbl was -69.8 ± 4.3 mV, and Va was 109.5 ± 7.2 mV, lumen positive. The control Rt was 11.2 ± 1.7 kOmega · cm. The control fRbl, 0.733 ± 0.038, indicated that 73.3% of the transcellular resistance resided at the basolateral membrane and 26.7% at the apical membrane. In the absence of leucokinin-VIII, the 10-fold reduction of the peritubular Cl- concentration yielded a transepithelial Cl- diffusion potential (DPCl-) of 8.2 ± 1.2 mV, indicating a modest transepithelial Cl- conductance under control conditions.


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  		Fig. 3.   Effects of LK-VIII on electrophysiological variables of isolated perfused Malpighian tubules of A. aegypti and the reversal of these effects by nifedipine. Control data were obtained in the presence of normal Ringer (1.7 mM Ca2+) on both sides of the epithelium (5-10 min). Peritubular bath flow was then switched to include LK-VIII (1 µM) for 5 min and then LK-VIII plus nifedipine (50 µM) for 20 min. Steady-state values are shown for each treatment period. DP<SUB>Cl<SUP>−</SUP></SUB>, transepithelial Cl- diffusion potential in response to a 10-fold reduction of the peritubular Cl- concentration; Vbl, basolateral membrane voltage of the principal cell; fRbl, fractional resistance of the basolateral membrane. Values are means ± SE (no. of tubules). a Statistical significance (P < 0.05) is referenced to the previous condition. b Statistical significance (P < 0.05) is referenced to the control condition.

The addition of leucokinin-VIII to the peritubular bath profoundly and significantly affected all measured variables (Fig. 3). Vt dropped to 2.7 ± 0.6 mV within a few seconds. In parallel, Vbl hyperpolarized to -86.9 ± 5.2 mV, and Va depolarized to 89.4 ± 5.0 mV. Inspection of the equivalent circuit of Fig. 1B shows that the hyperpolarization of Vbl concomitant with the depolarization of Va is expected from a decrease in the Rsh. Consistent with a decrease in Rsh in the presence of leucokinin-VIII was the large drop of Rt from 11.2 to 2.6 ± 0.3 kOmega · cm together with the significant 5.1-fold increase in DP<SUB>Cl<SUP>−</SUP></SUB> to 42.1 ± 5.4 mV. Leucokinin-VIII also affected the transcellular transport pathway through principal cells as indicated by the decrease of fRbl from 0.733 to 0.518 ± 0.020.

Nifedipine (50 µM) reversed the effects of leucokinin-VIII. All variables returned toward control values, some completely and others incompletely (Fig. 3). Although the return of Vt from 2.7 to 22.5 ± 3.7 mV is highly significant (P < 0.00005), Vt did not fully return to the control mean value of 42.8 mV. Likewise, DPCl- significantly (P < 0.00005) decreased from 42.1 to 16.4 ± 2.9 mV but did not reach control diffusion potentials of 8.2 mV. All other variables fully returned to control values. In particular, nifedipine completely reversed the effects of leucokinin-VIII on 1) Rt, which returned to 10.5 ± 1.5 kOmega · cm; 2) Vbl, which repolarized to -74.3 ± 5.1 mV; 3) Va, which repolarized to 96.2 ± 6.0 mV; and 4) fRbl, which returned to 0.709 ± 0.039 (Fig. 3). It took between 5 and 12 min for nifedipine to reverse the effects of leucokinin-VIII on both principal cells and transepithelial shunt conductance.

In the absence of leucokinin, i.e., under control conditions, nifedipine had no significant effects on Vt (control, 51.6 ± 6.0 mV; nifedipine, 53.2 ± 7.6 mV) and Rt (control, 12.1 ± 2.1 kOmega · cm; nifedipine, 9.9 ± 1.1 kOmega · cm) in seven isolated perfused Malpighian tubules.

