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
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ABSTRACT |
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 k
· 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
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INTRODUCTION |
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.
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MATERIALS AND METHODS |
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
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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.
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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
(
Va and
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
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(1)
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where
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.
Vt,x decreases along the
length of the tubule according to Eq. 2
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(2)
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where l is the length of the perfused tubule segment,
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
). 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.
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RESULTS |
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 k
· 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 k
· 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 k
· 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.
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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 k
· 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.
, 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.
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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 k
· cm together with the significant 5.1-fold increase in
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 k
· 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 k
· cm;
nifedipine, 9.9 ± 1.1 k
· 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.
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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 k
· 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 k
· 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 k
· 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 k
· 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.
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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 k
· 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 k
· 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 k
· 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.
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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 k
· 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 k
· cm,
and
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 k
· 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 |
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
),
a known activator of G proteins (43). The stimulation of
phospholipase C-
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 ,
aluminum tetrafluoride; PLC, phospholipase C- ; 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.
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|
 |
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 |
1.
Berridge, MJ.
Inositol trisphosphate and calcium signalling.
Nature
361:
315-325,
1993[ISI][Medline].
2.
Beyenbach, KW.
Energizing epithelial transport with the vacuolar H+-ATPase.
News Physiol Sci
16:
145-151,
2001[Abstract/Free Full Text].
3.
Beyenbach, KW,
Aneshansley JD,
Pannabecker TL,
Masia R,
Gray D,
and
Yu MJ.
Oscillations of voltage and resistance in Malpighian tubules of Aedes aegypti.
J Insect Physiol
46:
321-333,
2000[ISI][Medline].
4.
Beyenbach, KW,
and
Masia R.
Membrane conductances of principal cells in Malpighian tubules of Aedes aegypti.
J Insect Physiol.
48:
375-386,
2002[ISI][Medline].
5.
Blackburn, MB,
Wagner RM,
Shabanowitz J,
Kochansky JP,
Hunt DF,
and
Raina AK.
The isolation and identification of three diuretic kinins from the abdominal ventral nerve cord of adult Helicoverpa zea.
J Insect Physiol
41:
723-730,
1995[ISI].
6.
Blumenthal, EM.
Characterization of transepithelial potential oscillations in the Drosophila Malpighian tubule.
J Exp Biol
204:
3075-3084,
2001[Abstract/Free Full Text].
7.
Burg, M,
Grantham J,
Abramow M,
and
Orloff J.
Preparation and study of fragments of single rabbit nephrons.
Am J Physiol
210:
1293-1298,
1966[Free Full Text].
8.
Cady, C,
and
Hagedorn HH.
The effect of putative diuretic factors on in vivo urine production in the mosquito, Aedes aegypti.
J Insect Physiol
45:
317-325,
1999[ISI][Medline].
9.
Cady, C,
and
Hagedorn HH.
Effects of putative diuretic factors on intracellular second messenger levels in the Malpighian tubules of Aedes aegypti.
J Insect Physiol
45:
327-337,
1999[ISI][Medline].
10.
Clark, TM,
and
Bradley TJ.
Additive effects of 5-HT and diuretic peptide on Aedes Malpighian tubule fluid secretion.
Comp Biochem Physiol A Mol Integr Physiol
119:
599-605,
1998[ISI].
11.
Clark, TM,
Hayes TK,
Holman GM,
and
Beyenbach KW.
The concentration dependence of CRF-like diuretic peptide: mechanisms of action.
J Exp Biol
201:
1753-1762,
1998[Abstract/Free Full Text].
12.
Coast, GM.
Insect diuretic peptides: structures, evolution, and actions.
Am Zool
38:
442-449,
1998[ISI].
13.
Coast, GM.
Synergism between diuretic peptides controlling ion and fluid transport in insect Malpighian tubules.
Regul Pept
57:
283-296,
1995[ISI][Medline].
14.
Coast, GM,
Holman GM,
and
Nachman RJ.
The diuretic activity of a series of cephalomyotropic neuropeptides, the achetakinins, on isolated Malpighian tubules of the house cricket, Acheta domesticus.
J Insect Physiol
36:
481-488,
1990[ISI].
15.
Coast, GM,
Kay I,
and
Wheeler CH.
Diuretic peptide in the house cricket, Acheta domesticus (L): a possible dual control of Malpighian tubules.