Effects of leucokinin-VIII are independent of barium-sensitive K+ channels. The addition of Ba2+ (5 mM) to the peritubular Ringer solution had immediate and significant effects on the electrophysiology of the tubule and its principal cells (Fig. 4). Vt depolarized from 19.0 ± 4.2 to 17.1 ± 4.0 mV, Vbl hyperpolarized from -64.6 ± 6.5 to -71.0 ± 6.9 mV, Va hyperpolarized from 83.7 ± 7.6 to 92.3 ± 7.7 mV, and fRbl increased from 0.710 ± 0.030 to 0.800 ± 0.024, consistent with the block of basolateral membrane K+ channels observed previously in principal cells (28). The Ba2+ blockade of K+ channels did not significantly increase Rt, although a trend to higher values was observed (Fig. 4).


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  		Fig. 4.   Effects of LK-VIII in the presence of K+ channel blocker Ba2+. Experiments began with the perfusion of Malpighian tubules of A. aegypti with normal Ringer solution (1.7 mM Ca2+) present in the tubule lumen and the peritubular bath (control). After a control period of 5-10 min, the peritubular Ringer bath was changed to include 5 mM Ba2+ for 5 min. Thereafter, the bath was supplemented with LK-VIII in the presence of Ba2+ (1 µM) for 5 min and then with nifedipine (50 µM) in the presence of LK-VIII and Ba2+ for 20 min. Steady-state values are summarized for each treatment period. Values are means ± SE (no. of tubules). a Statistical significance (P < 0.05) is referenced to the previous condition. b Statistical significance (P < 0.05) is referenced to the preLK-VIII with Ba2+ condition.

Ba2+ did not impair the efficacy of leucokinin-VIII (Fig. 4). On the addition of leucokinin-VIII to the peritubular bath containing 5 mM Ba2+, Vt dropped immediately and significantly from 17.1 ± 4.0 to 0.9 ± 0.7 mV together with the drop in Rt from 6.7 ± 0.9 to 1.8 ± 0.2 kOmega · cm. Furthermore, Vbl significantly hyperpolarized from -71.0 ± 6.9 to -87.8 ± 6.7 mV, and fRbl significantly decreased from 0.800 ± 0.024 to 0.542 ± 0.030. The small depolarization of Va was not statistically significant.

As in the absence of Ba2+ (Fig. 3), nifedipine (50 µM) significantly reversed all effects of leucokinin-VIII in the presence of Ba2+ (Fig. 4). Vt significantly repolarized to 9.1 ± 2.0 mV, Rt significantly returned to 4.7 ± 0.4 kOmega · cm, Vbl significantly repolarized to -78.4 ± 6.5 mV, and fRbl significantly returned to 0.724 ± 0.026. It took 3-9 min for nifedipine to reverse the effects of leucokinin-VIII in the presence of Ba2+.

Effects of leucokinin-VIII depend on the presence of extracellular Ca2+. Because the peritubular addition of the Ca2+ channel blocker nifedipine reversed the effects of leucokinin-VIII (Figs. 3 and 4), we explored the role of extracellular Ca2+ in signal transduction by testing the effects of leucokinin-VIII in peritubular Ringer solution made Ca2+ free by deleting this ion and buffering trace Ca2+ with EGTA. As shown in Fig. 5 for a representative tubule experiment, the change from normal peritubular Ringer to Ca2+-free Ringer had no obvious effects on Vt (70.0 mV) and Rt (17.9 kOmega · cm). However, the effects of leucokinin-VIII were markedly diminished in the absence of peritubular Ca2+. On the addition of leucokinin-VIII to the Ca2+-free peritubular Ringer bath, Vt depolarized from 68.9 to 11.2 mV together with the drop of Rt from 18.1 to 4.6 kOmega · cm. However, these effects were only transient. Vt and Rt returned to values approximately one-half of control (44.3 and 61.4%, respectively) and then began to exhibit small oscillations. Characteristic of these oscillations was the parallel change of Vt and Rt. As Rt dropped, Vt depolarized toward zero, and as Rt rose, Vt repolarized (Fig. 5).