In: Molecular Comparative Physiology. Structure and Function of Primary Messengers in Invertebrates: Insect Diuretic and Antidiuretic Peptides, edited by Beyenbach KW.. Basel, Switzerland: Karger, 1993, p. 38-66.
16.
Cox, KJ,
Tensen CP,
Van der Schors RC,
Li KW,
van Heerikhuizen H,
Vreugdenhil E,
Geraerts WP,
and
Burke JF.
Cloning, characterization, and expression of a G-protein-coupled receptor from Lymnaea stagnalis and identification of a leucokinin-like peptide, PSFHSWSamide, as its endogenous ligand.
J Neurosci
17:
1197-1205,
1997[Abstract/Free Full Text].
17.
Davies, SA,
Huesmann GR,
Maddrell SH,
O'Donnell MJ,
Skaer NJ,
Dow JA,
and
Tublitz NJ.
CAP2b, a cardioacceleratory peptide, is present in Drosophila and stimulates tubule fluid secretion via cGMP.
Am J Physiol Regul Integr Comp Physiol
269:
R1321-R1326,
1995[Abstract/Free Full Text].
18.
Furuya, K,
Milchak RJ,
Schegg KM,
Zhang J,
Tobe SS,
Coast GM,
and
Schooley DA.
Cockroach diuretic hormones: characterization of a calcitonin-like peptide in insects.
Proc Natl Acad Sci USA
97:
6469-6474,
2000[Abstract/Free Full Text].
19.
Hayes, TK,
Holman GM,
Pannabecker TL,
Wright MS,
Strey AA,
Nachman RJ,
Hoel DF,
Olson JK,
and
Beyenbach KW.
Culekinin depolarizing peptide: a mosquito leucokinin-like peptide that influences insect Malpighian tubule ion transport.
Regul Pept
52:
235-248,
1994[ISI][Medline].
20.
Hayes, TK,
Pannabecker TL,
Hinckley DJ,
Holman GM,
Nachman RJ,
Petzel DH,
and
Beyenbach KW.
Leucokinins, a new family of ion transport stimulators and inhibitors in insect Malpighian tubules.
Life Sci
44:
1259-1266,
1989[ISI][Medline].
21.
Helman, SI.
Determination of electrical resistance of the isolated cortical collecting tubule and its possible anatomical location.
Yale J Biol Med
45:
339-345,
1972[ISI][Medline].
22.
Holman, GM,
Cook BJ,
and
Nachman RJ.
Isolation, primary structure, and synthesis of leucokinin VII and VIII: the final members of the new family of cephalomyotropic peptides isolated from head extracts of Leucophaea maderae.
Comp Biochem Physiol C Pharmacol Toxicol Endocrinol
88:
31-34,
1987[ISI].
23.
Holman, GM,
Cook BJ,
and
Nachman RJ.
Isolation, primary structure, and synthesis of two neuropeptides from Leucophaea Maderae members of a new family of cephalomyotropins.
Comp Biochem Physiol C Pharmacol Toxicol Endocrinol
84:
205-212,
1986[ISI].
24.
Holman, GM,
Nachman RJ,
and
Coast GM.
Isolation, characterization, and biological activity of a diuretic myokinin neuropeptide from the housefly, Musca domestica.
Peptides
20:
1-10,
1999[ISI][Medline].
25.
Holmes, SP,
He H,
Chen AC,
Ivie GW,
and
Pietrantonio PV.
Cloning and transcriptional expression of a leucokinin-like peptide receptor from the southern cattle tick, Boophilus microplus (Acari: Ixodidae).
Insect Mol Biol
9:
457-465,
2000[ISI][Medline].
26.
Iaboni, A,
Holman GM,
Nachman RJ,
Orchard I,
and
Coast GM.
Immunocytochemical localisation and biological activity of diuretic peptides in the housefly, Musca domestica.
Cell Tissue Res
294:
549-560,
1998[ISI][Medline].
27.
Kataoka, H,
Troetschler RG,
Li JP,
Kramer SJ,
Carney RL,
and
Schooley DA.
Isolation and identification of a diuretic hormone from the tobacco hornworm, Manduca sexta.
Proc Natl Acad Sci USA
86:
2976-2980,
1989[Abstract].
28.
Masia, R,
Aneshansley D,
Nagel W,
Nachman RJ,
and
Beyenbach KW.
Voltage clamping single cells in intact Malpighian tubules of mosquitoes.
Am J Physiol Renal Physiol
279:
F747-F754,
2000[Abstract/Free Full Text].
29.
Morgan, PJ,
and
Mordue W.
5-Hydroxytryptamine stimulates fluid secretion in locust Locusta migratoria Malpighian tubules independently of cyclic AMP.
Comp Biochem Physiol C Pharmacol Toxicol Endocrinol
79:
305-310,
1984[ISI].
30.
Morgan, PJ,
and
Mordue W.
The role of calcium in diuretic hormone action on locust Locusta migratoria Malpighian tubules.
Mol Cell Endocrinol
40:
221-232,
1985[ISI][Medline].
31.
O'Donnell, MJ,
Dow JA,
Huesmann GR,
Tublitz NJ,
and
Maddrell SH.
Separate control of anion and cation transport in Malpighian tubules of Drosophila melanogaster.
J Exp Biol
199:
1163-1175,
1996[Abstract/Free Full Text].
32.
O'Donnell, MJ,
Rheault MR,
Davies SA,
Rosay P,
Harvey BJ,
Maddrell SH,
Kaiser K,
and
Dow JA.
Hormonally controlled chloride movement across Drosophila tubules is via ion channels in stellate cells.
Am J Physiol Regul Integr Comp Physiol
274:
R1039-R1049,
1998[Abstract/Free Full Text].
33.
Pannabecker, TL,
Hayes TK,
and
Beyenbach KW.
Regulation of epithelial shunt conductance by the peptide leucokinin.
J Membr Biol
132:
63-76,
1993[ISI][Medline].
34.
Pietrantonio, PV,
Gibsona GE,
Strey AA,
Petzel D,
and
Hayes TK.
Characterization of a leucokinin binding protein in Aedes aegypti (Diptera: culicidae) Malpighian tubule.
Insect Biochem Mol Biol
30:
1147-1159,
2000[ISI][Medline].
35.
Putney, JW, Jr.,
and
McKay RR.
Capacitative calcium entry channels.
Bioessays
21:
38-46,
1999[ISI][Medline].
36.
Rosay, P,
Davies SA,
Yu Y,
Sozen A,
Kaiser K,
and
Dow JA.
Cell-type specific calcium signalling in a Drosophila epithelium.
J Cell Sci
110:
1683-1692,
1997[Abstract/Free Full Text].
37.
Schoofs, L,
Holman GM,
Proost P,
Van Damme J,
Hayes TK,
and
de Loof A.
Locustakinin, a novel myotropic peptide from Locusta migratoria, isolation, primary structure, and synthesis.
Regul Pept
37:
49-57,
1992[ISI][Medline].
38.
Terhzaz, S,
O'Connell FC,
Pollock VP,
Kean L,
Davies SA,
Veenstra JA,
and
Dow JA.
Isolation and characterization of a leucokinin-like peptide of Drosophila melanogaster.
J Exp Biol
202:
3667-3676,
1999[Abstract/Free Full Text].
39.
Torfs, P,
Nieto J,
Veelaert D,
Boon D,
van de Water G,
Waelkens E,
Derua R,
Calderon J,
de Loof A,
and
Schoofs L.
The kinin peptide family in invertebrates.
Ann NY Acad Sci
897:
361-373,
1999[Abstract/Free Full Text].
40.
Veenstra, JA,
Pattillo JM,
and
Petzel DH.
A single cDNA encodes all three Aedes leucokinins, which stimulate both fluid secretion by the Malpighian tubules and hindgut contractions.
J Biol Chem
272:
10402-10407,
1997[Abstract/Free Full Text].
41.
Wang, S,
Rubenfeld A,
Hayes TK,
and
Beyenbach K.
Leucokinin increases paracellular permeability in insect Malpighian tubules.
J Exp Biol
199:
2537-2542,
1996[Abstract/Free Full Text].
42.
Xu, W,
and
Marshall AT.
Control of ion and fluid transport by putative second messengers in different segments of the Malpighian tubules of the black field cricket Teleogryllus oceanicus.
J Insect Physiol
46:
21-31,
2000[ISI][Medline].
43.
Yu, MJ,
and
Beyenbach KW.
Leucokinin and the modulation of the shunt pathway in Malpighian tubules.
J Insect Physiol
47:
263-276,
2001[ISI][Medline].
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