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  		Fig. 5.   Dependence of the effects of LK-VIII on peritubular Ca2+ concentrations. The representative tubule experiment began in the presence of normal Ringer containing 1.7 mM Ca2+ on both sides of the epithelium. The peritubular bathing solution was then changed to Ca2+-free Ringer (no Ca2+ plus 1 mM EGTA). LK-VIII (prepared in Ca2+-free Ringer) was subsequently added to the Ca2+-free peritubular bath. Thereafter, CaCl2 was added to the peritubular bath to yield the free Ca2+ concentrations indicated. Dashed arrows point to the Vt when Rt was measured.

The addition of Ca2+ to the peritubular bath to yield a free Ca2+ concentration of 31 nM only transiently reduced Vt to 13.0 mV and Rt to 5.6 kOmega · cm before a return to oscillations again. A similar response was observed on increasing the peritubular free Ca2+ concentration further to 17 µM. But at this Ca2+ concentration, the initial parallel effect on Vt and Rt was stronger, and it lasted longer before returning to oscillations that now were more pronounced than those at lower Ca2+ concentrations. Only after the Ca2+ buffering capacity of peritubular EGTA was exceeded, i.e., in the presence of 1 mM Ca2+, were the full effects of leucokinin-VIII observed: permanent, steady-state low values of Vt (3.6 mV) and Rt (4.0 kOmega · cm). What is shown for the single tubule experiment in Fig. 5 was observed in 10 other Malpighian tubules (data not shown).

Thapsigargin duplicates the effects of leucokinin-VIII, and nifedipine reverses them. The oscillations of Vt and Rt in leucokinin-VIII-treated tubules bathed in Ca2+-free Ringer (Fig. 5) suggest a role of intracellular Ca2+ in signal transduction. For this reason, the effects of thapsigargin, a specific inhibitor of Ca2+ uptake by the intracellular stores, were of interest. The addition of 1 µM thapsigargin to the normal peritubular Ringer solution containing Ca2+ duplicated the effects of leucokinin-VIII (Fig. 6). Vt depolarized from 21.4 ± 4.4 to 3.0 ± 0.3 mV, and Rt decreased from 7.9 ± 1.4 to 2.6 ± 0.2 kOmega · cm. At the same time, Vbl hyperpolarized from -70.0 ± 5.7 to -80.6 ± 7.4 mV, Va depolarized from 91.2 ± 9.9 to 82.0 ± 8.1 mV, and fRbl decreased from 0.628 ± 0.056 to 0.486 ± 0.056. All of these effects were statistically significant (Fig. 6).


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  		Fig. 6.   Effects of thapsigargin on electrophysiological variables of isolated perfused Malpighian tubules of A. aegypti and reversal by nifedipine. Tubules were perfused in vitro with normal Ca2+-containing Ringer solution in the tubule lumen and the peritubular bath (control; 5-10 min). The peritubular bath was then switched to include thapsigargin (1 µM) for 5-10 min and then to include thapsigargin plus nifedipine (50 µM) for another 20 min. Only steady-state values are summarized for each treatment period. Values are means ± SE (no. of tubules). a Statistical significance (P < 0.05) is referenced to the previous condition. b Statistical significance (P < 0.05) is referenced to the control condition.

The kinetics of the thapsigargin effects was significantly slower than that of leucokinin-VIII. The effects of leucokinin-VIII on electrophysiological variables took only seconds (Figs. 2-5). In contrast, the effects of thapsigargin started with a delay of 2 min and developed slowly, taking 5-7 min with a hyperbolic time course to reach effects comparable with those of leucokinin-VIII.

Nifedipine (50 µM) reversed the effects of thapsigargin (Fig. 6) as it reversed the effects of leucokinin-VIII (Figs. 3 and 4). Vt significantly repolarized to 15.1 ± 4.0 mV but remained significantly different from control. All other variables fully returned to control values in the presence of nifedipine. Rt returned to 7.8 ± 1.5 kOmega · cm, Vbl repolarized to -71.5 ± 5.5 mV, Va repolarized to 87.2 ± 8.7 mV, and fRbl returned to 0.632 ± 0.056 (Fig. 6). It took between 4 and 10 min for nifedipine to reverse the effects of thapsigargin.

Effects of thapsigargin depend on extracellular Ca2+. To gain insights into the relative roles of intra- and extracellular Ca2+ in the signal transduction of leucokinin-VIII, we explored the effects of thapsigargin in the absence and presence of extracellular Ca2+. As shown in Fig. 7, thapsigargin (1 µM) had no significant effect on any of the six electrophysiological variables in the absence of peritubular Ca2+ (-Ca2+), not even after 10 min. Moreover, the oscillations of Vt and Rt that were observed with leucokinin-VIII in Ca2+-free Ringer (Fig. 5) were not observed with thapsigargin in Ca2+-free Ringer. However, after exposure to thapsigargin for 10 min, restoration of the Ca2+ concentration in the peritubular Ringer to 1.7 mM immediately restored the full effects of thapsigargin with significant effects on all variables: Vt decreased from 32.1 ± 4.4 to 3.2 ± 0.8 mV, Rt decreased from 8.2 ± 1.2 to 2.4 ± 0.4 kOmega · cm, and DP<SUB>Cl<SUP>−</SUP></SUB> increased from 19.2 ± 3.5 to 38.2 ± 3.5 mV (Fig. 7). In parallel with these changes, Vbl hyperpolarized from -67.9 ± 6.0 to -86.6 ± 3.1 mV, Va depolarized from 96.6 ± 5.5 to 89.7 ± 3.6 mV, and fRbl decreased from 0.733 ± 0.043 to 0.451 ± 0.071. Significantly, the effects of thapsigargin were reversed again when the peritubular Ringer solution was switched back to nominally Ca2+-free Ringer. Vt returned to 25.8 ± 4.7 mV, Rt returned to 7.4 ± 1.3 kOmega · cm, and DPCl- decreased to 20.4 ± 3.7 mV. Similarly, Vbl depolarized to -70.9 ± 5.1 mV, Va repolarized to 94.7 ± 5.6 mV, and fRbl increased to 0.740 ± 0.040. It took between 2 and 6 min for nominally Ca2+-free Ringer solution to reverse the effects of thapsigargin. Restoring the normal Ca2+ concentration in the peritubular bath for a second time elicited the full effects of thapsigargin again (data not shown).


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  		Fig. 7.   Dependence of the effects of thapsigargin on peritubular Ca2+. Experiments began with the perfusion of Malpighian tubules of A. aegypti with normal Ringer solution containing 1.7 mM Ca2+ in the tubule lumen and the peritubular bath (+Ca2+). After ~5 min, the peritubular bath was changed to Ca2+-free Ringer (-Ca2+) containing 1 µM thapsigargin for 10 min. Then the bath was changed to include 1.7 mM Ca2+ in the presence of thapsigargin (+Ca2+) for 5 min. Finally, the bath was returned to Ca2+-free Ringer (-Ca2+) containing 1 µM thapsigargin for 10 min. Only steady-state values are summarized for each treatment period. Values are means ± SE (no. of tubules). a Statistical significance (P < 0.05) is referenced to the previous condition. b Statistical significance (P < 0.05) is referenced to the control condition.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Leucokinins and electrophysiological and diuretic effects in Malpighian tubules. The leucokinins were first isolated by Holman et al. (23) from the heads of cockroaches (Leucophaea) and named "cephalo-myotropic peptides" for their anatomical origin and their ability to stimulate contractions in the cockroach hindgut. Diuretic properties of the leucokinins were first found in Malpighian tubules of the yellow fever mosquito, A. aegypti (20), and later also in the house cricket, locust, tobacco hornworm, fruit fly, and housefly (5, 14, 26, 31, 37). Because the secretion of electrolytes generates Vt in Malpighian tubules, measures of voltage and resistance have been useful in elucidating transport mechanisms and their regulation (2) and serve as a rapid bioassay in the search and isolation of new diuretic peptides and hormones (19). Although changes in voltage can reflect changes in ion transport, they must not necessarily do so. For example, Aedes leucokinin 2 depolarizes the Vt in the isolated Malpighian tubule, but it fails to trigger diuresis even at micromolar concentrations (40). The dissociation of electrophysiological effects from diuretic effects can also be a matter of dose. Aedes leucokinin 3 affects the Vt at concentrations as low as 10-10 M, but its diuretic effects begin at a concentration around 10-8 M and approach maximum at 10-6 M (40). Furthermore, at concentrations from 10-10 to 10-7 M, Aedes leucokinin 3 elicits only partial oscillations of the Vt, and low, steady-state depolarizations of the Vt are only observed at the diuretic concentration of 10-6 M (40). The situation is similar for the cockroach leucokinin-VIII used in the present study. Like the Aedes leucokinins, the cockroach leucokinin-VIII elicits only electrical effects (voltage depolarizations) at low concentrations; concentrations of 10-6 M and higher are needed to observe 1) low, steady-state depolarizations of the Vt and 2) diuretic effects (20). These functional similarities are not surprising in view of close structural similarities of the cockroach and mosquito leucokinins in the crucial COOH-terminal pentapeptide sequence necessary for bioactivity (Table 1). Kinin-dependent and dose-dependent effects on Vt and fluid secretion suggest multiple receptors and/or signaling pathways in mediating the actions of these peptides.

In Malpighian tubules of the fruit fly and the yellow fever mosquito, leucokinin increases the Cl- conductance of a transepithelial shunt located outside principal cells (32, 33). Two sites of the Cl- shunt have been proposed, stellate cells in Malpighian tubules of the fruit fly (32) and the paracellular pathway in Malpighian tubules of the yellow fever mosquito (33). In the present study, 1 µM leucokinin-VIII had the following effects: it decreased the Rt 4.3-fold, it short-circuited the Vt to values close to zero, and it increased transepithelial Cl- diffusion potential >5-fold (Figs. 2-5). These observations confirm the increase in a paracellular shunt Cl- conductance, with the effects of producing a "leaky epithelium" suitable for high transport rates under conditions of diuresis. The increase in the paracellular conductance is expected to hyperpolarize the Vbl of principal cells and to depolarize the Va that were observed (Figs. 1B, 3, and 4). Consistent with the effect on the paracellular pathway is the increase in the transepithelial permeability to two markers of the paracellular pathway, inulin and mannitol, induced by leucokinin in Malpighian tubules of Locusta and Aedes (12, 41).

Leucokinin and its Ca2+-dependent signal pathway. Wherever the signal transduction pathway of leucokinin has been studied, Ca2+ has been found to serve as second messenger (12). Ca2+ ionophores that bring Ca2+ into the cell from extracellular fluids duplicate the effects of leucokinin by stimulating fluid secretion in Malpighian tubules of the locust (30), house cricket (15), yellow fever mosquito (11), and black field cricket (42). Thapsigargin, which prevents Ca2+ uptake by intracellular stores, thereby increasing intracellular Ca2+ concentrations, increases fluid secretion in Malpighian tubules of the locust (13), fruit fly (36), housefly (26), house cricket (12), and black field cricket (42). Clamping intracellular Ca2+ concentrations at low levels with the intracellular Ca2+ chelator 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA)-AM abolishes the diuretic effects of leucokinin in Malpighian tubules of the house cricket (15) and the fruit fly (31). Actual measurements of intracellular Ca2+ concentrations show that leucokinin increases Ca2+ concentrations in stellate cells of the fruit fly Malpighian tubules (32) but increases Ca2+ concentration in principal cells of Malpighian tubules of the house cricket (12).

The ability of Ca2+ ionophore and thapsigargin to mimic the effects of leucokinin suggests roles of both extracellular and intracellular Ca2+ in signal transduction. The present study sought to clarify the relative roles of the two Ca2+ sources, extracellular fluid and intracellular stores, in mediating the diuretic effects of leucokinin in Malpighian tubules of the yellow fever mosquito, A. aegypti.

Previous studies in our laboratory have shown that Ca2+ ionophore A-23187 duplicated the effects of leucokinin-VIII in Aedes Malpighian tubules (11). The present study shows duplication of the effects of leucokinin-VIII by thapsigargin, an agent known to increase intracellular Ca2+ concentrations (Figs. 6 and 7). Like the effects of leucokinin-VIII, the effects of thapsigargin were critically dependent on peritubular Ca2+ as shown by the reversal of both leucokinin-VIII and thapsigargin effects on removal of Ca2+ from the peritubular Ringer bath or on the addition of the Ca2+ channel blocker nifedipine (Figs. 3-7). Significantly, both leucokinin-VIII and thapsigargin lowered the fRbl of principal cells, an effect that reversed toward control values by removing Ca2+ from or adding nifedipine to the peritubular Ringer solution (Figs. 3-7). These observations suggest the activation of basolateral membrane Ca2+ channels that are part of the signal transduction pathway of leucokinin. Apparently, a large number of Ca2+ channels must be activated in view of the substantial decrease in fRbl on the addition of leucokinin-VIII or thapsigargin. Primary or secondary effects of leucokinin on basolateral membrane K+ channels were ruled out in the present study, although K+ channels account for 64% of the basolateral conductance in principal cells of Aedes Malpighian tubules (4). After the blocking of K+ channels with the supramaximal dose of 5 mM Ba2+, the effects of leucokinin-VIII were not blocked (Fig. 4). Again, fRbl, Rt, Vt, and Vbl significantly decreased. Moreover, in the presence of Ba2+, the Ca2+ channel blocker nifedipine reversed these effects of leucokinin-VIII. Accordingly, leucokinin activates a substantial Ca2+ conductance in the basolateral membrane of principal cells.

Roles of intra- and extracellular Ca2+. The present study revealed that intracellular Ca2+ initiates and peritubular Ca2+ sustains the effects of leucokinin in Aedes Malpighian tubules. Thapsigargin duplicated the effects of leucokinin only if millimolar Ca2+ concentrations were present in the peritubular Ringer bath (Figs. 6 and 7). In the absence of extracellular Ca2+, thapsigargin had no effects on tubule and principal cell electrophysiology (Fig. 7). Thus the increase in cytoplasmic Ca2+ from intracellular stores may be sufficient to initiate the effects of leucokinin, but peritubular Ca2+ is needed to maintain the effects.

Inositol-1,4,5-trisphosphate (IP3) takes part in the signaling cascade, as shown in the laboratory of Hagedorn (9), where all three leucokinins of A. aegypti were found to increase IP3 concentration in isolated Malpighian tubules. Because IP3 is known to release Ca2+ from intracellular stores via IP3 receptor Ca2+ channels (1), the depletion of intracellular Ca2+ stores could trigger the activation of so-called store-operated Ca2+ channels in the basolateral membrane of principal cells (35). The hypothesis of store-operated Ca2+ channels in the basolateral membrane is supported by two observations in the present study. First, under conditions of intracellular Ca2+ depletion (Figs. 5 and 7), the addition of Ca2+ to the peritubular medium of leucokinin- or thapsigargin-treated tubules immediately restored their full effects, as if basolateral membrane Ca2+ channels were already open in these Ca2+-depleted cells (Fig. 7). Second, responses mediated via sudden Ca2+ store depletion by leucokinin-VIII are expected to be much faster than those mediated via slow Ca2+ store depletion by thapsigargin. When IP3 receptor Ca2+ channels of intracellular Ca2+ stores are activated by leucokinin-VIII, the electrophysiological responses to the tubule are immediate, within seconds (Figs. 2 and 5). In contrast, when the depletion of intracellular Ca2+ stores is achieved by inhibition of the Ca2+ pump with thapsigargin, electrophysiological responses of the tubule develop slowly over 10 min, reflecting the slow Ca2+ leak from intracellular stores.

In the absence of peritubular Ca2+, thapsigargin had no effect whatsoever on tubule electrophysiology, but leucokinin-VIII had partial and transient effects, consistent with the sudden release of Ca2+ from intracellular stores but in quantities insufficient to sustain the full and lasting effects of leucokinin (Figs. 5 and 7). The transients began as large parallel oscillations of the Vt and Rt with brief transitions to the leaky epithelium. These transitions diminished with time, returning the tubule to control values of the moderately "tight epithelium." The transitions had a frequency not uncommon for cyclical changes in intracellular Ca2+ concentrations, suggesting temporal displacements of Ca2+ release and reuptake by intracellular Ca2+ stores.

The oscillations of the Vt and Rt observed in leucokinin-treated tubules in the absence of peritubular Ca2+ (Fig. 5) were similar to spontaneous transients seen frequently in Aedes Malpighian tubules in the presence of peritubular Ca2+ (3). Similar voltage transients are observed in Aedes Malpighian tubules in the absence and presence of low Aedes leucokinin concentrations (40) and in Drosophila Malpighian tubules in the presence and absence of peritubular Ca2+ (6). BAPTA-AM, a chelator of cytosolic Ca2+, eliminates the voltage transients in Drosophila Malpighian tubules (6). Thus it appears that spontaneous cyclical changes in cytosolic Ca2+ concentrations are responsible to oscillating Vt and Rt, even in the absence of peritubular Ca2+ (Fig. 5).

Observations made in the present study allow a hypothetical model for the signal transduction pathway activated by high micromolar concentrations of leucokinin (Fig. 8). We propose a leucokinin receptor at the basolateral membrane on the basis of the recent identification of a leucokinin-binding protein in Aedes Malpighian tubules (33, 34). The receptor is probably heterotrimeric G protein coupled in view of 1) predictions from the primary sequences of the known leucokinin receptors from a pond snail (16) and a cattle tick (25) and 2) duplication of the electrophysiological effects of leucokinin in Aedes Malpighian tubules by aluminum tetrafluoride (AlF<UP><SUB>4</SUB><SUP>−</SUP></UP>), a known activator of G proteins (43). The stimulation of phospholipase C-beta to produce diacylglycerol and IP3 is indicated by the rise in intracellular IP3 concentrations measured in leucokinin-stimulated Aedes Malpighian tubules (9). IP3 is then thought to trigger the sudden release of Ca2+ from intracellular Ca2+ stores via IP3 receptor Ca2+ channels (1). The depletion of intracellular Ca2+ stores may then activate nifedipine-sensitive Ca2+ channels in the basolateral membrane of principal cells. Extracellular Ca2+ entering the cells then goes on to produce and hold the epithelium in the "leaky" condition as long as leucokinin is present. Exactly how Ca2+ entry leads to the increase in the shunt Cl- conductance and the maintenance of diuretic transepithelial transport rates is an intriguing question this study leaves unanswered.


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  		Fig. 8.   Hypothetical model of the LK signal transduction pathway in principal cells of Malpighian tubules of the yellow fever mosquito, A. aegypti. LK is known to increase rates of the transepithelial secretion of both NaCl and KCl, consistent with the increase in transepithelial Cl- permeability (33). Water follows by osmosis. Electrophysiological studies indicate the increase of the Cl- conductance of the paracellular, septate-junctional pathway between principal and/or stellate cells (33). LK-R, LK receptor; G, heterotrimeric G protein; AlF<UP><SUB>4</SUB><SUP>−</SUP></UP>, aluminum tetrafluoride; PLC, phospholipase C-beta ; PIP2, phosphatidylinositol-4,5-bisphosphate; IP3, inositol-1,4,5-trisphosphate; DAG, diacylglycerol; IP3-R, IP3 receptor; TG, thapsigargin; NP, nifedipine; A-23187, Ca2+/Mg2+ ionophore; +, activation; -, inhibition.


    ACKNOWLEDGEMENTS

We thank the National Science Foundation for supporting our studies.


    FOOTNOTES

We thank Dr. Ronald Nachman for the generous gift of synthetic leucokinin-VIII, Dr. Mark Brown for his kind gift of A. aegypti eggs, and Daniel S. Wu for fruitful discussions.

Address for reprint requests and other correspondence: K. W. Beyenbach, Dept. of Biomedical Sciences, VRT 8014, Cornell Univ., Ithaca, NY 14853 (E-mail: kwb1{at}cornell.edu).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

April 10, 2002;10.1152/ajprenal.00041.2002

Received 30 January 2002; accepted in final form 1 April 2002.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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M.-J. Yu and K. W. Beyenbach
Effects of leucokinin-VIII on Aedes Malpighian tubule segments lacking stellate cells
J. Exp. Biol., February 1, 2004; 207(3): 519 - 526.
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00041.2002v1
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